Frogs of the United States and Canada [2 ed.] 1421444917, 9781421444918

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Frogs of the United States and Canada

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Frogs SECOND EDITION

of the United States and Canada

C. Kenneth Dodd Jr.

Johns Hopkins University Press Baltimore

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© 2013, 2023 Johns Hopkins University Press  All rights reserved. Published 2023 Printed in China on acid-­free paper 987654321

Frontispiece: “Rain Response” by Audrey K. Owens Special discounts are available for bulk purchases of this book. For more information, please contact Special Sales at specialsales@jh​.­edu.

Johns Hopkins University Press 2715 North Charles Street Baltimore, Mary­land 21218 www​.­press​.­jhu​.­edu Library of Congress Cataloging-­in-­Publication Data Names: Dodd, C. Kenneth, author. Title: Frogs of the United States and Canada / C. Kenneth Dodd Jr. Description: Second edition. | Baltimore : Johns Hopkins University Press, 2023. | Includes bibliographical references and index. Identifiers: LCCN 2022011648 | ISBN 9781421444918 (hardcover ; acid-­free paper) | ISBN 9781421444925 (ebook) Subjects: LCSH: Frogs—­United States. | Frogs—­Canada. Classification: LCC QL668.E2 D57 2023 | DDC 597.8/90971—­dc23/ eng/20220420 LC rec­ord available at https://­lccn​.­loc​.­gov​/­2022011648 A cata­log rec­ord for this book is available from the British Library.

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 To the memory of Albert Hazen Wright and Anna Allen Wright, pioneers in the study of North American frogs, and my late colleagues Lauren E. Brown, Gary Fellers, Joseph C. Mitchell, and Raymond D. Semlitsch for their contributions to amphibian conservation

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In the Eyes of a Toad  In the eyes of a toad ­there is no vision of tomorrow, nor a shadow image of bygone years. ­There is merely and simply: ­Here; and, Now. Oh, ­there ­will, no doubt, be a golden fleck or two of memory shining ­there—­a sort of photo­graph ­album drifting about the periphery of the pupil like a ­silent movie reel . . . . . . ​flip-­flash; flip-­flash . . . disjointed moments when spring breeding and winter hibernation have been recorded. Perhaps ­there also ­will be a random cast to the windowed orb, streaks and shadows of camouflaging pattern, a permanent file of instinct-­plus-­genetics; subtle markings; unobtrusive identity; a natu­ral logic and nurturing reason why toads do whatsoever it is that toads do— without question, doubt, or hesitation. It is not the man-­trail changes that shine reflection, but rather, it is the unchanged traits and symbiotic habits of a timeless world that are recorded in the marvelous eyes of the toad. Marian Lovene Griffey

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Contents

Preface to the First Edition  ix Preface to the Second Edition  xi Introduction xiii Abbreviations xxxiii

SPECIES ACCOUNTS

Family Ascaphidae Ascaphus montanus  1 Ascaphus truei  7

Family Bufonidae Anaxyrus americanus  15 Anaxyrus baxteri  40 Anaxyrus boreas  44 Anaxyrus californicus  61 Anaxyrus canorus  65 Anaxyrus cognatus  72 Anaxyrus debilis  81 Anaxyrus exsul  85 Anaxyrus fowleri  89 Anaxyrus hemiophrys  105 Anaxyrus houstonensis  111 Anaxyrus microscaphus  117 Anaxyrus monfontanus  122 Anaxyrus nelsoni  124 Anaxyrus nevadensis  127 Anaxyrus punctatus  129 Anaxyrus quercicus  136 Anaxyrus retiformis  141 Anaxyrus speciosus  143 Anaxyrus terrestris  146 Anaxyrus williamsi  156 Anaxyrus woodhousii  158 Incilius alvarius  167 Incilius nebulifer  171 Rhinella marina  176

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Family Craugastoridae Craugastor augusti  182

Family Eleutherodactylidae Eleutherodactylus cystignathoides  186 Eleutherodactylus guttilatus  188 Eleutherodactylus marnockii  190

Family Hylidae Acris blanchardi  193 Acris crepitans  206 Acris gryllus  213 Dryophytes andersonii  221 Dryophytes arenicolor  225 Dryophytes avivoca  230 Dryophytes chrysoscelis  234 Dryophytes cinereus  246 Dryophytes femoralis  256 Dryophytes gratiosus  261 Dryophytes squirellus  267 Dryophytes versicolor  274 Dryophytes wrightorum  288 Hyliola cadaverina  291 Hyliola regilla  296 Pseudacris brachyphona and Pseudacris collinsorum  311 Pseudacris brimleyi  317 Pseudacris clarkii  319 Pseudacris crucifer  323 Pseudacris feriarum  338 Pseudacris fouquettei  346 Pseudacris illinoensis  351 Pseudacris kalmi  354 Pseudacris maculata  357 Pseudacris nigrita  370 Pseudacris ocularis  375

Pseudacris ornata  379 Pseudacris streckeri  383 Pseudacris triseriata  387 Smilisca baudinii  393 Smilisca fodiens  395

Family Leptodactylidae Leptodactylus fragilis  399

Family Microhylidae Gastrophryne carolinensis  402 Gastrophryne mazatlanensis  410 Gastrophryne olivacea  412 Hypopachus variolosus  418

Family Rhinophrynidae Rhinophrynus dorsalis  421

Family Ranidae Lithobates areolatus  424 Lithobates berlandieri  431 Lithobates blairi  437 Lithobates capito  444 Lithobates catesbeianus  450 Lithobates chiricahuensis  477 Lithobates clamitans  483 Lithobates fisheri  508 Lithobates grylio  512 Lithobates heckscheri  516 Lithobates kauffeldi  520 Lithobates okaloosae  523 Lithobates onca  527 Lithobates palustris  531 Lithobates pipiens  540 Lithobates septentrionalis  568 Lithobates sevosus  576

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viii  Contents

Lithobates sphenocephalus  580 Lithobates sylvaticus  596 Lithobates tarahumarae  631 Lithobates virgatipes  635 Lithobates yavapaiensis  641 Rana aurora  647 Rana boylii  656 Rana cascadae  664 Rana draytonii  670 Rana luteiventris  678 Rana muscosa  688 Rana pretiosa  693 Rana sierrae  701

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Family Scaphiopodidae Scaphiopus couchii  708 Scaphiopus holbrookii  714 Scaphiopus hurterii  725 Spea bombifrons  729 Spea hammondii  737 Spea intermontana  742 Spea multiplicata  748

Introduced Species Dendrobates auratus  759 Eleutherodactylus coqui  761

Eleutherodactylus planirostris  764 Glandirana rugosa  768 Litoria caerulea  771 Osteopilus septentrionalis  773 Xenopus laevis  779 Xenopus tropicalis  782 Glossary 785 Bibliography 789 Index of Scientific and Common Names 947 Index of Potential Stressors  953

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Preface to the First Edition

­ here are two reasons I became a biologist. First, nature T fascinates me. I have always been astonished at the diversity of life and how the sum total of its parts, from basic chemistry through physiology and ge­ne­tics all the way to im­mense ecosystems, still cannot explain the essence and “why” of life. Through herpetology, I have tried to make sense of how even a small portion of nature works, and I have never understood ­people who have no interest in what makes them and our world what they are. ­There is nothing more fascinating than the organ­ization and evolution of life. The second reason for becoming a biologist was the dread of working in an office building. I wanted to be outdoors, not anchored to a desk; if ­there ­were canyons and forests and wild animals “out ­there,” why be inside? I was not always successful at avoiding the tedium of paperwork and administration—­but then I was able to take to the woods, creeks, deserts, and mountains. Many ­people are drawn to the beauty of nature, but I was drawn also to its silence. I am never happier than when I am in some wild beautiful quiet place. I cannot say when I saw my first frog, but I must have been very young. Growing up in mostly rural northern ­Virginia in the late 1950s and 1960s provided a wealth of habitats to explore. I remember hearing the singing American Toads and catching leopard frogs along the creek near my ­house in places that no longer exist. That started me on a long journey that has taken me to six continents, fifty states, three Canadian provinces, and many Ca­rib­bean islands. At ­every turn I found frogs, and each place left its own special memories: listening to Bird-­voiced Treefrogs in a Mississippi swamp, trying to photo­graph a large ranid in ­Kenya and suddenly realizing I was lying on an ant mound (do the dance!), searching unsuccessfully for gastric-­brooding frogs in Australia’s tropical rainforest, finding rare frogs in the Seychelles ­under a starry sky with fruit bats squawking in the trees, finding Scutiger tadpoles at 5,000 m in Tibet and foam-­nesting Rhacophorus in Taiwan, and seeing my first poison dart frogs in Costa Rica. I still get a thrill seeing a Barking Treefrog or hearing the soft chirping of Eleutherodactylus planirostris around our front porch in Florida.

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Frogs are truly gentle beings and, as the New Zealanders say, “Fact: the survival of the Earth depends on frogs.” Throughout my ­career, I have been fortunate to work with many creative, enthusiastic, and knowledgeable friends, colleagues, and students. Although they may not have contributed directly to this book, it could not have been written without their guidance and friendship at least at some point over the past 30 years. I specifically thank Butch Brodie, my major advisor at Clemson University so many de­cades ago. ­There is nothing to stimulate interest and excitement better than someone who is interested and excited about their work. Ronn Altig secured my first teaching position, Hobart Smith provided encouragement when I needed it, Jim Williams got me a job in conservation, Ernie Liner talked books and made ­great food, Bob Shoop and Carol Ruckdeschel offered perspective and wine, and Dick Franz got me started again in research ­after an eight-­ year administrative stint. Thanks to you all. I thank the following persons, in par­tic­u­lar, for providing help with lit­er­a­ture and information used in the first edition of this book: Kraig Adler, Ronn Altig, Kim Babbitt, Jamie Barichivich, Breck Bartholomew, Aaron Bauer, James Bettaso, Jeff Briggler, Robin Jung Brown, Charles Bursey, Bruce Bury, Brian Butterfield, Christine Campbell, Celia Chen, Michael Conlon, Steve Corn, Christopher Distel, Nathan Engbrecht, Edward Ervin, Gary Fellers, Don Forester, Tony ­Gamble, Dana Ghioca, James Gibbs, Harry Greene, Patrick Gregory, Jackson Gross, Gavin Hanke, John Himes, Steve Johnson, Larry Jones, Tom Jones, William Karasov, Vicky Kjoss, Kenney Krysko, Michael Lannoo, James Lazell, Emily Lemmon, Lawrence Licht, Ernie Liner, Lauren Livo, Mickey Long, Clark Mahrdt, Chris McAllister, Malcolm McCallum, Jonathan Micancin, Joseph Mitchell, John Moriarty, Erin Muths, Max Nickerson, Deb Patla, Thomas Pauley, Greg Pauly, Chris Pearl, David Pilliod, Andrew Price, Louis Porras, Michael Redmer, Phil Rosen, Mark Roth, Ray Saumure, David Seburn, Lynnette Sievert, Hobart Smith, Lora Smith, Scott Smyers, Mike Sredl, Dennis Suhre, Dean Thompson, Stan Trauth, Susan Walls, Kent Wells, and Richard Yahner.

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x  Preface to the First Edition

I thank Monica McGarrity for preparing the maps, Camilla Pizano and Breck Bartholomew for the illustrations, and Audrey Owens for the beautiful frontispiece. I appreciate the photo­graphs sent by many colleagues, even though I was unable to use them all. Specific thanks go to: Ronn Altig, Jennifer Anderson-­Cruz, Adam Backlin, Jamie Barichivich, Sean Barry, Breck Bartholomew, James Beck, Dawne Becker, Steve Bennett, David Bishop, Steve Brady, Koen Breedveld, Christopher Brown, Jennifer Brown, Travis Brown, Jason Butler, Robert Byrnes, Jan Caldwell, Alessandro Catenazzi, Adam Clause, Steve Corn, John Cossel, Alan Cressler, Eric Dallalio, Nina D’Amore, Ray Davis, Chris Dellith, David Dennis, Dana Drake, Andrew Durso, Anna Farmer, Gary Fellers, Mike Forstner, Justin Garwood, Carl Gerhardt, Jim Godwin, Earl Gonsolin, Tom Gorman, Mike Graziano, Matt Greene, Kerry Griffis-­Kyle, Kyle Gustafson, Robert W. Hansen, James Harding, Chris Harrison, Marc Hayes, Valentine Hemingway, Aubrey Heupel, Jody Hibma, Brian Hubbs, John Jensen, Steve Johnson, Mike Jones, James Juvik, Kris Kendell, Ceal Klingler, Roland Knapp, Fred Kraus, Brian Kubicki, Jeff LeClere, Twan Leenders, Kirk Lohman, Patrice Lynch, Bryce Maxwell, Jonathan Mays, Brome McCreary, Melanie McFarland, Maija Meneks, Gabe Miller, Joe Mitchell, Martin Morton, Gary Nafis, Robert Newman, Justin Oguni, Charles Painter, Cindy Paszkowski, David Patrick, Seth Patterson, Joe Pechmann, Ryan Peck, Maria Pereyra, Jeff Petersen, Todd Pierson, David Pilliod, Brian Pittman, Jesse Poulos, Jim Rorabough, Francis Rose, Kevin Rose, Paddy Ryan, Rob Schell, Sara Schuster, Cecil Schwalbe, Betsy Scott,

Richard Seigel, Brad Shaffer, Nathan Shepard, Brent Sigafus, Bill Stagnaro, David Steen, Cameron Stevens, Dirk Stevenson, Jim Stuart, Dennis Suhre, Cynthia Tate, Robert A. Thomas, Stan Trauth, John Tupy, Michael van Hattem, Sara Viernum, Laurie Vitt, Kenny Wray, and Bob Zappalorti. Lee Brumbaugh of the Nevada Historical Society, Kraig Adler, and Mark Jennings ­were helpful in obtaining photos of Lithobates fisheri habitat in the Las Vegas Valley; Stephanie Munson of Cornell University Press granted permission to reproduce the Wrights’s photo­graphs of Lithobates fisheri. Ellin Beltz permitted me to use her information on amphibian etymologies and updated generic names. Other colleagues provided advice and technical assistance, particularly Kraig Adler, Jamie Barichivich, Sherry Bostick, Priya Nanjappa, Ken Sulak, Susan Walls, and Jim Williams. This book could never have been completed without the outstanding ser­vices provided by the University of Florida library system, particularly in finding t­ heses, dissertations, and sometimes very obscure papers. Vincent Burke of the Johns Hopkins University Press got me into this proj­ect in the first place, and Helen Myers ensured the readability and accuracy of the manuscript—­I c­ an’t thank her enough. To all of you, I offer my sincere thanks. Fi­nally, I thank Marian Griffey, Dick Daniell, Antoinette Marie, Benjamin Silas, Christopher Robin, Etta Mae, Fallicity Sue, Frederick Hercules, Guinevere Faye, Gwyneth Grace, Herman Bartholomew, MacMurphy O’Leary, Sir Reginald Michael, and Mortisha Marie for their special support and encouragement.

Preface to the Second Edition

In 1933, Anna and Albert Wright published the first continent-­wide review of the biology of North American frogs. It was a slim volume of 231 pages. A second edition of 286 pages was published in 1942, with a third edition in 1949 of 640 pages. It is in­ter­est­ing to note the si­mul­ta­ neously occurring world events—­the ­Great Depression, World War II, and the aftermath of the war as the economy recovered. Information increased rapidly in the late 1940s as world events allowed for focus in North Amer­i­ca on the natu­ral world, ­free from the threat of total economic collapse and the horrors of war. I suspect that the ­earlier restrictions on travel and finances, however, also gave the Wrights the opportunity to assem­ble their data and make them available to their colleagues, their students, and the public. Much of the preparation of the second edition of Frogs has taken place during the coronavirus pandemic of late 2019–2021. Although many parts of the world have been in a lockdown state for considerable periods of time—­causing all sorts of prob­lems—­the lockdown offered me the opportunity (and excuse) to focus my full attention on revising the text. I have incorporated a large number of publications from ca. 2011, when the original text was submitted, to mid-2021; more than 1,100 additional publications have been examined. This book is not a field guide, but a compendium about what is known about our North American frog fauna. I hope it stimulates both research and conservation. As the pace of scientific knowledge is accelerating, so is our knowledge of the biology and status of our North American frogs. Unfortunately, the news is not good, as our frogs continue to experience unpre­ce­dented threats and declines as the Anthropocene continues unabated. During the coronavirus pandemic, the real­ity of human-­nature interconnectedness became apparent, but it remains to be seen ­whether the importance of biodiversity, ecosystem function, and climate ­will be recognized, and ­whether the need to curb population, toxic substances, and the ­wholesale waste of our planet’s resources ­will lead to a reexamination of what is impor­tant in life. Solutions are in science and in ­will, not in the misinformation of current politics. Archie Carr was an optimist; I am not.

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An undertaking of this magnitude (i.e., the revision of Frogs) cannot be approached in isolation. Unlike the Wrights, I have been fortunate to have nearly instantaneous access to ­people, publications, and information that it would have taken them months or years to assem­ble. In addition to ­those acknowledged below, I thank the University of Florida library system that somehow managed to function during the tightest days of COVID lockdown. I ­don’t know how they did it, but thanks a million. My wife, Marian Griffey, granted permission to republish her poem In the Eyes of a Toad, originally published in Dire Elegies, 59 Poets on Endangered Species in North Amer­i­ca (2006, Foothills Publishing). Monica McGarrity again prepared the maps, and Audrey Owens graciously offered a new frontispiece, “Rain Response,” for the second edition featuring the Arizona Treefrog. ­These three deserve special credit for making this volume a success. I thank the following individuals for providing information and advice during preparation of the second edition: Jim Andrews, David Blackburn, Vernon Bleich, Erica Crespi, Marty Crump, Charles Drost, Nathan Engbrecht, Jeremy Feinberg, Jason D. Gibson, Andy Gluesenkamp, Jim Godwin, Colin Goodman, Jeff Hall, Robert Hansen, David Hillis, Andrew Holycross, Jason Hoverman, Mark Jennings, Mike Lannoo, David Laurencio, Melissa Lawton (Museum of Northern Arizona), Scott Lindemann, Dale Marcum, Chris McMartin, Walter Meshaka, Erin Muths, Matthew Niemiller, John Orr, Audrey Owens, John Palis, Dennis Parmley, Tom Pauley, Trevor Persons, Dave Pfennig, Steve Price, Steve Roble, Mark-­Oliver Rödel, Jim Rorabaugh, Brian ­Sullivan, Jeffrey Wimsatt, and Fred Zaidan. The following persons offered photo­graphs for the second edition: Ronn Altig, Marion Anstis, Jamie Barichivich, Tim Burkhardt, Jan Caldwell, David Dennis, Andrew Didiuk, Kevin Enge, Jeremy Feinberg, Brad Glorioso, Colin Goodman, Marian Griffey, Clint Guadiana, David Hillis, Jake Hutton, Steve Johnson, Thomas R. Jones, Kris Kendell, Alexa Killion, Erik Kiviat, Patrick Kleeman, Kenney Krysko, Chris Lechowicz, Emily Lemmon, John MacGregor, Jessica Martin, Gary Nafis, Piotr Naskrecki, John Palis, David Pfennig, Michael

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Introduction

I have always liked frogs . . . ​I like the looks of frogs, and their outlook, especially the way they get together in wet places on warm nights and sing about sex.

Archie Carr, The Windward Road As this book is completed (1 July 2021), ­there are 7,370 species of frogs known worldwide with new species being described at a rapid pace. North of the United States–­México border, however, only 106 species occur naturally (1.4% of the world total), and 13 of them barely enter the United States. All of Hawai’i’s frogs are introduced species, and only 27 species occur in Canada, with many only just entering the country. Only the genera Acris, Ascaphus, Hyliola, Pseudacris, and Spea may be considered endemic to our region, although Hyliola, Pseudacris, and Spea also cross the border into northern México. Centers of species richness of frogs in temperate North Amer­i­ca include the Atlantic and Gulf Coastal Plains, the Pacific Northwest, and the arid Southwest. Although it might appear that the diversity of North American frogs is reasonably well known, one new species from the Southeast has yet to be described (see Lithobates catesbeianus account), a new species was described in 2012 from near New York City, three new toad species ­were described from the western United States in 2017 and 2020, and a new Pseudacris species was described from Alabama in 2020; a number of other subspecies or isolated populations seem likely to be reevaluated as full species. No one can ­mistake a frog for any other vertebrate. All frogs are tetrapods and have the same basic body plan (Figs. 1, 2), with a short head, large bulging eyes (reduced in some fossorial forms), ­little or no neck, and a short compact body. The front legs have four toes on each foot; the rear legs have five toes on each foot and are often webbed to some extent, except in terrestrial forms. Frogs, of course, lack tails as adults. Their short power­ful bodies with specialized hind legs are made for hopping, jumping (sometimes in ­great leaps), ­running, climbing, swimming, or burrowing. Toads (Anaxyrus, Incilius, Rhinella), for example, usually walk or hop, whereas the so-­called true frogs (Lithobates, Rana) and chirping frogs (Eleutherodactylus) are expert leapers;

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Fig. 1.  Basic body plan of an aquatic frog. Illustration by Camila Pizano

Fig. 2.  Basic body plan of a toad. Illustration by Camila Pizano

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xiv  Introduction

Scaphiopus and Spea are burrowers with specialized digging tubercles (Fig. 3); Xenopus are swimmers; and Dryophytes, Smilisca, and Osteopilus are expert climbers. The toes of frogs normally are long and thin, except within the chorus and treefrog ­family (Hylidae). In this group, the ends of the toes are expanded slightly in cricket and chorus frogs (Acris, Hyliola, and Pseudacris) and greatly in treefrogs (Dryophytes, Osteopilus, Smilisca). The treefrogs use their expanded toe tips to climb trees and smooth surfaces; the expanded surface area of the toe pad exerts hydric friction ­toward the surface on which the frog walks, enabling it to hold on and climb. The largely ground-­ dwelling chorus frogs do not need such expanded toe pads, although Spring Peepers climb high in trees in Florida and elsewhere. Aquatic species usually have a membranous web between their hind toes that facilitates swimming, but Xenopus laevis has webs between the front toes as well. A comparison of the hind feet of bufonids, hylids, and ranids is provided in Fig. 4. Frogs do not have external ears, although they are very attuned to sound. The opening to the inner ear is covered by a thin, membranous sheath of skin, the tympanum, located ­behind the eyes. The tympanums of American Bullfrogs, for instance, are rounded and large; they are con­spic­u­ous on ­either side of the head. The tympanums of some other species are less easily seen or may not be pre­sent (Ascaphus). Many frogs have “warts”—­bumps or ridges on the back (the dorsum)—­dorsolaterally, or on the upper portions of the limbs. ­These bumps and ridges usually contain mucous or

Fig. 4.  A comparison of the hind feet of bufonids (left), hylids (center), and ranids (right). Note the differences in the extent of webbing and the presence of toepads in hylids. Illustration by Camila Pizano

granular glands and are impor­tant for moisture retention and defense. The parotoid glands of toads are kidney bean–­shaped structures located on the head ­behind the eyes, and their shape and size are impor­tant, in conjunction with the configuration of the cranial crests, for the identification of the dif­fer­ent species of toads (see Fig. 2). Male frogs of many species can be distinguished from females by the presence of vocal pouches and a darkened coloration on their throats, at least during breeding season. Males also develop enlarged and roughened thumbs during the reproductive season, which are useful while amplexing females (that is, when the male clasps the female during courtship). Female frogs are often much larger than males, and usually do not produce the “warning croak” when picked up. Eggs may be vis­i­ble through the ventral body wall. Outside the breeding season, differentiating the sexes may be difficult. Juveniles usually resemble miniature adults.

Anuran Evolution

Fig. 3.  Configuration of the spade on the left hind foot in the genus Scaphiopus (left) and Spea (right). In Scaphiopus the tubercle is cycle s­ haped, whereas in Spea it is wedge ­shaped. Illustration by Camila Pizano

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The first vertebrates to leave the ­water did so during the Devonian Period, some 350–370 million years ago. ­These animals ­were transitional between the lobe-­finned fishes and the true Amphibia. ­These amphibian ancestors moved around on land, based on the fossilized trackways that they left in soft muds. Their bony fossils clearly demonstrate that they ­were not adapted solely for living in the ­water, although they may have had fish-­like tails. Some of the transitional forms, such as the ­giant flat-­headed Ichthyostega, had impressive rows of teeth and a vertebral structure designed for flexibility and mobility outside ­water. The first true amphibians, the Labyrinthodonts, appeared in the Carbo­ niferous Period and survived ­until the Early Cretaceous Period, a span of about 230 million years. The Labyrinthodonts gave rise to both the modern Amphibia and the reptiles, and ­were

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Introduction xv

pre­sent throughout the age of the dinosaurs. Another group of primitive amphibians, the Lepospondyls, became extinct in the Early Permian Period, and left no modern descendants. Modern amphibians evolved at least by the mid-­Mesozoic Era, and Holman (2003) has discussed the importance of paedomorphosis and locomotion in the evolution of the first frogs. Frog-­like amphibians, Triadobatrachus massinoti (from Madagascar; Rage and Roček, 1989) and Czatkobatrachus (from Poland; Evans and Borsuk-­Białynicka, 2009), are known from the Early Triassic Period (about 225–245 million years ago). ­These proto-­frogs certainly resembled modern frogs in terms of body plan, but they possessed very dif­fer­ent morphological characters, such as having up to 26 vertebrae (modern frogs have 4–9), of which 10 formed a tail in Triadobatrachus. Their pelvic bones suggest ­these animals swam by kicking their hind legs and ­were unable to jump as do modern frogs. The earliest known true frog (Prosalirus bitis) was found in Lower Jurassic Kayenta Formation deposits in Arizona (Shubin and Jenkins, 1995). ­These deposits date from 190 million years ago. Thus, both primitive and modern amphibians ­were pre­sent throughout the Mesozoic Era, and the first known true frog likely evolved in what is now North Amer­i­ca. The basic body plan of modern frogs was set nearly 200 million years ago, although the earliest frog-­like amphibians likely walked more than they hopped or jumped. When dinosaurs ruled the world, frogs called from the swamps. More information on the early evolution of frogs can be found in Volume 4 of the Amphibian Biology series (particularly Roček, 2000; Roček, and Rage, 2000) and in Carroll (2009).

entirely aquatic species in North Amer­i­ca are the introduced African Clawed Frog, Xenopus laevis, and the Tropical Clawed Frog, X. tropicalis. All members of the mostly tropical genus Eleutherodactylus deposit eggs terrestrially. The larval period occurs entirely within the egg, and hatchlings resemble miniature adults. Only a few other frogs depart from the typical pattern. In the nonindigenous Dendrobates auratus of Hawai’i, eggs are oviposited in moist situations and fertilized by the male who then guards them ­until they hatch. He then carries the tadpoles on his back to an appropriate developmental site, such as a treehole or bromeliad cup. In Leptodactylus fragilis, eggs are oviposited into a foam nest and tadpoles are released as ­water eventually fills the shallow nest depression. All frogs in Canada and the United States have external fertilization except for Ascaphus (internal fertilization via an intromittent organ) and Eleutherodactylus (through cloacal apposition). The pro­cess of holding onto a female during reproduction is termed amplexus. The male frog grabs the female dorsally ­either ­under the armpits (axial or pectoral amplexus; Fig. 5) or just in front of her hind legs (inguinal or pelvic amplexus) and holds on as strongly as he can. The location where the male grabs the female is species specific. As eggs are extruded from the female’s vent, the male releases clouds of sperm over them. Male frogs often amplex the wrong species, other males, or inanimate objects. If another male is amplexed, he gives a warning croak and/or vibration to let the courting male know that he has erred in his mate choice. Other­wise, a male amplexing an inappropriate object often has a long and frustrating reproductive season. Spent females also may give a warning vibration, but

Life History North of the Mexican border, frogs occur from sea level to the high Rocky Mountains, and from the south Florida Keys to the Arctic Ocean. They occur in tropical lowlands, grassland prairies, deserts, and in alpine-­tundra habitats. Although thought of as entirely freshwater in nature, a few frogs and tadpoles have been found associated with saline habitats near oceans and in isolated desert wetlands. Some species have restricted habitat requirements, whereas ­others occur contiguously from the arid ­Great Plains (Pseudacris maculata) or humid Southeastern forests (Lithobates sylvaticus) to the high tundra. In the “typical” amphibian life cycle of most US and Canadian frogs, adults move to ­water to breed and lay eggs, then emigrate back to terrestrial or other aquatic habitats to forage and overwinter. The eggs develop into larvae that remain in ­water for a period of time, then the larvae metamorphose, and the tiny juveniles disperse. The only

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Fig. 5.  Axillary (pectoral) amplexus in aquatic frogs. Illustration by Camila Pizano

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xvi  Introduction

many amplexing males seem to be able to determine ­whether a female is gravid by her girth and perhaps the firmness of her body as she carries eggs. Nearly all North American anurans lay eggs in ­water. A few (Dendrobates auratus, Eleutherodactylus sp.) deposit their eggs in moist terrestrial locations. The eggs are coated by one or more jelly-­like capsules surrounded by a firm membrane (Fig. 6). The capsules and membrane help protect the eggs from infection and desiccation while allowing for the exchange of oxygen, carbon dioxide, and nitrogenous wastes (mostly ammonia). Eggs may be deposited individually, in small clusters, in strings, in thin sheaths that float on the surface of ­water, or in large compact jelly masses (Figs. 7–9). In addition, eggs may

Fig. 8.  Surface film of frog eggs. A number of ranids and microhylids deposit their eggs in single-­layer surface films. Illustration by Camila Pizano

Fig. 6.  Diagram of a frog egg. The vitellus is surrounded by two jelly envelopes enclosed within encapsulating membranes. Illustration by Camila Pizano Fig. 9.  Toad egg strings. Note that in some species the eggs are partitioned into separate chambers (left), whereas in ­others the eggs are continuous, forming one or more rows without partitioning (right). Illustration by Camila Pizano

Fig. 7.  A frog egg mass. Egg masses of this type are typical of many ranids. Illustration by Camila Pizano

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or may not be attached to aquatic vegetation or to the wetland bottom. Depending on the species, the number of eggs deposited can be from a few to as many as tens of thousands. All ­these characteristics are impor­tant in helping to identify species in the field, and they are mentioned in more detail in the species accounts.

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Introduction xvii

In the early stages of development, most anuran eggs have a dark animal pole and a light cream to yellowish vegetal pole, although in a few species (Eleutherodactylus sp.) the eggs are entirely white. The cells of the animal pole ­will eventually overgrow the vegetal pole and form the developing embryo, whereas the yolk forms from cells of the vegetal pole. The nutrient-­rich yolk supplies all the energy needed to complete development prior to hatching. Hatching occurs as partially developed larvae (tadpoles) in aquatic situations, and as miniature adults in terrestrial nests (Eleutherodactylus sp.). All larvae have gills for respiration; in larval frogs, the gills are hidden shortly ­after hatching within an internal chamber covered by an operculum on the side of the head. As the larva grows, it eventually reaches the stage where it metamorphoses into an adult. Metamorphosis involves a complex set of morphological and physiological changes that radically alter the body plan. It is also the time when the animal is most likely to be affected by chemicals or pollutants in the environment that interfere with the intricate developmental changes. Anurans remain as larvae from a few days (e.g., Eastern Spadefoot) to 3–5 yrs (e.g., Coastal and Rocky Mountain Tailed Frogs). Most larvae, however, transform within a few months to a year. If breeding occurs in spring or early summer, transformation usually occurs in late summer to autumn. If breeding occurs in the fall, the larvae overwinter and metamorphose the following spring or summer. ­There is a ­great deal of variation in the larval period among species, however. Some of the variation undoubtedly is derived from hereditary ­factors, although the availability of high-­quality food and the duration of the hydroperiod also influence the length of the larval period. Unlike salamanders, ­there are no paedomorphic frogs. Tadpoles are mostly detritivores in that they eat a wide variety of benthic organic ­matter. They graze on algae or aquatic vegetation using a specialized mouth structure that rasps small bits of vegetation into the mouth. This material is funneled into the gut for digestion. Some tadpoles are filter feeders and feed in the ­water column or at the surface, specializing on phytoplankton and zooplankton, but ­others are carnivorous or even cannibalistic. Adult frogs are all carnivorous, usually eating a wide array of insects, spiders, and other invertebrate animals. A few species, such as American Bullfrogs, consume large prey such as small mice or snakes, whereas ­others, such as the ant-­loving Eastern Narrow-­mouthed Frog, are specialist feeders. Larvae are largely ­free swimming and do not form social aggregations. Spadefoots, some toads (Anaxyrus sp.), and the River Frog (Lithobates heckscheri) are exceptions, however, in that their tadpoles form large schools that move and feed together throughout a pond or lake. Schooling helps to churn up the bottom and expose more food for

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consumption, provides a mea­sure of protection, and maintains a favorable thermal environment. Spadefoot schools in par­tic­u­lar are initially comprised of kin—­that is, tadpoles that hatched from eggs deposited by the same parents. They apparently keep in contact via pheromones, chemicals that aid them in sibling recognition. Tadpoles also are known to respond to alarm chemicals within their environment. If another tadpole is attacked and injured, the chemical released by the injured tadpole alerts the rest of the tadpoles to danger in the area. Larval anurans have a smooth and moist skin that readily allows ­water to enter and waste products to be expelled. During the terrestrial phase of their life cycle, however, frogs must remain in moist situations ­because their skins are only semi-­impermeable to ­water loss. In dry conditions, moisture is lost to the surrounding air from the animal’s body, and frogs ­will desiccate if too much ­water is lost. Anuran skins are usually moist to the touch ­because of mucous secreted to keep them damp. The skins of some species, such as toads, may appear rather dry, however. A few species are able to spread a lipid-­based secretion across the body to impede ­water loss. All North American frogs possess sac-­like lungs, but gas exchange takes place across the skin of larvae and adults. Some areas of the skin are particularly well vascularized (e.g., the pelvic patch of certain frogs), and gases and ­water readily diffuse across the thin skin membranes. Frog skin also is remarkable in terms of defense. Many species possess ­either toxic (Dendrobates auratus, Rhinella marina) or noxious (e.g., Anaxyrus sp., Scaphiopus sp., Lithobates palustris) secretions that help them avoid predators and parasites. Noxious and toxic skin secretions are produced from granular glands in the skin of frogs. Such glands may be concentrated, such as in the “warts” of toads, or they may be more diffusely distributed over the dorsal part of the body. Toads (Fig. 2) and spadefoots (Fig. 10) have kidney bean–­shaped glands ­behind the head that contain glands that produce noxious secretions. ­These parotoid glands have even been observed to squirt secretions in the direction of an attacking predator, especially as the predator bites down on the frog. Other anurans (many Lithobates, Rana, Ascaphus sp.) have antimicrobial peptides in their skin that help ward off disease and fungi. Chemical communication is not as impor­tant in terrestrial frogs as it appears to be for salamanders, at least as adults. Larvae and perhaps aquatic adults seem to be keenly aware of chemical cues in ­water. Most male frogs in the United States and Canada gather at breeding sites from winter to late summer and send out chirps, peeps, and trills depending on species. Only Ascaphus is truly ­silent. The calls serve two main functions: to entice females to the pond and to inform other males that the caller

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xviii  Introduction

Fig. 10.  Head morphology of a Spadefoot showing the position of the boss, parotoid gland, tympanum, and vertical eye pupil. Illustration by Camila Pizano

has occupied a par­tic­u­lar portion of the breeding site. Thus, what sounds to a ­human listener like a single call conveys multiple messages to conspecifics. For example, a female may assess male quality by the sound of his call, how long it is, how loud it is, or how deep it is. Inasmuch as some frogs are territorial at breeding sites, the calls also serve to alert other males that the caller is ready to defend his calling space. Females often prefer the largest males when presented with a choice. However, “satellite” males (males that do not call but sit near a calling male) sometimes intercept females on their way to breed, and thus avoid the competitive chorus. Frogs generally return to the same ponds or wetlands to breed from one year to the next, although ­there are exceptions. In some frogs, both sexes may use the same ponds each breeding season. In other species, the males are site specific, but the females are not. In this way, a female can choose the best-­fit male among all the males that she can hear, regardless of where he is located. Male pond-­breeding frogs usually arrive at the ponds first and establish their calling sites and territories, and such males may overwinter closer to the ponds than females, presumably so they can get to the breeding sites early. Males also stay at ponds longer than females, who frequently stay only long enough to mate and deposit their eggs. In explosive breeders (that is, frogs with short reproductive periods in which nearly all adults breed at the same time), males and females arrive nearly si­mul­ta­neously. At least some species of frogs breed in about any type of wetland. Frogs that breed in small temporary or semiperma-

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nent ponds do so ­because ­these habitats lack fish and may have fewer invertebrate predators, especially early in the season. In areas lacking aquatic vertebrate predators, larvae are often con­spic­u­ous; in habitats where predators are abundant, larvae are secretive and cryptic. Many of the ponds and streams in North Amer­i­ca are temporary or intermittent, especially in the ­Great Plains and the West. Even in the East, small breeding pools often dis­appear by summer. As the year progresses, ­these habitats dry up, making them unsuitable for breeding. Drought and low-­ rainfall years can also pose prob­lems, and in some years frogs may not be able to breed. Thus, whereas frogs breeding in temporary wetlands may reduce predation risk to their larvae, ­there is a trade-­off as to ­whether the wetlands ­will fill and allow sufficient time for the larvae to develop and metamorphose. Many frogs travel short distances from their breeding sites, whereas ­others travel hundreds of meters or even several kilo­meters from where they passed the larval stage. Terrestrial habitats may be as close as streamsides, making certain frogs (e.g., Ascaphus) seem semiaquatic. Other frogs (for example, the Western Toad, Anaxyrus boreas) travel extensively before establishing a nonbreeding terrestrial home range. ­Here, they spend their time in surface, subsurface, cliff face, or arboreal habitats before returning to the area where, presumably, they hatched. As a rule, the shorter lived a frog is, the sooner it begins to breed. Still, not ­every adult frog breeds ­every year, and ­little is known about reproductive periodicity through their life-­span. I have observed a female Florida Eastern Narrow-­mouthed Frog skip a breeding season, even though the radioactive-­tagged animal was near a pond, and the life-­span of this species is only about 4 yrs. Females, in par­tic­u­lar, seem to skip breeding in years of harsh environmental conditions and poor food availability, and instead keep their energy reserves to stay alive. Anurans are especially active at night, although ­those species with noxious secretions (especially toads) and a few ­others (e.g., Acris) are often active by day. Aquatic frogs (e.g., Lithobates, Rana) are especially active diurnally on cloudy and rainy days, but they are usually in ­water and tend to be very alert. Frogs congregate around all kinds of wetlands, emerge from ­under rocks and logs, and forage arboreally or through the terrestrial leaf litter or along stream edges. With a flashlight, one can often spot frogs sitting on buildings, at the mouths of burrows and hiding places, and in ­water or along the shoreline waiting for unsuspecting invertebrates. At night, larvae also become more con­spic­u­ous as they leave daytime refugia for open and shallow ­water. Nocturnal activity presumably makes some frogs less prone to predation, especially by visually oriented

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Introduction xix

predators. Only on rainy nights can the abundance of anurans be appreciated as they call and forage.

Anuran Conservation Frogs play significant roles in ecosystem function, particularly as predators of invertebrates, prey for a vast array of other species, and in nutrient and energy transfer to and from habitats rich in nutrients (e.g., ponds) to areas poor in nutrients, such as sandy uplands (Earl et al., 2011; Hocking and Babbitt, 2014; Capps et al., 2015). Unfortunately, frogs are now at greater peril worldwide than at any time in recent geologic history (Stuart et al., 2004; Sodhi et al., 2008; Wake and Vredenburg, 2008). In the United States alone, overall amphibian population declines reached 3.7% between 2002 and 2011 (Adams et al., 2013). Indeed, the Earth’s biota may already be well into the sixth mass extinction event since life began on this planet (Barnosky et al., 2011). It is beyond the scope of this introduction to discuss in depth the many threats to anuran populations in Canada and the United States and the ways to mitigate and manage ­these threats. Much more detail can be found in Stebbins and Cohen (1995), Kingsbury and Gibson (2002), Semlitsch (2003), Lannoo (2005), Bailey et al. (2006), Mitchell et al. (2006), Pilliod and Wind (2008), Dodd (2010), and in the references in the species accounts. Threats to anuran populations derive from local, regional, and global sources. Wetland and associated terrestrial habitats are being lost and fragmented (or shredded, as some biologists have termed it) at alarming rates ­because of expanding ­human populations and economic and po­liti­cal considerations fostering rapid and often poorly regulated development (Hamer and McDonnell, 2008). In the United States alone, 185,400 ha of wetlands ­were destroyed per year from the mid-1950s to the 1970s, with another 117,400 ha lost per year from the mid-1970s to the 1980s; another 155,200 ha ­were lost from 1986 to 1997 (Dodd and Smith, 2003). In the ­Great Plains, optimal amphibian habitat declined by 22% from 2007 to 2012, or 3.8 million ha to 2.9 million ha (Mushet et al., 2014). ­These numbers are frightening when considering the potential loss of aquatic diversity and individual frogs. Habitat changes affect amphibians over a long time period, so it is necessary to consider both historic and current landscape effects on frog populations in order to conserve them (Piha et al., 2007). Anurans, with their unprotected eggs, aquatic larval development, permeable skins, complex endocrinological and morphologic changes associated with metamorphosis, diverse life histories, and biphasic life cycles requiring both terrestrial and aquatic habitats, are being saturated by a host of lethal and sublethal pesticides and other toxic substances

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(Quaranta et al., 2009). Emerging infectious diseases such as amphibian chytrid fungus (Batrachochytrium dendrobatidis), ranaviruses, and novel alveolate pathogens threaten worldwide impacts (e.g., Fisher and Garner, 2007; Vredenburg et al., 2010b; Gray and Chinchar, 2015; James et al., 2015; Earl et al., 2016; Kolby and Daszak, 2016; Isidoro-­Ayza et al., 2017; Rollins-­Smith, 2020). Indeed, amphibian chytrid is now so widespread that it can be considered endemic across North Amer­i­ca (Lannoo et al., 2011). Unfortunately, ­there are still no effective methods to reduce the impacts of chytridiomycosis, and the strategies that have been proposed have not been tested in the field and face a variety of economic, logistic, societal, and biological hurdles to their implementation (Woodhams et al., 2011; Garner et al., 2016). In addition to diseases, many malformations have appeared in amphibian populations (Lannoo, 2008; Reeves et al., 2013). Threats such as global and regional climate change affect both temperature and precipitation patterns and further imperil frogs, especially ­those with ­limited distributions and dispersal capabilities (Corn, 2005; Lawler et al., 2009; Barrett et al., 2014). Invasive plants and animals offer additional novel threats to amphibian habitats and species (Bucciarelli et al., 2014). As with all biota, threats faced by amphibians do not act in a vacuum. Threats may act synergistically, that is, the ­factors are additive, thus accelerating the rate of decline (Blaustein et al., 2011; Hof et al., 2011; Buck et al., 2012; Battaglin et al., 2016; Grant et al., 2016, 2020). Fully one-­third of all amphibians are now considered threatened worldwide (Stuart et al., 2004), and 168 amphibians have become extinct within the last several de­cades. Clearly, ­these are treacherous times for many frogs, as many of the species accounts ­will indicate. Anuran conservation requires an integrated landscape approach to management, rather than a species-­oriented focus, except ­under dire circumstances (e.g., Semlitsch, 2000; Grant et al. 2013; Thompson et al., 2015). The reason for this is ­simple: frogs do not live in nature in a biotic or physical vacuum. They frequently move from wetlands to terrestrial sites, and movements may cover hundreds of meters. This necessitates the protection of breeding sites, movement corridors, and surrounding terrestrial habitats used for foraging and overwintering (Cayuela et al., 2020). Simply “protecting” breeding sites ­will not conserve anuran populations. It once was thought that if frogs and habitats could be protected against take or destruction, then frog populations would survive. Threats from disease, toxic chemicals, and climate change clearly have demonstrated the inadequacy of such an approach. The ge­ne­tic consequences of habitat fragmentation also are much better understood, so connectivity among even protected sites must be maintained

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despite serious habitat fragmentation (Cushman, 2006; Becker et al., 2007). Habitats cannot be surrounded by a fence or even be inside well-­protected parks with the assumption they ­will maintain their ecological integrity (Nori et al., 2015). Frog management options range from the rather ­simple and inexpensive to the very complex and expensive. When planning, the overriding consideration should be “first, do no harm” to the species, its community, or its habitat. High technology–­based approaches may work no better than ­simple and inexpensive approaches, and care should be exercised to maximize the benefits from the ­human resources and funds available. The primary objectives of management should always focus on the species or community of concern, and not on peripheral or extended objectives, such as positive publicity. Even when extensive restoration and other management efforts are undertaken, it may take a long time for them to show positive results (e.g., Walls et al., 2014b). All state and provincial governments have statutes aimed at protecting endangered and threatened species. The effectiveness of regulations varies considerably among jurisdictions, and rarely is habitat considered on an equal basis with individual protection. For listed species, wildlife and conservation agencies frequently attempt to develop conservation and recovery plans, but ­these plans are vastly underfunded. When compared to highly publicized species such as manatees, frogs receive a pitiable amount of financial support for research and conservation. Unfortunately, many individuals and organ­izations oppose environmental land purchase and protection ­under the guise of economic recovery, which masks—­such as in Florida—­a deep-­seated opposition to impediments to wealth accumulation. Such myopic foresight ­will have devastating consequences for frogs and their habitats. As of April 2022, the following species are protected ­under provisions of the US Endangered Species Act of 1973, as amended, as Endangered or Threatened: Wyoming Toad (Anaxyrus baxteri), Arroyo Toad (A. californicus), Yosemite Toad (A. canorus), Houston Toad (A. houstonensis), ­Dixie Valley Toad (A. williamsi), Chiricahua Leopard Frog (Lithobates chiricahuensis), Dusky Gopher Frog (L. sevosus), California Red-­legged Frog (Rana draytonii), Mountain Yellow-­legged Frog (R. muscosa, both northern and southern populations), Oregon Spotted Frog (R. pretiosa), and Sierra Nevada Yellow-­legged Frog (R. sierrae). The following should be considered candidate species: Railroad Valley Toad (A. nevadensis), Hot Creek Toad (A. monfontanus), Illinois Chorus Frog (Pseudacris illinoensis), and Relict Leopard Frog (Lithobates onca). In 2002, the Government of Canada enacted the Species at Risk Act, its first endangered species act. ­Under that legisla-

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tion, the Committee on the Status of Endangered Wildlife in Canada (COSEWIC) decides which species are at risk. COSEWIC is funded by Environment Canada, but it other­wise operates in­de­pen­dently of the government. COSEWIC commissions studies of native species whose survival in Canada might be at risk. Based on that research, it places them in one of five categories: extinct, extirpated, endangered, threatened, or special concern. The Rocky Mountain Tailed Frog (Ascaphus montanus), Fowler’s Toad (Anaxyrus fowleri), ­Great Basin Spadefoot (Spea intermontana), Oregon Spotted Frog (Rana pretiosa), Northern Leopard Frog (Lithobates pipiens, Rocky Mountain population), Blanchard’s Cricket Frog (Acris blanchardi), and Western Chorus Frog (Pseudacris triseriata) are listed as endangered or threatened species by COSEWIC. The rise of ­human population coupled with agricultural and industrial development has been termed the Anthropocene. Although lasting only a few thousand years, direct and indirect ­human activity during the Anthropocene has dramatically threatened the environment of frogs and indeed much of nature. At the same time, ­human action is necessary to prevent the continued loss of frogs and biodiversity in general; if frogs are to survive, ­humans must take effective mea­sures to ensure their ­future. ­There are many ways to do this, from active participation in research to joining organ­izations trying to conserve what remains of Earth’s biotic diversity. Concerned individuals may become “citizen scientists,” learning the methods of research and cooperating with state, provincial, and national conservation organ­izations and government agencies in monitoring species’ status (Lee et al., 2021). ­People need to be aware of how land-­use proposals impact the function and diversity of nature, and to oppose ­those that are detrimental to natu­ral systems. Individuals must learn to think critically, keep abreast of scientific developments, develop a passion for the value of nature unto itself, and not be afraid to oppose ignorance and ideologically based hyperbole, such as that surrounding climate change and the impact of pesticides on sexual development, health, and ecosystem function. In short, be involved.

The Frog in North American Culture Frogs have had a very long association with ­humans throughout the world, ­whether through my­thol­ogy, spirituality, art, lit­er­a­ture, or scientific investigation (Crump, 2015). Their effigies occur on totem poles in Alaska, pueblo pottery, and on pipes ceremonially smoked by Native ­peoples in the Midwest and Southeast (Figs. 11–12). In post-­European contact, images of frogs from North Amer­i­ca first appeared in Catesby’s magnificent The Natu­ral History of Carolina, Florida, and the Bahamas, published in parts from 1729 to

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Introduction xxi

Fig. 11.  Zuni bowl featuring tadpole motif. Pitt Rivers Museum, Oxford, ­England. Photo: C.K. Dodd, Jr.

Fig. 13.  Green Treefrog (Dryophytes cinereus) on skunk weed. Mark Catesby: Natu­ral History of Carolina, Florida and the Bahama Islands. Issued in parts 1731–1743.

Fig. 12.  Frog pipe, Hopewell Culture, central United States. British Museum, London. Photo: C.K. Dodd, Jr.

1742 (Fig. 13). Frogs continue to provide us with im­mense inspiration and enjoyment with their calls that signal an end to winter and the rebirth of dormant nature. They eat massive quantities of injurious insects; they are subjects of research in development, regeneration, tissue repair, and ge­ne­tics; and they are prey for countless other animals within their community. Despite the presence of sometimes noxious or toxic secretions, frogs and toads are gentle creatures that can be handled safely. Archie Carr, a well-­known Florida naturalist and scientist, once noted that “any damn fool knows a catfish.” The same

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could be said of frogs. ­Because of their sheer abundance during the breeding season and at metamorphosis, and their often sudden appearance associated with rainfall, frogs are familiar to ­people throughout the continent, and likely have been since ­humans first set foot in North Amer­i­ca. Universally, they are associated with fecundity (abundance) and rebirth (the end of drought, the arrival of spring ­after a long winter). Sometimes they are representative of good, whereas at other times—­particularly in Eu­ro­pean cultures—­they are associated with evil or the underworld. In fact, they are simply benign animals ­going about their business of life. Frogs occur frequently in the stories and my­thol­ogy of First Nations ­peoples (Crump, 2015) where their biphasic life cycle suggests spiritual transformation. They ­were (and are) symbols of wisdom, knowledge, wealth, and protection to some Nations; for example, the Haida of coastal Canada and the Pacific Northwest carved frogs on ­house posts to prevent the ­house from falling down. Frogs possess the power to go between worlds and live in both the natu­ral and super­natural. Frog images appear frequently on feast bowls

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xxii  Introduction

and totem poles and in Haida art. An excellent collection of ­these is on display at the Museum of Anthropology at the University of British Columbia. Frogs occur frequently in oral traditions. In the Frog Mountain Story of the Sinixt Nation, a frog tells the ­People how to survive a ­great drought, with the frogs coming to feed the ­People ­until the land recovers. One day a frog tells the ­People in their protective cave to no longer eat the frogs, for the danger has passed. The frog gives the ­People a final gift, transforming into a ­great mountain (Frog Mountain). The moral: “We are small creatures who do not seem to impact life, threaten life, or support life in any way. We have shown through our love and our offering to you that even something believed to be small and unimportant can become a power­ful being in both deed and symbol” (see http://­ sinixtnation​.­org​/­content​/­swarakxn​-­chaptikwl​-­frog​-­mountain​ -­story). ­There certainly is a lesson ­there for every­one. In addition to symbolism, frogs also are impor­tant in the history of First Nations ­peoples. For example, the Kiks.adi (Frog) Clan of the Tlingit Nation built a fort at Sitka, Alaska, in 1804 to defend themselves temporarily against invasion by Rus­sian colonists (Urban and Car­ter, 2021; also see https://­www​.­akherpsociety​.­org​/­kiksadihistory​.­htm). In the Native American Southwest, frogs feature prominently on pottery and in legends and stories, particularly ­those associated with rain and Creation. The four frogs motif can be found on Navajo rugs (Fig. 14) and sand

Fig. 14.  Four Frogs. Navajo sandpainting weaving by Lorraine Tallman. 1.13 m × 1.13 m. Author’s collection.

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paintings. (The frog is a deity who can make floodwaters recede and who plays a role in fertility.) For the Hopi, kachinas are mythical ancestors who became spirits of rain and the promise of rain. As such, they, like the frogs of the Northwest, are intermediaries between ­humans and the spirit world. Frogs ask the kachinas for rain on behalf of the P ­ eople. Crump (2015) recounts stories of the Zuni on the origin of turtles, frogs, and snakes. ­These beings became the Council of the Gods, to whom the Zuni prayed for rain to sustain their crops. Not surprisingly, the Zuni frequently decorate their pottery with frogs and tadpoles (Fig. 12). In the Navajo story Frog Creates Rain, a frog saves the ­People from a dangerous fire by bringing ­water to put out the fire. As a reward, First ­Woman then give frogs the responsibility to call the rain, which they do to this day when they sing (Crump, 2015). In the East, frogs played impor­tant roles in the Adena, Hopewell, Mississippian, and prob­ably all other Native cultures, and their images are common on pottery, shells, and carvings. Frogs are prominent in the my­thol­ogy of the Ojibwa, Chippewa, Creek, Iroquois, Catawba, Cherokee, and many ­others (see https://­www​.­firstpeople​.­us​/FP​-­Html​-­Legends/). In a Catawba story, for example, the toad is a medicine man (Speck, 1934). As in the West, frogs are associated with ­water. The earliest Eu­ro­pean references to a recognizable frog species are Boucher’s (1664), Thomas’ (1698), Beverley’s (1705) and Lawson’s (1709) mention of bullfrogs (‘Bull Frog’). Beverley (1705) states that “Last year I found one of ­these near a stream of fresh ­water, of so prodigious a magnitude, that when I extended its legs, I found the distance betwixt them to be seventeen inches and a half.” Catesby (1731–1747) illustrates several identifiable species; Brickell (1737) mentions the ‘Bull-­Frog,’ a ‘Green Frog’ that climbs up trees, and a ‘Land-­Frog’ that is “like a Toad, only it leaps and is not poisonous”; and Winterbotham (1796) rec­ords the “toad, bull-­frog, water-­frog, green tree-­frog, bell-­frog, and small green-­frog” as inhabitants of the United States. Brickell (1737) also notes that frogs are baked and beaten into a powder and, when mixed with orrice-­root (Iris germanica), used to cure “Tympany [i.e., when the chest contains ­free air or the abdomen is distended with gas] and many other Disorders.” Brickell (1737) appears to have copied much of his information directly from Lawson (1709). In South Carolina, Ramsay (1809) noted that toads, bull frogs, frogs, and green frogs inhabited the state, setting the stage for Holbrook’s (1842) first scientific treatise on North Amer­i­ca herpetology. In con­temporary Western culture, frogs are everywhere: in trinkets, art,fashion, as symbols for wineries and breweries, as Muppets (Kermit the Frog), on coins and postage stamps (Figs. 15a, b), as school mascots (the Bullfrogs of Lake Worth High School, Texas), and in lit­er­a­ture. They are featured in famous short stories (e.g., Mark Twain’s The

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Introduction xxiii

Fig. 15a.  Canadian coins and stamps depicting frogs. Author’s collection.

Fig. 16.  Frog festival memorabilia. Top: souvenir envelope from Toad Suck Daze, Conway, Arkansas, 5 May 1996. Bottom left: “Let’s Get Associated” stamp No. 124 commemorating the first frog jumping contest in Calaveras County, California. Tidewater Oil Co. and Flying A Gas, 1930s. Bottom right: Centennial (1865-1965) commemorative token from the Calaveras County International Frog Jump, Calaveras County, California, 20-23 May 1965. Author’s collection.

over the invasion of migrating frogs (Weaver, 1857). ­There are even festivals for toads and frogs (Rayne Frog Festival, Louisiana; Toad Suck Daze, Arkansas; Calaveras Jumping Frog Jubilee, California; Fig. 16), and ­there are statues of frogs, such as in Regina, Saskatchewan, and Loveland, Colorado. Clearly, frogs have captured the imagination of ­people throughout our region and through time.

Etymology

Fig. 15b.  United States stamps depicting frogs. Author’s collection.

Notorious Jumping Frog of Calaveras County), nature writing (e.g., George Constantz’s Hollows, Peepers & Highlands, An Appalachian Mountain Ecol­ogy, 2004, West ­Virginia University Press; Glen Rounds’ The Snake Tree, 1966, The World Publishing Com­pany), poetry (e.g., Susan Fromberg Schaeffer’s The Rhymes & Runes of the Toad, 1975, MacMillan Publishing), humor (e.g., Waldie and Frobish’s Fair Play for Frogs, 1977, Harcourt Brace Jovanovich), numerous ­children’s books (e.g., Robert McClung’s Bufo, The Story of a Toad, 1954, William Morrow and Com­pany), and anthologies of toad and frog information through the ages (e.g., Gerald Donaldson’s Frogs, 1980, Van Nostrand Reinhold Com­pany; Robert DeGraaff ’s The Book of the Toad, 1991, Park Street Press). They are on the city seal of Windham, Connecticut, commemorating the ­great “­Battle of the Frogs” in 1758 when townsfolk took up arms

Dodd_Canada_int_5pgs_B1&B2.indd 23

Etymology refers to the derivation, or meaning, of a name. Species names are always italicized or underlined. They consist of both a genus and specific epithet, with the genus, but not the specific epithet, capitalized (e.g., Acris gryllus). Species names are derived from Latin or Greek roots and in accordance with Classical grammar. When they are used to honor a person or place, specific epithets must conform in gender and number to both the generic name and the name of the person or place they honor. Although ­there are a number of frivolous names in the scientific lit­er­a­ture, most authors attempt to convey some impor­tant characteristic of the animal, its location or habitat, or the p ­ eople impor­tant to its recognition and discovery. The etymologies of specific epithets are provided in the species accounts. In order to avoid duplication, generic etymologies are provided below, including the names proposed by Frost et al. (2006a); ­these are derived from Beltz (2007; personal communication) and other sources. Especially with older names, authors often did not provide an etymology with the description, so the exact meaning of the name is sometimes unclear. Acris. From the Greek akris, meaning ‘locust.’ The name refers to the call of cricket frogs, which sounds like insect calls.

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xxiv  Introduction

Anaxyrus. Possibly from the Greek anax, meaning ‘leader’ or ‘chief,’ and urus, meaning ‘tail.’ What Tschudi was referring to when applying this name is unclear. ­There are several other pos­si­ble meanings, including the Greek an, meaning ‘without,’ the Greek axyr, meaning ‘being cut’ or ‘an anchor,’ and the Greek ax, meaning ‘axel.’ Ascaphus. From the Greek a, meaning ‘without,’ and skaphis, meaning ‘spade.’ Literally, without a spade, denoting the absence of a metatarsal spade. Craugastor. From the Greek krauros, meaning ‘hard’ or ‘brittle,’ and gaster, meaning ‘of the belly.’ Dendrobates. From the Greek dendro, meaning ‘leaf,’ and bates, meaning ‘one who treads or climbs’; literally, leaf-­climber. Dryophytes. From the Greek dryos, meaning ‘tree,’ and phytes, meaning ‘plant.’ Eleutherodactylus. From the Greek eleutheros, meaning ‘­free’ or ‘unbound,’ and dactylos, referring to ‘fin­ger’ or ‘toe.’ ­These frogs lack a web between their digits. Gastrophryne. From the Greek gastros, meaning ‘belly,’ and phryne, meaning ‘toad.’ The name may refer to the fat bellies of ­these frogs. Glandirana. The name literally means ‘glandular frog’ and is in reference to the numerous glands found on both tadpoles and postmetamorphs. Hyliola. A diminutive form of Hyla. Frogs of the genus Hyla are usually larger than ­those within the genus Hyliola. Hypopachus. From the Greek hypo, meaning ‘­under,’ ‘beneath,’ or ‘lesser,’ and pachos, meaning ‘thickness.’ Incilius. Unknown. Meaning not stated in Cope (1863). Lithobates. From the Greek lithos, meaning ‘rock,’ and bates, meaning ‘one who treads or climbs’; literally, rock climber. Leptodactylus. From the Greek leptos, meaning ‘fine,’ ‘slender,’ or ‘thin,’ and daktylos, meaning ‘fin­gers’ or ‘digits.’ Osteopilus. From the Greek osteon, meaning ‘bone,’ and pileos, meaning ‘cap.’ The name refers to the co-­ossified skin on the top of the head. Pseudacris. From the Greek pseudes, meaning ‘false’ or ‘deceptive,’ and akris, meaning ‘locust.’ Hence, ‘false Acris,’ that is, a false cricket frog. Rana. Latin for ‘frog.’ Rhinella. From the Greek rhinos, meaning ‘nose,’ and ella, meaning ‘diminutive’; literally, ­little nose. Rhinophrynus. From the Greek rhinos, meaning ‘nose,’ and phrynos, meaning ‘toad.’ Scaphiopus. From the Greek skaphis, meaning ‘shovel’ or ‘spade,’ and pous, meaning ‘foot.’ The reference is to the spade of each hind foot, which is used in digging. Smilisca. From the Greek smiliskos, meaning ‘­little knife.’ The name refers to the sharp pointed fronto-­parietal pro­cesses.

Dodd_Canada_int_5pgs_B1&B2.indd 24

Spea. From the Greek speos, meaning ‘cave’ or ‘cavern.’ This prob­ably refers to the burrowing habits of spadefoots. Xenopus. From the Greek xenos, meaning ‘foreign’ or ‘stranger,’ and pous, meaning ‘foot.’

About the Book As with the first edition, this volume is not a field guide. ­There are no keys to the species of anurans, although impor­tant distinguishing characteristics are provided in each species account. Many regional, state, and provincial field guides provide keys to adults and tadpoles, and additional information regarding identification can be found on numerous Internet sites. General references to many other aspects of the biology of frogs are included in the list of books, Internet sites, and atlases at the end of this section. My objective in this book is to synthesize the lit­er­a­ture on all frogs of North Amer­i­ca north of the Mexican border through June 2021. In that regard, I have concentrated on what is known about frogs within our region, even when the range extends farther south as it does for many tropical species that reach their northern limits just north of the México–­United States border. The life history of tropical species may be dif­fer­ent at the northern periphery of their range than it is hundreds or thousands of kilo­meters southward. Summary references to the biology of peripheral and nonnative species are provided in the species accounts. The following synopsis outlines the basis for the information provided, in order to delineate the book’s scope and limitations. Nomenclature. The generic names of toads (Bufonidae), treefrogs (Hylidae), and ­water frogs (Ranidae) are in a considerable state of flux. The inclusion of molecular analyses involving mtDNA and nDNA—­using large datasets and often dif­fer­ent and rapidly evolving methods of analy­sis and interpretation—­has led to intense debate about how to name phyloge­ne­tic lineages. Should genera be named based on strict monophyly, and at what level of branching? Which names should be used in order to best reflect evolution and to provide information concerning ­those lineages? Should long-­used and familiar names (e.g., Bufo, Hyla, Rana) be changed to reflect phyloge­ne­tic branching, which in turn provides biogeographic and temporal information about when, where, and how the lineages evolved? Should old names inclusive of large numbers of species be used with subgenera (e.g., Bufo (Anaxyrus) americanus) to incorporate phyloge­ne­tic information without relinquishing scientific rigor and the historical connection to a vast lit­er­a­ture on ­these species? In this book, I do not attempt to ­settle the debate concerning anuran nomenclature; that would be an impossible task, considering the level of acrimony among

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Introduction xxv

distinguished professional biologists. No ­matter which nomenclature I use, someone ­will take offense. I generally follow anuran nomenclature as outlined on the web page Amphibian Species of the World (http://­research​ .­amnh​.­org​/­vz​/­herpetology​/­amphibia​/­index​.­php) and by Crother (2017, Scientific and Standard En­glish and French Names of American Amphibians and Reptiles of North Amer­i­ca North of Mexico, with Comments Regarding Confidence in our Understanding. Eighth Edition. Society for the Study of Amphibians and Reptiles, Herpetological Circular No. 43). I have included the subgenera (e.g., Aquarana, Novirana) recognized by Yuan et al. (2016) within the list of synonymies in the ranids, although some lineages ­were not given formal subgeneric names. I use Lithobates for our eastern, central, and southwestern ranid frogs, especially since Pyron and Wiens (2011) and Yuan et al. (2016) show a clear and impor­tant phyloge­ne­tic split between eastern/ central North American ranids and western North American / Asian ranids. Arguments in ­favor of lumping Rana and Lithobates are not particularly persuasive to me (although ­others disagree vehemently), especially considering the long recognition of the distinctive evolutionary history within ­these separate lineages, which is supported by morphological and molecular analyses. If nomenclature is to provide information and reflect evolutionary lineage, then this usage is informative and justified, as it is for Anaxyrus, Hyliola, and Dryophytes. Taxonomy is not static (Dominguez and Wheeler, 1997), regardless of ­whether names change. Just as biologists have adapted to changes in the nomenclature of many snakes and salamanders, so ­will they reconcile to well-­supported changes in anuran nomenclature. I continue to use Anaxyrus for the toads. The genus Dryophytes replaces Hyla for our North American species, and Hyliola replaces Pseudacris for the two western species (cadaverina, regilla) formerly placed in the genus Pseudacris in line with the revision by Duellman et al. (2016). En­glish common names follow Crother (2017), French common names follow Green (2012), and Hawaiian common names follow McKeown (1996). I include the scientific names in the popu­lar field guides of Powell et al. (2016) and McGinnis and Stebbins (2018) where differences occur, inasmuch as ­these guides are widely in use. I also provide a brief list of species synonyms in the scientific lit­er­a­ture. Full synonymies are published in the Cata­logue of American Amphibians and Reptiles published by the Society for the Study of Amphibians and Reptiles and online through the American Museum of Natu­ral History (see below). Some accounts contain additional nomenclatural notes, particularly when ­there is confusion in the lit­er­a­ture concerning species identification. The Amphibian Species of the World web page is particularly helpful in sorting through the often confusing nomenclatural histories of many taxa.

Dodd_Canada_int_5pgs_B1&B2.indd 25

Etymology. As noted above, most etymologies follow Beltz (2007). In a few cases, I have added additional information or amended Beltz’s accounts for clarification. Identification. The identification of most species of North American frogs is relatively straightforward, and the verbal descriptions and photo­graphs of postmetamorphs should prove sufficient for identification. However, many species have multiple color morphs and dorsal patterns that change among and even within individuals. Tadpoles also are highly variable in coloration and pattern depending upon growth stage, population, and region. Lighting and humidity influence the appearance of both adults and tadpoles of many species. In a few species, juveniles vary greatly from adults. Any typological description is likely to be insufficient to identify some individuals. Regional field guides may have multiple photo­graphs of some variants, but in this work it is impossible to include more than a hint of the color and pattern variation within frog populations. I have, however, pointed out some of the most common variants within the accounts. The identification of tadpoles is truly an art as much as a science. Many tadpoles look alike, especially when small, and even good diagnostic characters may only be apparent among the larger size classes. Researchers often rely on the features of the oral apparatus to recognize species (Fig. 17). In this book, however, I have not provided much information on labial teeth, tooth rows, and the position and number of oral papillae. Although impor­tant, ­these characters of tadpole identification have been covered in more detail by Altig and McDiarmid (2015). Nomenclature for tadpole external morphology follows Altig and McDiarmid (1999:35) (Fig. 18). Field experience and knowledge of natu­ral history are the best teachers when identifying larvae. A combination of verbal description, known range, breeding site, and season of observation ­will help in identifying tadpoles. Distribution. Species range is based on the latest field guides, atlases, and primary systematic lit­er­a­ture available as this work is completed. Readers should keep in mind that all range accounts and maps are approximate. Published and online maps in one state or province often do not agree with distributions in adjacent states or provinces, and even “authoritative” maps often disagree about species ranges. Another prob­lem is the blind ac­cep­tance of museum rec­ords based on online searches. If a museum database has not been completely updated taxonomically, an online search may give rather inaccurate results. For example, if the leopard frog species complex has not been updated, an online search might turn up Northern Leopard Frog locations where phenotypically similar species actually occur, particularly in the Southwestern United States. Lack of attention to verifying specimens likely has resulted in inaccurate maps

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xxvi  Introduction

Fig. 17.  Oral disk (mouthparts) of a tadpole. AL = anterior (upper) labium; A-1 and A-2 = first and second anterior (upper) tooth rows; A-2 GAP = medial gap in second anterior tooth row; LJ = lower jaw sheath; LP = lateral pro­cess of upper jaw sheath; M = mouth; MP = marginal papilla; OD = oral disk; PL = posterior (lower) labium; P-1, P-2, P-3 = first, second, and third posterior (lower) tooth rows; SM = submarginal papilla; UJ = upper jaw sheath. Terminology is that of Altig and McDiarmid (1999:35). Reprinted with permission of the University of Chicago Press.

and may explain some anomalies in historical versus pre­sent distribution patterns. Although maps may appear contiguous, frog populations are not evenly distributed within an area, especially since many species are found only in specific habitats that are themselves patchily distributed. Even in “ideal” habitats, colonization and extinction may change distribution patterns, even over a short time period; frogs may be pre­sent at a site one year but absent the next. More detailed information on distribution is contained in many regional field guides and online atlases, especially ­those that are maintained and georeferenced with dot distribution maps. I do not mention the field guides by Powell et al. (2016) and McGinnis and Stebbins (2018) as impor­tant references on distribution within each account; their inclusion should be considered automatic and unnecessary to repeat. Fossil Rec­ord. Most information on fossil frogs is based on Holman (2003). More detailed information and additional references are in Holman’s book. Systematics and Geographic Variation. Information on systematics comes from the latest primary peer-­reviewed

Dodd_Canada_int_5pgs_B1&B2.indd 26

Fig. 18.  Body morphology of a tadpole. TL = total length; BL = body length; TAL = tail length; TMH = the maximum height of the tail musculature; MTH = maximum height of the tail, including both tail fins and tail musculature; IND = distance between the narial apertures (internarial distance); IOD = distance between the eyes (the interorbital distance); TMW = maximum width of the tail. Terminology is that of Altig and McDiarmid (1999:26). Reprinted with permission of the University of Chicago Press.

lit­er­a­ture. Impor­tant geographic phenotypic variation is included, ­whether regionally based or as unusual morphs, such as the dif­fer­ent genet­ically based color patterns of Lithobates pipiens. Information on hybridization is included within this section. Adult Habitat. The adult habitat refers to the macroenvironment of the species, specifically the physiography and vegetative components of its ecosystem. Terrestrial and Aquatic Ecol­ogy. This section focuses on the microenvironmental components of a species’ life history. Information on daily and seasonal activity, movement patterns, juvenile dispersal, overwintering, physiological mechanisms employed to cope with harsh environmental conditions, orientation, and sensory perception is included within this heading. The title for this section may change depending upon the life history of the species ­under discussion (e.g., some species are entirely terrestrial, hence the section labeled Terrestrial Ecol­ogy). Calling Activity and Mate Se­lection. Seasonal and daily calling patterns, descriptions and specifications of calls,

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Introduction xxvii

environmental ­factors affecting calling, male calling be­hav­ ior, female response be­hav­ior, courtship, and intraspecific aggression with regard to calling are covered in this section. Breeding Sites. The physical and biotic components of breeding sites are covered, including the types of habitats used and physical and biotic characteristics of ­those habitats. Reproduction. This section includes information of reproductive activity, such as breeding season, environmental ­factors stimulating the onset of breeding, oviposition, clutch size, total fecundity, and time to and size at hatching. I include information from throughout the range of the species to illustrate the effects that latitude, elevation, and changes in weather patterns have on reproductive be­hav­ior. Despite this, ­little information is available on the variability of reproductive traits, and what is known is often based on small sample sizes from one or at best a few locations. This caveat applies to virtually all data on anuran life history from North Amer­i­ca and makes it difficult to assess the importance of differences among lit­er­a­ture reports. ­There is an urgent need for more extensive information over long time periods from multiple locations. Larval Ecol­ogy. All aspects of larval ecol­ogy are included within this section, such as the duration of the larval period, growth, ­factors influencing larval ecol­ogy and be­hav­ior, and the influence of predators on larval activity. I also include data on sizes of tadpoles and recent metamorphs. Some aspects of larval ecol­ogy may be included ­under the sections labeled Predation and Defense or Community Ecol­ogy, depending upon the nature of the interaction. Diet. Information on the diet of tadpoles, recent metamorphs, and adults is presented in this section. Most amphibians consume a wide array of invertebrates, particularly beetles. Even when information is not reported in the lit­er­a­ture, invertebrates prob­ably are the main prey. Predation and Defense. All amphibians are preyed upon opportunistically by a wide array of vertebrates (snakes, birds, mammals). Much information in the lit­er­a­ture, however, reflects opportunistic sightings rather than detailed analyses. Tadpoles face countless aquatic predators, such as turtles, snakes, and predaceous aquatic invertebrates. To avoid predation, larval and adult anurans have evolved a variety of defensive morphological, color-­related, behavioral, and biochemical responses. ­These are discussed in this section. Population Biology. Information on growth and demography is included ­here, with emphasis on population structure, age and size at maturity, sex ratios, recruitment, the effects of immigration and emigration, population ge­ne­tics, survivorship, and longevity. Community Ecol­ogy. This section focuses on interactions between anuran species within the aquatic and terrestrial

Dodd_Canada_int_5pgs_B1&B2.indd 27

habitat with specific emphasis on competition and habitat partitioning. It does not contain a list of other frogs within the area or same habitat type, nor does it attempt to list all potential biotic interactions. Indeed, so ­little is known about the community ecol­ogy of most anurans in North Amer­i­ca that this section is sometimes omitted from species accounts. Diseases, Parasites, and Malformations. As the heading implies, information is provided on anuran diseases, endo-­ and ectoparasites, and malformations. Malformations include ­those induced by unknown developmental ­causes as well as ­those thought to be induced by toxic substances or parasites. Malformations due to injuries are not reported. I have attempted to update parasite nomenclature, but it is likely I have inadvertently used synonyms on occasion. Susceptibility to Potential Stressors. Many amphibians are affected by substances (e.g., toxic chemicals) or environmental variables (e.g., acidity, conductivity) arising from both anthropogenic and natu­ral sources. ­These stressors can have both lethal and sublethal effects that differ among species and life stage. Some stressors act paradoxically, that is, small amounts may have more serious effects than large amounts. In this section, I report on potential stressors, as not all studies of stressors on anurans have been demonstrated to have effects on wild populations. The section includes subheadings on metals, chemicals (pesticides, industrial chemicals, drugs), nitrates and nitrites (e.g., fertilizers), pH (acidity), conductivity, alkalinity, UV radiation, and ­others as appropriate. I have made no attempt to standardize mea­sure­ ments; the mea­sure­ment units are ­those reported in the publication cited. Status and Conservation. This final section covers all aspects of the status and trends of species as understood based on published lit­er­a­ture. ­There is a ­great deal of information that is unpublished or available only in reports, files, and non-­peer-­reviewed Internet sites. In addition, the ­legal and regulatory status of species is constantly being revised and updated by federal, state, and provincial wildlife agencies. Each governmental organ­ization maintains an Internet site with up-­to-­date information on the protected status of species within their jurisdiction. Readers are advised to check on regulations prior to searching for frogs. Bibliography. The bibliography includes references from the 1690s to mid-2021. All references have been examined for accuracy of citation and content. Many authors apparently had not examined original copies of the papers they referenced, since I found many incorrect citations of titles, journals, books, dates, and page numbers as I read the more than 8,500 references included in the bibliography (also see Dodd, 2018, 2022). As it is, I made no attempt to cite ­every paper, note, thesis, or dissertation ever published on North American anurans; as my friend Ernie Liner once told me,

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xxviii  Introduction

“You ­can’t get them all!” I have tried to cite all pertinent papers, however, and of course I take responsibility for oversights. Readers are urged not to take my word for the brief summaries provided in the species accounts, but to verify information in which they are interested. Unfortunately, I could not list the authors for each species account for ­every edited state/regional guide (e.g., Jensen et al., 2008; Krysko et al., 2019) as it would increase space/word count substantially; readers should consult the books for individual species’ account authorship. Many obscure publications are available on the Internet, but by no means all. Just ­because information is not readily available does not mean it should be ignored or that it is not impor­tant or insightful.

Mea­sure­ments, Precision, and Generalizations Mea­sure­ments of body size, mass, distance, and area have been converted to metric units throughout the book as appropriate. Values for toxicological effects are largely given in LC50 values, and ­these generally are based on exposures for 24–96 hrs. Most amphibian biologists recognize that short-­term LC50s are grossly inadequate to assess the effects of toxic substances on anurans. Toxic substances may have both lethal and sublethal effects (on activity, feeding be­hav­ior, survivorship when faced by predators, body mass, larval period, postmetamorphic fitness) that cannot be mea­sured using traditional short-­term toxicological protocols. Toxics may affect eggs, hatchlings, larvae, and postmetamorphs differently and in paradoxical ways (lower concentrations may actually have more serious consequences than higher doses; eggs may be far less prone to toxic effects than late-­stage larvae). Results may vary between even closely related species, and the effects or lack of effects on one species cannot be extrapolated to another species. Clearly, toxicological studies need to be more rigorous. While combing through literally thousands of research papers, it became evident that authors frequently ­were imprecise in the terminology they used, a situation that has sometimes led to apparent contradictions in the lit­er­a­ture. In order to clarify and standardize information, I have attempted to interpret what the authors meant when confusion or contradiction seemed apparent. Below, I provide several examples of such confusion. As ­these examples demonstrate, precise meaning is impor­tant. •  Calling versus oviposition. Authors frequently confuse calling with breeding (oviposition). Many species call early and continue to call well ­after oviposition has ended. In fact, many species are opportunists with short breeding bouts over an extended period of months. Just ­because a species was heard calling does not mean oviposition occurred.

Dodd_Canada_int_5pgs_B1&B2.indd 28

•  Breeding season. Some authors assume breeding occurs continuously if rec­ords are available in spring and autumn (“I heard calls in March and October, therefore the breeding season extends March to October”). In fact, some species have a biphasic breeding cycle (spring and autumn), and the strength of the biphasic cycle may vary with latitude. For example, the breeding season may be biphasic in southern Arizona but monophasic during midsummer in northern Colorado. Researchers who work on single populations often fail to recognize and appreciate this distinction. •  Clutch size versus fecundity. Authors often confuse clutch size with fecundity when reporting egg counts. Clutches may be oviposited minutes, days, weeks, or even months apart. Total fecundity (total number of eggs deposited during a breeding season) may actually be a compilation of many bouts of egg deposition. Thus, some species appear to have widely varying “clutch sizes” when the authors actually ­were reporting on dif­fer­ent phenomena. Uncertainty about what constitutes a clutch occasionally influences conclusions about ­whether a species oviposits multiple clutches. •  Time to sexual maturity and larval duration. Authors often use dif­fer­ent methods of reporting ­these time-­based traits. For example, one author might say “the time to maturity is two years” whereas another might say “the time to maturity is three years,” yet both authors apparently mean the same ­thing. For example, a frog metamorphosing in Au­ ntil June 2013, which is gust 2011 might not reach maturity u a period of nearly two years, although the frog ­will be in its third year of life. Some authors report a two year larval duration, whereas ­others report three years in the same way. Authors often fail to make this distinction. Time confusion frequently leads to apparent contradictions in the lit­er­a­ture. In preparing this volume, I endeavored to determine where data originated, especially when recounted in general field guides. I quickly discovered that the basis for many reported life history traits rested on ­little empirical data; once a “fact” got into the lit­er­a­ture it tended to be repeated. Unfortunately, sample sizes ­were often very small, and data frequently originated from a single population or study. In one species, for example, the clutch size was repeatedly said to be 600–800 eggs in book ­after book. I soon discovered that the ­actual number was based on counts of 1 ovary from 2 individuals, then doubled to get counts of 684 and 760 (thus, 600–800). Such imprecision makes it difficult to understand life history evolution or build predictive models. Researchers need to understand the basis and limitations of data used to test hypotheses, or their meta-­analyses ­will not be of much value. Although much statistical analy­sis and theory is based on variance, many researchers still do not appreciate its importance and are quick to make generalizations. The

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Introduction xxix

validity of models and data interpretation depends on understanding variance. Life history traits of North American frogs are poorly known. Season, year, sex, climate, physiography, latitude, and elevation all contribute to the local population dynamics, status, and trends of North American frogs. Just ­because a species has a larval duration of 2–3 yrs in Ohio, for example, does not mean it does so in Louisiana or Québec (think of American Bullfrogs). All too often, biologists consciously or unconsciously embrace the concept of ecological typology whereby if a species does something in one location, it must do the same ­thing elsewhere. Such narrow thinking needs to be countered by comprehensive long-­term studies in a variety of locations and ­under varying environmental conditions. Amphibian biologists need to understand ­whether observed variance is based on sampling errors and biases, resource-­based variation, ge­ne­tics, or landscape differences. Unfortunately, such work is often tedious and consigned to “natu­ral history” that does not immediately lead to generalizations suitable for publication in prestigious journals (Ríos-­Saldaña et al., 2018). Researchers also should not be hasty to suggest inaccuracy in the work of ­others when results do not coincide. As the reader ­will soon learn when using this work, an initial impression of a ­great amount of information quickly fades to a realization about how ­little is known about the basic natu­ral history of North American frogs. Natu­ral history study has lost much of its following as a result of the impact of ge­ne­tics, population modeling, meta-­analysis, and a shift in focus to short-­term experimental studies with high potential for research funding. Regardless, natu­ral history data form the basis upon which hypotheses are conceived and conservation planned. I hope that this volume stimulates interest in the natu­ral history of anurans before many of ­these species dis­appear from our land-­and soundscapes.

cally the genera cannot be distinguished using external characters. I provide a key to the genera of North American frogs below (excluding Glandirana, which has been introduced into Hawai’i), but keys to species should be consulted in regional, state, or provincial guides. Another useful resource is the Key to the Herpetofauna of the Continental United States and Canada, Third Edition (Powell et al., 2019). Note that this key ­will not work outside the United States and Canada. Class Amphibia Skin may be smooth or warty; scales lacking; true claws not pre­sent. Order Anura Forelimbs smaller than hindlimbs and less robust; hindlimbs usually larger than forelimbs (for jumping and propelling the body if walking, hopping, or swimming); tails lacking in postmetamorphs (but note that male Ascaphus have an everted cloaca that superficially resembles a tail).

1a.

Body streamlined and fully aquatic; small head; lateral line system prominent; small upward-­pointing eyes; rear toes fully webbed; inner three toes have a black, keratinized sharp tip that superficially resembles a claw; tongue lacking���������������������������������������​Xenopus

1b.

Toes may or may not be webbed; no structure on toes that resembles a claw; tongue pre­sent������������������������ 2

2a.

Body squat (for walking or hopping rather than jumping); parotoid glands obvious; skin usually dry and warty (glandular) in appearance; eye pupil horizontal������������������������������������������������������������������ 3

2b.

Body squat or streamlined; parotoid glands may or may not be pre­sent; skin moist; eye pupil vertical or horizontal������������������������������������������������������������������ 6

3a.

Parotoid glands as long as head, tapering ­toward ​ the rear���������������������������������������������������������������������� 4

3b.

Parotoid glands shorter than the head and, if large, not tapering posteriorly�������������������������������������������� 5

4a.

Prominent cranial crests; very large parotoids; very large toad������������������������������������������������������� Rhinella

4b.

Low cranial crests; small toad; greenish������� Anaxyrus [only debilis, retiformis]

5a.

Thigh with 1–3 enlarged dorsal glands; deep valley between cranial crests��������������������������������������� ​Incilius

5b.

Thigh lacking enlarged dorsal glands or, if pre­sent, glands only slightly enlarged; valley between cranial

Key to the Genera of North American Frogs Dichotomous keys may be useful to determine the identity of frogs, but ­there is a ­great deal of phenotypic and geographic variation within certain species making keys less than perfect. A few species, particularly among the leopard frogs and hylids, may be impossible to identify without referring to much more detailed descriptions. For example, no ­simple descriptive definition can separate all individual Acris from Pseudacris accurately, and I can find no phenotypic description at all to separate Hyliola from Pseudacris (­these genera ­were separated based on molecular analyses). Likewise, Rana and Lithobates can be separated using internal morphology and molecular analy­sis, but phenotypi-

Dodd_Canada_int_5pgs_B1&B2.indd 29

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xxx  Introduction

​ naxyrus [all other crests lacking or only shallow�������� A species]

15a. Head very small and pointed in relation to the body������������������������������������������������������������������������ 16

6a.

Body squat (for hopping and digging); parotoid, if pre­sent, indistinct; skin moist and only slightly granular; eye pupil vertical; spade (dark sharp-­edged metatarsal tubercle) pre­sent�������������������������������������� ​7

15b. Head not very small in relation to the body; outermost toe thicker than ­others; male with “tail” (everted cloaca); streams of Pacific Northwest������������Ascaphus

6b.

Body streamlined; parotoid glands not pre­sent; skin moist and smooth, or only slightly rugose; spade (dark sharp-­edged metatarsal tubercle) absent�������������������� 8

7a.

Spade on hind foot elongated and sickle ­shaped���������� Scaphiopus

7b.

Spade on hind foot short, rounded, and wedge ­ shaped����������������������������������������������������������������� Spea

8a.

Toe pads pre­sent; toes with at least some webbing between�������������������������������������������������������������������� 9

8b.

Toe pads absent������������������������������������������������������ 14

9a.

Toe pads reduced, often barely wider than the toe; webs on feet significantly reduced �������������������������� 10

9b.

Toe pads obvious and wider than toes; webs on feet ca. ½ the length of the longest toe �������������������������� 12

10a. Small squat frog ( 1,981 m in elevation (Bury, 1968; Bosakowski, 1999). TERRESTRIAL AND AQUATIC ECOLOGY

The Coastal Tailed Frog is a species of the cool, moist, mature, and old-­growth forests of the Pacific Northwest. Adults frequently leave the ­water and forage terrestrially, especially along a stream bank. Most terrestrial activity

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Ascaphus truei 9

occurs in the autumn (September–­October), with ­little summer activity (Wahbe et al., 2004). Some juveniles and adults even wander overland (Bury and Corn, 1988a), with juveniles being found in clearcuts far more often than adults, who tend to remain in old-­growth forest (Wahbe et al., 2004; Matsuda and Richardson, 2005). Most individuals stay in proximity to streamside habitats, where they feed along creek banks and on the nearby forest floor to within 100 m of the stream. In the ­water, Coastal Tailed Frogs are found in shallow ­water (mean 4.1 cm in depth) ­under large rocks (mean cover area of 1,011 cm2) (Bury et al., 1991b), where they crawl rather than swim across the stream bottom. The thermal and ­water relations of this species are likely similar to A. montanus (Claussen, 1973a, 1973b), indicating a preference for cold mountain streams. Ascaphus truei prefers specific habitat conditions, although they may be found at lower densities at less favorable sites. In Washington, for example, Coastal Tailed Frogs are associated with higher elevations, moderate stream gradients, cobble substrates, substrates covered by 10% coarse woody debris, consolidated substrates, canopy covers >50%, and northern aspects (Bosakowski, 1999; Adams and Bury, 2002). The adjacent forest is often composed of Douglas fir (Pseudotsuga menziesii), red cedar (Thurja plicata), firs (Abies amabilis, A. procera), and western hemlock (Tsuga heterophylla) with mixed hardwoods such as red alder (Alnus rubra). Ascaphus truei are found at dif­fer­ent abundances in vari­ous stream segments and stream ­orders. For example, older life stages are most often found in higher elevation headwater streams than younger life stages. Higher-­order streams tend to have more evidence of reproduction than low-­order streams, and many first-­order streams may be too small to maintain Ascaphus populations. Ascaphus are also vulnerable to fish predation in lower-­order streams, and can be maintained only where ­there is sufficient flow for at least 600 m above where fish distribution stops (Hayes et al., 2006c). Higher-­order stream basins usually have more of such areas than do lower-­order stream basins. Large-­scale seasonal movements also are evident in some areas, as both larvae and adults move upstream in late summer from positions occupied early in the summer. For example, Hayes et al. (2006c) found that larvae moved a median distance of 733 m upstream by late summer, and adults moved 406 m upstream from portions of the stream they occupied ­earlier in the year. Hayes et al. (2006c) speculated that adults moved downstream before breeding activity and returned upstream immediately afterward. In contrast, Burkholder and Diller (2007) found that most individuals ­were site philopatric, with ­little upstream or downstream movement. Movements ­were usually 0–30 m

Dodd_Canada_int_5pgs_B1&B2.indd 9

within the stream channel, with adult females making longer movements than males and immatures of both sexes. Still, occasional adult movements of up to 110 m ­were noted both up-­and downstream. Coastal Tailed Frogs are likely to be monotonically photonegative in their phototactic response to white light, as is A. montanus, suggesting that they prefer to avoid daylight (Jaeger and Hailman, 1973). They likely show a U-­shaped spectral response and do not likely use color vision in phototaxis (Hailman and Jaeger, 1974). CALLING ACTIVITY AND MATE SE­L ECTION

Ascaphus truei does not have a mating call, as it would be useless over the roar of the cascades in which it lives. It is not known how the male and female locate each other, although they may move to a favored nest location where they are likely to come into contact. Visual cues are not used, but waterborne chemical cues may assist in locating mates or drawing them to a nest location (Asay et al., 2005). A male touches a female and holds onto her leg as he slowly works his way into position to grasp her thigh in inguinal amplexus. Axial and midbody attempts at amplexus are unsuccessful. During this time, the female is quiet with her legs extended; her thighs form a channel directed ­toward her vent. The male clasps his hands around the female’s body and then turns his “tail” inward and arches his back to insert the tip of the “tail” into the female’s cloaca. This occurs about 1.5 hrs following initial amplexus. During this time, the female keeps her nictitating membrane tightly closed as if in a trance and remains immobile. Amplexus lasts from 70 hrs to 7 days ­under laboratory conditions (Wernz, 1969; Brown, 1975c). ­After mating, the female dislodges the male and moves rapidly away, although the male may attempt to remain clasped. The mating sequence is described by Slater (1931), Noble and Putnam (1931), and Wernz (1969). BREEDING SITES

Oviposition usually occurs in shallow tributaries of mainstream channels. In Washington, females move into the tributaries in July for a late July oviposition. Mainstream channel ­waters are usually 4–10°C, with tributaries slightly warmer. However, eggs may be oviposited in both the main stream and tributaries. Eggs are placed ­under large flat rocks or boulders. REPRODUCTION

Mating occurs from spring to fall, with oviposition occurring in the summer ­after the spring snowmelt and before the winter rains. For example, Gaige (1920) found females with eggs and males in breeding condition from late June to early

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10  Ascaphidae

Adult female Ascaphus truei extruding eggs. Photo: Amber Palmeri-­Miles

September, although Wernz (1969) found gravid females in early April. Burkholder and Diller (2007) suggested most breeding in northern California occurs in spring with egg deposition from July to August. In western Washington, Palmeri-­Miles et al. (2010) found eggs at a communal site in late July, and Bury et al. (2001) found nests with eggs from late July to mid-­August. In the North Cascades, oviposition occurs from mid-­to late July (Brown, 1990). Noble and Putnam (1931) reported clasping pairs in June–­July, but Nussbaum et al. (1983) thought most reproduction takes place in the autumn. Karraker et al. (2006) reported oviposition from early June to late August. As previously noted, fertilization is internal. Eggs are oviposited in nests on the underside of stream boulders and cobbles, where they readily stick to the surface ­because of their sticky outer membrane (Gaige, 1920; Noble and Putnam, 1931; Brown, 1975c; Adams, 1993; Karraker and Beyersdorf, 1997; Bury et al., 2001; Palmeri-­Miles et al., 2010). Substrates ­under nest boulders may be sand, silt, or gravel. ­Under laboratory conditions, oviposition takes from 15 to 25 hrs with a fertilization rate of >90%. Nussbaum et al. (1983) reported that copulation requires 24–30 hrs. During oviposition, females aggregate with other females; Brown (1975c) reported 5–20 females ­under a single rock with developing embryos in early stages of cleavage. Nests may contain more than 1 clutch (Bury et al., 2001; Palmeri-­ Miles et al., 2010); ­these latter authors reported 4 females and 183 eggs beneath a single boulder. Bury et al. (2001) noted nests with 96 and 182 eggs. The clutch size of A. truei may be smaller than A. montanus (mean 41.9 vs. 66.6; Karraker et al., 2006), but females of both species may deposit eggs only ­every other year in some populations (Metter, 1967; Burkholder and Diller,

Dodd_Canada_int_5pgs_B1&B2.indd 10

2007). In contrast, Bury et al. (2001) suggested populations in the Olympic Peninsula deposited fewer eggs but on a yearly cycle. Clutch size is reported from 37 to 82 eggs in the Washington Cascades (Brown, 1975c). Adams (1993) reported a single clutch of 27 eggs in Oregon, and P.S. Corn (in Adams, 1993) found a clutch of 38 eggs. Karraker and Beyersdorf (1997) reported a single clutch of 28 eggs in California. In the Olympic Mountains, clutch size averaged 37 eggs (range 28–47) (Noble and Putnam, 1931) and 48 eggs (range 40–55) (Bury et al., 2001), whereas Gaige (1920) observed clutches of 35 and 49 eggs. Palmeri-­Miles et al. (2010) ­later reported clutch sizes of 70, 68, 47, and 24 eggs, also on the Olympic Peninsula. In Karraker et al.’s (2006) summary, clutch sizes ­were 15–182, but the larger clutches (>89) likely represented multiple clutches; most clutch sizes ­were 19°C, and only about 50% develop normally at temperatures 1,500 cm3/sec, low siltation, high gradients, and low ­water temperature (Welsh and Ollivier, 1998; Diller and Wallace, 1999), and have even been found in waterfalls (Gaige, 1920). They use large rocks for cover, with a mean area of 415 cm2, and they are found in deeper ­water but ­under smaller rocks than adults (Bury et al., 1991b). Hawkins et al. (1988) noted a preference for substrate cobble 10–30 cm in dia­meter, with highest larval densities in open regions just below headwater forests. In

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Ascaphus truei 11

Tadpole of Ascaphus truei. Photo: Brome McCreary

British Columbia, larval densities decrease with increasing levels of fine sediment, rubble, detritus, and wood; larval densities increase with bank width (Dupuis and Steventon, 1999). Cool ­waters are necessary for larval development. During the first year of life, laboratory observations suggest larvae prefer ­water 50% of the males visit dif­fer­ent breeding sites both within and between seasons, traveling as far as 230 m in a straight line between wetlands over a period of a few days (Ewert, 1969). Female movement between breeding sites is very rare within a season, as most females are amplexed rapidly at a breeding site and deposit all of their eggs. However, females may visit dif­fer­ent breeding sites between years (Ewert, 1969); 1 female moved 762 m between breeding sites in successive years. When displaced, toads usually orient ­toward the breeding site from which they ­were removed, instead of moving to the nearest breeding site (Oldham, 1966). In a study of toad ge­ne­tics, ­there ­were significant differences in mitochondrial DNA haplotypes between 5 breeding sites located at least 500 m apart. Significant differences between haplotypes ­were not recorded from one year to the next within a breeding pond, however, confirming site philopatry (Waldman et al., 1992). ­These results seem at odds with the movements recorded by Ewert (1969) and suggest the possibility of regional or population differences in ge­ne­tic structure and within-­and between-­season movement patterns. CALLING ACTIVITY AND MATE SE­L ECTION

Male American Toads attract mates both by calling and by active searching, which they alternate at wetland breeding sites (Fairchild, 1984; Wells and Taigen, 1984; ­Sullivan,

Calling male Anaxyrus americanus. Photo: David Dennis

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Anaxyrus americanus 23

1992a). Males vary considerably in the amount of time they spend calling, searching for females, and in clasping attempts. Aerobic capacities of American Toads are higher than ­those of other anurans (Taigen and Pough, 1981; Taigen et al., 1982), and allow this species to vigorously pursue mates during the very short reproductive period. However, aerobic capacity is not correlated with the time spent in any be­hav­ior, although individual males differ significantly in their aerobic capacity. Neither is aerobic capacity correlated with male body size. Thus, ­these physiological variables do not account for differences in the levels of male activity (Wells and Taigen, 1984). Calling occurs along the shoreline of breeding sites or in shallow ­water, and certain locations within a breeding site may be favored over other locations. For example, 85% of an Ohio chorus called from an area constituting only 30% of the available shoreline (Fairchild, 1984). Toads may switch calling sites, ­going from shoreline to ­water and back, for instance. Calls primarily serve as a species identifier and indicate the location of a breeding chorus. The call may also function to stimulate the final stages of oogenesis, and thus bring the sexes together during the optimum time for reproduction (Christein and Taylor, 1978). The call is a high-­pitched trill lasting 8–30 sec. Calling activity is short, usually occurring over a period of only a few days (Gatz, 1981a; ­Sullivan, 1992a; Howard and Young, 1998). Howard and Young (1998) classified 50.5% of the males at an Indiana breeding site as callers, and 33–64% of the males pre­sent ­were chorusing at any one time over 7 breeding seasons. In Ohio, calling activity could not be correlated with successful amplexus, as many noncalling males ­were also successful (Gatz, 1981a). Call characteristics vary among individuals and populations, and may or may not be correlated with morphology and environmental variables. For example, dominant call frequency (1400–1900 cps) decreases with body length (­Sullivan, 1992a) and mass, but only weakly with age. ­Water temperatures from 10–23°C have no effect on dominant call frequency (Howard and Young, 1998). On the other hand, pulse rates (normally 18–59 pulses/sec; Zweifel, 1968a) are weakly correlated with body length and mass, but not age. Pulse rates are, however, strongly correlated with ­water temperatures near a calling male, and with body temperature (Zweifel, 1968a). In discrimination tests, females did not discriminate between high and low call frequencies, but instead preferred high calling effort (duration x call rate) (­Sullivan, 1992a). The dominant frequency of the release call is also correlated negatively with male mass and body size (­Sullivan, 1992a). Regardless, males make the same release call ­whether they are amplexed by conspecifics or by artificial means. The same pulse rates and dominant frequen-

Dodd_Canada_int_5pgs_B1&B2.indd 23

cies are used for release calls regardless of the species of the amplexing intruder (Leary, 1999). Call duration varies annually (but not within a season; ­Sullivan, 1992a), and ­there is no correlation with male body mass, length, condition, or age. Call duration is weakly correlated with ­water temperature and negatively correlated with pulse rate, but it is unrelated to dominant call frequency (Howard and Young, 1998). Call durations lasted a mean of 7.8 sec (range 1.1–18.6 sec), with 1.6 calls per min in 1 Indiana population (Howard and Palmer, 1995), whereas in the northeast, call duration was 4–11 sec (Zweifel, 1968a). Mean and maximum call rates varied between 0 and 3.2 calls per min, but rates ­were not related to age, body length, or mass (Howard and Young, 1998). ­These authors noted that the social context of calling affected characteristics of the call, such as a reduction in dominant frequency when nearby males’ calls overlapped. In such circumstances, females tend to choose the male with the lowest dominant frequency. On the other hand, call duration was not affected by the presence of a nearby calling male, and females do not have preferences for certain dominant frequencies regardless of their age or past breeding experience (Howard and Palmer, 1995). They do, however, tend to prefer the leading caller when mixed choruses or paired males are pre­sent. Male American Toads have individual differences in mating calls that are evident at the population level, such that females may be able to discriminate relatives from non-­relatives at a breeding pond. Only 2 of 86 pairings ­were by toads genet­ically confirmed as relatives in ponds in Mas­sa­chu­setts, a ratio far below what would be expected if mating was completely random (Waldman et al., 1992). ­After adjusting for temperature and size, pulse duration, interpulse interval, pulse rise time, and call duration varied among breeding ponds (Waldman et al., 1992). Individuals that ­were genet­ically similar had similar call characteristics (except for dominant frequency), with genet­ically dissimilar individuals having much more varied call characteristics. ­These variables help ensure that despite breeding site philopatry, matings between relatives likely rarely take place. In areas where closely related members of the americanus group of toads come into contact and hybridize, call characteristics are intermediate between the parental species (Zweifel, 1968a; Cook, 1983). For example, A. woodhousii x A. americanus hybrids in Oklahoma call at 57 pulses per sec (versus 29–33 pulses per sec for A. americanus and no distinct pulses for A. woodhousii) and 5.3 sec in duration (versus 7.5–14.5 sec for A. americanus and 0.9–2.6 sec for A. woodhousii) at 20.5°C (Blair, 1956a). In Ontario at 16°C, A. americanus calls at a mean of 33 pulses per sec for a mean of 6.5 sec (or 216 pulses per call) compared with

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24  Bufonidae

A. fowleri (90 pulses/sec, 1.8 sec in duration, 157 pulses per call) and hybrids (49 pulses/sec, 4.2 sec call duration, 203 pulses/call) (Green, 1982). The intermediate nature of the calls of hybrids is likely due to the intermediate morphology of the laryngeal cartilages that affect call pulse (Green, 1982). Females move through a group of chorusing and non-­ chorusing males as they migrate to a breeding site. If not amplexed, they move ­toward a calling male and initiate clasping by touching him. ­Sullivan (1992a) noted that females always choose calls with higher rates or longer duration in discrimination tests. Licht (1976) proposed that size-­assortative mating occurs in A. americanus, but ­there is no evidence of this (Wilbur et al., 1978; Gatz, 1981a; Kruse, 1981a; Howard and Young, 1998). Instead, females generally select large males over smaller males, particularly while they are chorusing, indicating a degree of sexual se­lection by the female. It is not entirely clear why she does so, however, since neither male size nor arm length (for holding on during amplexus) is correlated with egg clutch size or mating success. Whereas ­there are heritable differences in the larval offspring among dif­fer­ent mated pairs in terms of survival to metamorphosis, mass at metamorphosis and timing of metamorphosis, ­these fitness variables are not correlated with size or age of the male. Thus, ­there appears to be no support for the hypothesis that adult size and age are correlated with larval ge­ne­tic superiority or mating success (Kruse, 1981a; Kalb and Zug, 1990; Howard et al., 1994). ­Either sex may initiate reproduction at the breeding site, including noncalling males. In Indiana, females initiated 43% of the 68 pairings observed by Howard and Young (1998), with males initiating 57%; an ­earlier study noted that females initiated reproduction in 31% of the encounters observed (Howard, 1988a). Prior breeding experience had no effect on the size of the males that females chose, and females did not choose larger males with successive pairings. Females may mate with the same male more than once (22% of observed pairings), and as many as 3 times (Howard and Young, 1998). As many as 70% of the females approaching a pond are intercepted by terrestrial males in Mary­land, and ­these are often smaller than calling males. Thus, small males may have mating opportunities despite the fact that their calls are not as attractive to females (Forester and Thompson, 1998). Many males are not successful. Gatz (1981a) estimated an average of 0.2 matings per male per breeding season in Ohio. Mating success may vary depending on the number of toads calling. Fairchild (1984) suggested that in large populations, calling males and noncalling males have an equal probability of finding a mate. In small populations (50 m from a forest edge may be insurmountable ­under adverse weather conditions (heat, direct sun, exposure to desiccating wind) during emigration. Open areas thus may form barriers to dispersal by juveniles, even though they do not do so for adults. In keeping with ­these results, Gravel et al. (2012) reported that the abundance of dispersing American Toad juveniles increased with ­percent canopy cover in New Brunswick. Adult American Toads immigrate and emigrate to and from ponds non-­randomly, but the patterns may change from one year to the next (Homan et al., 2010). Adult dispersal occurs diurnally, especially in areas where nighttime

Dodd_Canada_int_5pgs_B1&B2.indd 28

temperatures drop below freezing, or nocturnally. Toads may use riparian corridors to move among habitats, although adult toads do not prefer to travel through inundated swamps and wetlands (Burbrink et al., 1998). Rainfall is not required to initiate adult dispersal. In a study of 16 female A. americanus tracked by radiotelemetry, Mary­land toads dispersed from 250–1,000 m from the breeding pond, and most traveled >400 m. Dispersal was non-­random and linear, with toads interrupting travel with sedentary periods (Forester et al., 2006). Most long-­distance dispersal took place within 10 days of leaving the breeding pond, with 1 toad moving 610 m in a single day. Movements usually begin within a week ­after chorusing ends. In Minnesota, Ewert (1969) recorded dispersal taking place over 109 days one year and for >73 days the next. One toad took 22 days to move 701 m and was still moving away from its breeding site when last observed. Another toad traveled 1,005 m to its overwintering site, and still another took 68 days to reach its final overwintering location. Movements of >200 m per 24 hr period ­were common in Ewert’s population; most movements ­were ca. 26 m per 24 hr period, however, as toads moved to overwintering sites. Summer movements ­were much the same as in Forester et al.’s (2006) Mary­land population. DIET

Larvae begin feeding at Gosner (1960) stage 25. Tadpoles eat filamentous algae, blue-­green algae, periphyton, diatoms, soft tissues from vascular plants, eggs of crustaceans, carrion (including other tadpoles), and fecal material (Munz, 1920). Larvae also filter detritus suspended in the ­water column (Test and McCann, 1976), and detritus is an impor­tant component of their diet. In fact, the ingestion of phytoplankton may have more to do with an open canopy cover (leading to greater availability) than preference, as tadpoles seem to prefer feeding on leaf detritus (Earl and Semlitsch, 2012). Feeding ceases about Gosner stage 42. In experimental ­trials, American Toad tadpoles chose foods higher in calcium than in controls, which suggests a degree of selectivity in tadpole food choice (Botch et al., 2007). Juveniles eat a wide variety of small invertebrates (Hamilton, 1930). For example, the diet of recently metamorphosed American Toads consisted primarily of springtails (Collembola), mites (Acarina), parasitoid wasps and ants (proctotrupoid and formicid Hymenoptera), and beetles (staphylinids and carabids) in a study in Québec (LeClair and Vallières, 1981). Other prey included spiders, larval lepidoptera, larval and adult flies, and vari­ous other insects (Hemiptera, Homoptera, Neuroptera, Trichoptera, Thysanoptera). American Toads are both active and sit-­and-­wait foragers. Adults eat a ­great variety and quantity of invertebrates,

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Anaxyrus americanus 29

including ants, beetles, grasshoppers, isopods, millipedes, spiders, gypsy moths, tent caterpillars, diptera larvae, snails, slugs, butterflies, moths, wasps, and virtually any small animal that can be captured with its protractible tongue (Kirkland, 1897, 1904; Garman, 1901; Dickerson, 1906; Miller, 1909a, 1909b; Hine, 1911; Surface, 1913; Smith and Bragg, 1949; Larochelle, 1974; Gilhen, 1984; Klemens, 1993; Bellocq et al., 2000; Gibson and Ivanov, 2020). ­There is no relationship between dietary niche breadth and among-­individual diet variation in this species (Cloyed and Eason, 2017). Their large appetite makes them valuable in controlling agricultural pests. Kirkland (1897; repeated by Dickerson, 1906, and Patch, 1918), reported that a single toad could eat 9,936 injurious insects and 368 beneficial insects in a single season, making its economic value $19.88 in 1897 dollars! Miller (1909a) disputed that claim somewhat, recalculating the yearly value of a toad at $5.00. In contrast, some invertebrates, such as bombardier beetles, are effective at thwarting ingestion due to their chemical defenses (Dean, 1980). PREDATION AND DEFENSE

Chief among the antipredator defenses of American Toads are the granular glands that secrete poison when mechanically stimulated, as when the toad ­faces a severe threat from a large predator. During normal ­handling, a toad does not secrete poison. The granular glands are concentrated dorsally as somewhat elliptical or oblong parotoids ­behind the eyes, and in the warty protuberances of the skin. Scatterings of granular gland occur elsewhere, although they are not prominent. Juveniles also have ­these glands, and when handled by invertebrate predators, juveniles become immobile and are often released as the predator is exposed to the skin secretion (Brodie et al., 1978). Details of glandular structure of the American Toad are in Muhse (1909). American Toads, both larvae (Kats et al., 1988) and postmetamorphs, are toxic or noxious to many potential predators. Interestingly, recently hatched larvae and tadpoles approaching metamorphic climax are unpalatable to both vertebrate and invertebrate predators, but intermediate stage larvae are not (Brodie et al., 1978; Brodie and Formanowicz, 1987). The toxicity serves to discourage predation, rather than to deliver a lethal toxin to the predator. Toxin is also deposited in the eggs via the female’s blood. They may be eaten by certain salamander larvae, but they are often spit out ­after initial contact, whereupon the larva, rather than the jelly capsule, is consumed. Eggs are eaten by the ­Giant Apple Snail (Pomacea maculata) ­under experimental conditions (Car­ter et al., 2018), but not by mosquitofish (Zeiber et al., 2008).

Dodd_Canada_int_5pgs_B1&B2.indd 29

Larvae are also unpalatable to many potential predators, but the degree of unpalatability may depend upon the predator (Voris and Bacon, 1966; Walters, 1975; Kruse and Stone, 1984; Holomuzki, 1995; Smith et al., 1999; Zeiber et al., 2008) as well as the larval stage of development. Paradoxically, larvae may be more successful in certain ponds with fish predators than in ponds where fish are removed. Fish depress the abundance of other tadpole predators, and when predaceous fish are removed, increases in the level of invertebrate predation result in decreased larval American Toad survival (Walston and Mullin, 2007b). American Toad larvae approaching metamorphosis have an immobility response whose duration is inversely correlated with stage of development (Dodd and Cupp, 1978). When disturbed, they draw their limbs into the body and refrain from any movement. However, just prior to metamorphosis, the immobility response is very short, as all attention is directed ­toward metamorphosis. Recent metamorphs, however, again adopt an immobile response which allows concealment from certain predators. The mosquito Culex territans feeds on American Toads and even can use the toad’s call to locate its next blood meal (Bartlett-­Healy et al., 2008). Other predators include Chipping Sparrows (Spizella passerina) on recent metamorphs (Gorham, 1964), and Largemouth Bass (Micropterus salmoides), Eastern Hognose Snakes (Heterodon platirhinos), Eastern Garter Snakes (Thamnophis sirtalis), Northern Watersnakes (Nerodia sipedon), Blotched Watersnakes (N. erythrogaster), Queensnakes (Regina septemvittata), Snapping Turtles (Chelydra serpentina), hawks, grackles, geese, chickens, guinea fowl, owls, herons and waterfowl, skunks (Mephitis mephitis), and raccoons (Procyon lotor) on adults (Miller, 1909b; Evermann and Clark, 1916; Burt, 1935; Oldham, 1966; Schaaf and Garton, 1970; Forester et al., 2006; Palis and Mendenhall, 2021). Heterodon in par­tic­u­lar is a toad specialist, containing specialized dentition for manipulating and deflating toads. Several incidents regarding mass predation on breeding toads have been reported, with ­either raccoons or striped skunks (Mephitis mephitis) implicated as responsible (Groves, 1980). Predators of larvae include leeches (Desserobdella picta), odonate larvae (Anax, Libellula, Leucorrhinia, Sympetrum, Eurythemis, Plathemis), backswimmers (notonectids), predacious ­water bugs and diving beetles (Belostoma; Dytiscus), waterscorpions (Ranatra), other frog larvae (Lithobates), salamanders (Ambystoma opacum, Notophthalmus viridescens), and Least Sandpipers (Brockelman, 1969; Walters, 1975; Stangel, 1983; DeBlieux and Hoverman, 2019). American Toads, which breed 4–10 weeks ­after Wood Frogs in North Carolina, avoid breeding ponds inhabited by Wood Frog larvae (Petranka et al., 1994). Not

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30  Bufonidae

surprisingly, Wood Frog tadpoles totally consumed eggs and hatchlings of the toad very quickly in a series of natu­ral and mesocosm experiments, and none survived (Petranka et al., 1994). Larval American Toads respond to the potentially debilitating cercariae of Echinostoma trematodes by alternating periods of swimming quickly with extremely rapid twisting, turning, and tumbling movements (Taylor et al., 2004). ­These movements have been described as “massive, violent and multiplanar” (Taylor et al., 2004). Such rapid movements last 4–8 sec. Presumably ­these rapid and vigorous movements help to dislodge cercariae crawling on the skins of tadpoles and lodging in places where they could cause limb malformations. Taylor et al. (2004) observed cercariae attempting to enter the spiracle of toad larvae, where buccal pumping rates are not as high as they are in other species (Wassersug and Hoff, 1979). The fact that toad larvae are distasteful prob­ably allows them to move more vigorously (and thus conspicuously) in their attempts to dislodge ­these parasites, more so than a less palatable species might do. In the presence of potential predators, ­there are conflicting reports on the be­hav­ior of American Toad larvae. American Toad larvae do not modify their activity levels or swimming be­hav­ior in experimental ­trials where the olfactory cues of fish, odonate larvae, or combinations thereof ­were pre­sent, but the predators themselves ­were not in direct physical proximity (Richardson, 2001; Smith et al., 2009). Likewise, toad larvae did not reduce the amount of time swimming in the presence of nonindigenous mosquito fish (Gambusia affinis) or native bluegill (Lepomis macrochirus), nor did they spend more time in vegetation than exposed on gravel substrates (Smith et al., 2008; Smith and Awan, 2009). In contrast, Holomuzki (1995) reported reduced larval activity in the presence of fish, and Relyea (2001b) found that toad larvae reduced their level of activity in the presence of both fish (Umbra) and dragonfly larvae (Anax), but not in the presence of newts (Notophthalmus). In addition, morphological changes took place in response to the predator’s presence. Tadpoles developed shallower and longer tails in the presence of Umbra, and shallower tails with Anax, than they did with newts and predaceous beetles (Dytiscus) (Relyea, 2001b). Presumably, this phenotypic plasticity in response to predator presence has some adaptive component, although most of ­these predators tend to avoid Anaxyrus tadpoles. Likewise, American Toad larvae decrease activity levels in the presence of Anax predators, as well as when food availability is increased. Activity levels of larvae exposed to vari­ous types of predators do not change with predator density or satiation in laboratory ­trials, or with larval body size despite differences in noxiousness (Anholt et al., 1996). In contrast, activity levels increase in the

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presence of ­water beetles (Dysticus) (Smith and Awan, 2009). Body size further affects the activity of larvae in the presence of predators; small tadpoles are more active and use open ­water more often than large tadpoles. Chemoreception plays an impor­tant role in larval defense. Larvae are able to recognize some predators innately using chemoreception. American Toad larvae, both wild-­caught and laboratory reared, decrease activity and increase aggregation in the presence of water-­borne cues emanating from fish (Lepomis macrochirus) and odonate (Anax) predators. This response does not require previous experience with the predators, as both naïve and experienced larvae exhibit the same be­hav­ior. In contrast, no such be­hav­iors are observed in response to chemical cues from the newt Notophthalmus viridescens (Gallie et al., 2001). In addition, larvae are able to recognize cues released from injured conspecifics, and ­will avoid an area where injured tadpoles occur (Petranka, 1989). In ­trials featuring beetles (Dytiscus), fish (Umbra), newts (Notophthalmus), and dragonfly larvae (Anax), larval toads had low levels of predation. Toads ­were easily captured by Umbra and Anax, but ­were immediately rejected; however, the most common response by ­these predators was to ignore the tadpoles. Newts also found toad larvae unpalatable, whereas Dysticus readily captured and ate toad tadpoles in about half their attempts (Relyea, 2001a). The beetles also took a long time to consume the tadpoles they ate (mean 17.2 sec). Adults make use of the following be­hav­iors when confronted by a predator: fleeing rapidly, remaining immobile, taking a crouching stance, tucking the chin downward ­toward the pectoral region, body inflation, digging, hiding, kicking, and backing away (Marchisin and Anderson, 1978; Hayes, 1989; Heinen, 1994; 1995; Hartzell, 2016). Body postures and inflation presumably make the toad appear larger or more difficult to ­handle. Postures may be oriented ­toward an approaching predator, perhaps to direct the dorsum containing the parotoid glands and skin secretions to the face (and eyes) of the predator. By remaining immobile, both adult and recently metamorphosed juveniles may be able to avoid detection, especially when the potential predator is visually oriented. American Toads also exhibit an ability to blend into surrounding litter by slowly changing their background coloration. POPULATION BIOLOGY

Newly metamorphosed American Toads are 10 mm and weigh approximately 0.05 g in New York (Miller, 1909a; Hamilton, 1934; Raney and Lachner, 1947; Pough and Kamel, 1984). They are 12–15 mm in Indiana (Minton, 2001), but only 6–8 mm in Louisiana (Dundee and Rossman,

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Anaxyrus americanus 31

Adult Anaxyrus americanus charlesmithi. Photo: David Dennis

1989). By first overwintering ­after metamorphosis, they are approximately 20 mm SUL, and grow to 60 mm SUL in their first full season. They add approximately 18 mm during the second full season, with negligible growth thereafter (Hamilton, 1934). Of course, ­these growth rates ­will vary geo­graph­i­cally depending on the extent of seasonal activity, but the pattern of initial rapid growth followed by decreasing growth to sexual maturity should be consistent among populations. In a series of controlled outdoor mesocosm experiments ­under semi-­natural conditions, juvenile American Toad survivorship varied annually and by season. Survivorship was lower from metamorphosis to September (mean 21.1%) compared to a mean of 42.4% from September to the following May (Earl and Semlitsch, 2015). Growth rates also varied from metamorphosis to September (12.4 mg/d) versus 5.6 mg/d from September to May. Both microclimate (temperature and soil moisture) and microhabitats (logs, canopy cover, leaf litter depth) ­were impor­tant for juvenile growth and survivorship, but not the type of forestry practices (Earl and Semlitsch, 2015). Growth rates tended to increase with increasing temperature and decrease with soil moisture content from metamorphosis to September. From September to May, growth rates increased with leaf litter depth. The best predictor of metamorph survival was the quality of the terrestrial environment (Earl and Semlitsch, 2013). Earl and Semlitsch (2013) also found no support for life history carry-­over effects from the larval environment to the postmetamorphic terrestrial environment. Toads tend to grow slowly, at least in upstate New York. Small males (63–72 mm SUL) grew at an annual rate of 4.8 mm/yr, 73–76 mm males grew 4.0 mm/yr, 77–81 mm males grew 2.9 mm/yr, and 82–94 mm males grew 0.1 mm/ yr (Raney and Lachner, 1947). ­These authors suggested that

Dodd_Canada_int_5pgs_B1&B2.indd 31

male toads in this region reached their maximum length in 3 or more years, but they did not have comparable data for females. ­There may be some confusion in the lit­er­a­ture concerning age at maturity, since a toad reaching maturity in one season may not actually breed ­until the following season. Most reports of American Toads in breeding choruses are 2–4 yrs in age, with a few individuals reaching 5 yrs (Acker et al., 1986) or older (Jennette et al., 2019). In Pennsylvania (Meshaka et al., 2017a, 2020a), males reach sexual maturity at 2 yrs (minimum size 55–56.7 mm SUL), whereas females reach maturity at 3 yrs (minimum size 68.9 mm SUL). In Indiana, males first breed when they are 2 yrs of age, and females at 3 yrs. Males in an active chorus ranged from 2 to 6 yrs of age, and ­were 49–70 mm SUL (mass 14–48 g) (Howard and Young, 1998). Indiana females varied between 3 and 6 yrs in age, with older females generally larger than younger females. Body length is positively correlated with both age and mass. The growth rates reported by Hamilton (1934) suggest a similar age at first breeding in upstate New York. In ­Virginia, skeletochronology revealed most of the breeding population consisted of males at 3–4 yrs, and females at 4–5 yrs (Kalb and Zug, 1990); age was not correlated with size in this southern population. In Mary­ land, most toads ­were 3–5 yrs, with a few reaching 6–7 yrs (Jennette et al., 2019). Males sometimes ­were found at breeding ponds at 2 yrs of age, but very few. Females ­were still common in urban populations at 5 yrs. Toads surviving in urban and suburban settings may have dif­fer­ent demographics and life history characteristics from toads living in rural or more natu­ral settings. Jennette et al. (2019) found that American Toad males from rural areas ­were 14.6% larger than males from urban populations, and females ­were 12.7% larger than ­those from urban settings. Males from rural areas ­were larger than ­those from suburban populations. Males from rural areas ­were also 3.9–8.4% heavier in body mass than ­those from urban populations. Paradoxically perhaps, females from urban areas produced larger clutch sizes than rural toads, and the dry weight of clutches also was larger in urban populations. Age structure also varied, with urban toads slightly older in general than rural and suburban toads. COMMUNITY ECOLOGY

Both predators and interspecific competitors may have significant effects on larval size, the length of the larval period, and survivorship, depending on the densities of the vari­ous members of the community (Wilbur and Fauth, 1990; Smith and Dibble, 2012). In experimental ­trials, the growth rates of American Toads are negatively affected by the density of other larvae (Lithobates palustris, L. sphenocephalus,

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32  Bufonidae

Scaphiopus holbrookii), including conspecifics (Wilbur, 1977b; Alford and Wilbur, 1985; Alford, 1989a, 1989b, but see Wilbur, 1987), or even the density of snails (Holomuzki and Hemphill, 1996). In turn, the density of American Toad tadpoles has a negative effect on the density of Dryophytes chrysoscelis tadpoles (Alford, 1989a), and on snail reproduction (Holomuzki and Hemphill, 1996), but no effect on L. palustris, regardless of when tadpoles enter the pond (Alford, 1989b). Lithobates sphenocephalus, in par­tic­u­lar, significantly affects both the abundance and biomass of Anaxyrus tadpoles, but the effects vary depending on which species bred first in the pond (Alford and Wilbur, 1985); when this species is not pre­sent or eliminated from experimental mesocosms by predation, the mass and survivorship of Anaxyrus increases. By day 51 of development, however, American Toad tadpoles are of a sufficient size that even the presence of the newt Notophthalmus viridescens has no effect on them. For tadpoles surviving to this stage, metamorphosis occurs at greater mass than if newts had not been pre­sent to reduce initial densities of tadpoles. Even when D. chrysoscelis tadpoles ­were reduced in number, the biomass of American Toads increased, suggesting a certain degree of competition between ­these species (Alford, 1989a). As might be expected, the effects of competition are more pronounced at higher tadpole densities. Wilbur (1987) achieved similar results to the ­later study by Alford (1989a) regarding interspecific competition, although he also superimposed an experimental situation in which mesocosms ­were drawn down simulating pond desiccation. At low densities, most American Toad tadpoles completed metamorphosis, but at high densities, competition between the toads and L. sphenocephalus larvae prevented successful metamorphosis. The addition of newts mediated ­these results. Although fewer (28%) American Toads survived at low densities in the presence of N. viridescens as ­water was drawn down, 44% actually survived at higher densities as newt predation removed toad tadpoles during the desiccation pro­cess (Wilbur, 1987). ­These complementary studies illustrate how density, competition, community composition, predation, and environmental stochasticity interact in larval survival. Other taxa that compete with American Toad tadpoles in similar habitats, such as snails (Physella integra) in small streamside pools, tend not to co-­occur within even spatially proximate pools if tadpoles are pre­sent (Holomuzki and Hemphill, 1996). In this case, both compete for benthic algae as food, and their presence or absence affects the species composition of algae within the shallow pools. By avoiding each other, the snails and tadpoles eliminate the potential for competition. ­These results are in contrast to Smith et al.

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(2012) who found a more complex relationship between snails and tadpoles. When tadpoles ­were pre­sent, especially at high densities, they reduced the amount of periphyton pre­sent for the snails and thus decreased snail abundance (Smith et al., 2012); at low densities, the tadpoles had no appreciable effect on snail abundance. The presence of snails had no effect on survivorship or mass of A. americanus tadpoles in experimental ­trials, but ­there was a negative effect on the number of days to metamorphosis. If tadpoles ­were eliminated—or ­after tadpoles metamorphosed—­snail abundance again increased as more food became available. Note that Smith et al. (2012) used lower snail densities in their experiments than Holomuzki and Hemphill (1996), thus perhaps decreasing the competitive effect. Larval toads tend to metamorphose at smaller sizes in the presence of a predator, such as dragonfly larvae (Anax), newts (Notophthalmus), or mosquitofish (Gambusia), as well as when food is scarce, when compared to situations where predators are absent and food is adequate (e.g., Smith and Dibble, 2012). In experimental ­trials, the presence of dragonfly larvae did not have an effect on the larval period, but food scarcity significantly increased the duration of the larval period. The presence of potential predators affects larval be­hav­ior, as larvae tend to decrease activity and segregate themselves away from contact with the predator. As a result, larvae in the presence of a predator ­will have a decreased growth rate. The metamorphic response of tadpoles is mediated through behavioral effects on growth, which in turn affects size at metamorphosis (Brockelman, 1969; Wilbur, 1987; Skelly and Werner, 1990). Larvae also may metamorphose ­earlier (Wilbur and Fauth, 1990) or ­later (Smith and Dibble, 2012) in the presence of an aquatic predator, suggesting a trade-­off between ­future fitness (i.e., larger size at metamorphosis) and current survivorship. Habitat complexity may affect vari­ous life history aspects of larval development. In contrast to D. versicolor, for example, complex habitat structure increased time to metamorphosis, decreased mass at metamorphosis, and increased survivorship among treatments of A. americanus tadpoles, an effect that was compounded by the addition of L. pipiens tadpoles to experimental enclosures (Purrenhage and Boone, 2009). With Lithobates, mass at metamorphosis decreased at high densities compared with low competitor densities, regardless of the amount of time to metamorphosis. Survivorship also increased in the presence of Lithobates competitors in complex habitat structures, especially at low competitor densities. Wetland vegetation may have profound effects on tadpole development, and recent experiments suggest that nonindigenous species may alter the survival, developmental rates, and diet of American Toad tadpoles (Maerz et al., 2005b;

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Anaxyrus americanus 33

Brown et al., 2006). In both laboratory and field experiments, tadpoles developed more slowly in the presence of both ­water extracts and vegetative debris composed of Eurasian purple loosestrife (Lythrum salicaria) compared with native cattails (Typha latifolia). Survival and developmental rates ­were more variable in the presence of loosestrife than cattails, although survival was not affected in mesocosms and field enclosures. Dif­fer­ent plant species result in dif­fer­ent algal communities, and thus food quality and quantity available to tadpoles; this in turn affects per­for­ mance (Brown et al., 2006). Purple loosestrife did this by reducing the nutrients available for algae. Maerz et al. (2005b) and Brown et al. (2006) further suggested that Lythrum caused toxicity due to its high tannin concentrations, perhaps due to damage of the gills. In another example of inhibitory effects by plants, larval American Toads failed to develop through metamorphosis when exposed to a species of the green alga Spirogyra. Exposure to this alga not only inhibits development in natu­ral populations, but extracts from the alga, built up through time, can be acutely toxic to developing larvae (Wylie et al., 2009). In addition to larval effects, recently metamorphosed American Toads may experience competition from other recently metamorphosed species. Recently metamorphosed Green Frogs (L. clamitans) do not eat recently metamorphosed A. americanus, but when reared together, American Toads have a smaller body mass and lower survivorship when in proximity to Green Frogs than when they are reared separately; activity levels also are reduced. Sams and Boone (2010) suggested that interspecific competition between recently metamorphosed anurans may occur, but that in nature such effects are ameliorated by spatial segregation in terrestrial habitats. In addition to other frogs, other predators may impact populations of recently metamorphosed American Toads. When habitats are invaded by the nonnative plant Japa­nese stilt grass (Microstegium vimineum), toadlet survivorship decreases despite favorable microclimatic conditions. It turns out that spider populations increase in the stilt grass, leading to a decrease in prey available to the metamorphosed toadlets. However, DeVore and Maerz (2014) demonstrated that despite prey reductions, it was the increased spider per­sis­tence that decreased toadlet survival in the invaded habitats. Invasion of the grass affected the trophic interactions of both toadlet and spider populations, even though toadlet growth was not affected. Toadlets grew at a mean rate of 16.53 mg/d. DISEASES, PARASITES, AND MALFORMATIONS

Xanthoma cancer has been reported from A. americanus (Counts and Taylor, 1977). Pathogens include viruses,

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bacteria, and fungi. ­Water molds (Saprolegnia) form on eggs, especially unfertilized eggs (Miller, 1909b), and tadpoles (Bragg, 1962a). The fungi Basidiobolus ranarum and Dermosporidium penneri have been recorded from American Toads in the Midwest (Nickerson and Hutchison, 1971; Jay and Pohley, 1981), and dermosporidiosis is suspected in toads from ­Virginia (Green et al., 2002). Batrachochytrium dendrobatidis (Bd) has been found on A. americanus (14.6%) from Maine where it occurs on the toe-­webbing and the skin of the pelvis (Longcore et al., 2007). Bd was reported in toads from Québec (4.3%), spanning a period between 1960 and 2001 (Ouellet et al., 2005a), and from Minnesota (Martinez Rodriguez et al., 2009; Wolff et al., 2012). Other rec­ords are from Connecticut, Illinois, Mary­land, Ohio, Oklahoma, North Carolina, and ­Virginia (Davidson and Chambers, 2011b; Krynak et al., 2012; Muelleman and Montgomery, 2013; Hughey et al., 2014; Phillips et al., 2014; Watters et al., 2016; Marhanka et al., 2017; Tupper et al., 2017; Fuchs et al., 2018; Watters et al., 2018, 2019; Lentz et al., 2021). When exposed to Bd and low food availability, survivorship of American Toad metamorphs decreases. Based on ­these results and by using a population model, Rumschlag and Boone (2020a) estimated that Bd would reduce population growth by 14% when food availability was high, and by 21% ­under conditions of low food availability. Based on experimental ­trials, this species should be very prone to mortality via Bd infection (Gahl et al., 2011; Ortiz-­Santaliestra et al., 2013; Wise et al., 2014). Newly metamorphosed toadlets are more susceptible to infection than 4 week old juveniles, and can experience up to 75% mortality in 2 weeks ­under laboratory conditions (Wise et al., 2014). In nature, Bd was more prevalent on adults (28%) than metamorphs (1.6%) in Ohio, suggesting that infections occur primarily post-­metamorphosis (Rumschlag and Boone, 2020b). Adult infection was reduced with increasingly open-­canopied habitats across spatial scales of 100–1,000 m, but increased with the proportion of forested habitats over small (100 m) spatial scales (Rumschlag and Boone, 2020b). Toads infected by Bd display behavioral fever—­they increase their body temperature through behavioral thermoregulation (Karavlan and Venesky, 2016). Toads with higher Bd infection intensities had higher body temperatures than toads with lower infection intensities. Presumably raising the body temperature helps combat the infection, as Bd does not survive temperatures >30°C. In experimental ­trials, Bd had no interactive effects with sublethal pesticide mixtures (involving atrazine, acetochlor, glyphosate, 2,4-­D, carbaryl, chlorpyrifos, endosulfan, permethrin) on survivorship or on infection loads, but Bd reduced larval survivorship even in the absence of the pesticide mixture (Jones et al., 2017).

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34  Bufonidae

The deadly pathogen ranavirus occurs in A. americanus (Wolf et al., 1968; Homan et al., 2013; Watters et al., 2018; Lentz et al., 2021). American Toads are sensitive to ranavirus, but do not appear to be affected much by FV3 (Hoverman et al., 2011). Ranavirus infection does not increase the vulnerability of tadpoles to attack by dragonfly larvae (Anax) or ­giant ­water bugs (Belostoma) (DeBlieux and Hoverman, 2019). The bacterium Aeromonas hydrophila was reported from a wild population (Dusi, 1949). Another bacterium found on this species’ eggs and juveniles is Janthinobacterium lividum. It produces a potent antifungal metabolite that might contribute to an animal’s immune response to Bd (Standish et al., 2018). American Toads are parasitized by a wide variety of protozoans and protozoan-­like organisms, including Hepatozoon climate (Kim et al., 1998), Opalina obtrigonoidea (Odlaug, 1954; Delvinquier and Desser, 1996), Myxidium (McAllister and Trauth, 1995), and Toxoplasma (Stone and Manwell, 1969). As noted by Green (2005), blood parasitism is common, especially involving Trypanosoma fallisi. The species is parasitized by the Myxosporean protist Cystodiscus sp. (McAllister et al., 2014b). Nematodes include Gyrinicola batrachiensis on tadpoles (Adamson, 1981a, 1981b, 1981c) and Cosmocercoides sp., C. dukae, C. variabilis, Oswaldocruzia leidyi, O. pipiens, Physaloptera sp., Physaloptera ranae, Rhabdias sp., R. americanus, R. bufonis, and R. ranae on adults (Baker, 1977, 1978a, 1978b, 1979a; Ashton and Rabalais, 1978; Vanderburgh and Anderson, 1987a, 1987b; Joy and Bunten, 1997; Dyer, 1991; Yoder and Coggins, 2007; McAllister and Bursey, 2012; McAllister et al., 2014b). Additional helminths include the trematodes Allassostomoides sp., Clinostomum marginatum, Echinoparyphium sp., Echinostoma sp., Fibricola texensis, Glypthelmins quieta, Gorgodera bilobata, Gorgoderina attenuata, G. bilobata, G. translucida, Haematoloechus similiplexus, Megalodiscus temperatus, Mesocoelium monas,and Ostiolum medioplexus (Bouchard, 1951; Ulmer, 1970; Brooks, 1975; Coggins and Sajdak, 1982; Dyer, 1991; Cross and Hranitz, 1999; Yoder and Coggins, 2007; McAllister et al., 2008, 2014b) and the cestodes Cylindrotaenia americana, Distoichometra bufonis, Ophiotaenia saphena, Mesocestoides sp. and ­others (Ulmer and James, 1976a; Dyer, 1991; Yoder and Coggins, 2007; see reference list in Green, 2005; McAllister et al., 2014b). The trematode Echinoparyphium does not reduce the prevalence of ranavirus infection in this species, nor does it reduce the viral load compared with individuals not infected by Echinoparyphium (Wuerthner et al., 2017). An unknown species of acanthocephalan has also been reported (McAllister et al., 2014b). Biflagellated algae similar to Chlorogonium have been found along the lateral margins of American Toad tadpoles

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in Missouri (Drake et al., 2007) and Arkansas (Tumlison and Trauth, 2006). The alga may function to increase the amount of oxygen available to tadpoles, especially during thermal stress. In turn, the facultative symbiosis benefits the alga by making CO2 available to it from the tadpole’s waste products. American Toads may be parasitized by the green blowflies Lucilia elongata and L. bufonivora, but infections do not appear to be common (Anderson and Bennett, 1963; Bleakney, 1963; Briggs, 1975; Bolek and Coggins, 2002; Whitworth et al., 2021). Mortality is high, however, in infected individuals. Other ectoparasites include the leeches Desserobdella picta (Briggler et al., 2001; Bolek and Janovy, 2005) and Macrobdella decora (Blais, 2016), and the chigger Hannemania dunni (McAllister and Durden, 2014). Unidentified leeches and chiggers have been reported from toads in ­Virginia (Gibson and Sattler, 2020b). American Toads living in areas where ­there are high concentrations of pesticides often have a considerable prevalence of hind limb deformities. In Québec, Ouellet et al. (1997) found that 17% of 252 metamorphosing American Toads had ectromelia (missing or partly missing hind limb) or ectrodactyly (missing toes) from 3 locations adjacent to agricultural fields; at 1 location, the percentage was 69%. However, malformations have also been found on national wildlife refuges, especially affecting the hind limbs and feet (Converse et al., 2000). In Minnesota, malformations ­were found in 6.7% of American Toads examined (Hoppe, 2005). Toad larvae exposed to “cocktails” of pesticides also may have delayed metamorphosis, experience low survivorship, or, at severely impacted sites, be unable to breed. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. Aluminum toxicity is inversely correlated with pH in American Toads (Freda et al., 1991). However, total aluminum (31–1,155 µg/L) and labile aluminum (1–1,073 µg/L) had no effect on transplanted A. americanus embryos or tadpoles at 16 ponds in Ontario (Freda and McDonald, 1993). In laboratory ­trials, embryos ­were most stressed by aluminum at pH = 4.2. Cadmium at environmentally relevant doses has no effect on adult survival, percentage of body mass lost during winter dormancy, or locomotor per­for­mance (James et al., 2004). However, toads fed mealworms with 4.7µg/g dry weight had only a 56% survivorship compared with 100% for controls. Aqueous cadmium decreased larval survivorship ­under experimental conditions but had no effect on larval body mass at metamorphosis. Instead, the time it took to reach metamorphosis was increased (James et al., 2005). ­These authors hypothesized that cadmium has indirect effects on larvae by decreasing the abundance of detritus and periphyton. Doses

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Anaxyrus americanus 35

of cadmium >18 µg/L appear to be very toxic. The presence of American Toads is negatively correlated with nickel concentrations in Ontario (Glooschenko et al., 1992). American Toad larvae do not avoid detrimental concentrations of lead ­under laboratory conditions (Steele et al., 1991). In experimental ­trials, lead has no effects on gross tadpole locomotor per­for­mance. Likewise, pre-­exposure to lead did not increase sensitivity to subsequent encounters with lead (Steele et al., 1999). Mercury. Mercury is known to bioaccumulate in American Toads (Bergeron et al, 2010a). At a site contaminated by mercuric sulfate in ­Virginia, toad blood had a gradient of 82–4,235 ng/g wet weight of mercury, toes ranged from 32–602 ng dry weight, and eggs ranged from 15–205 ng/g dry weight (Bergeron et al., 2010b; Todd et al., 2012a).Total mercury concentrations of ≥775 ng/g dry weight are sufficient to cause sublethal maternal effects in offspring in laboratory ­trials (Bergeron et al., 2010b, 2011a). This is the equivalent of 209.7 ng/g dry weight in toes. Toadlets from ­mothers experimentally exposed to mercury had 2.5–5 times the prevalence of spinal malformations compared with controls, and their smaller size at metamorphosis meant that they did not hop as well as controls (Todd et al., 2011b, 2011c). Mercury-­exposed larvae also are vulnerable to predators ­because of their smaller size and the presence of spinal malformations that affect escape be­hav­ior (Todd et al., 2011c). Tadpoles in their natu­ral habitats ­were found to have a mean of 2,132 ng/g of total mercury. Mercury resulting from maternal exposure also results in reduced hatchling success, since mercury is transferred to the eggs (Bergeron et al., 2010a); ­there is a negative correlation between egg hatching success and mercury concentration (Bergeron et al., 2011b). Paradoxically, surviving larvae had a greater metamorphic success. ­These and other results (below) emphasize the need for multiple assessments of contaminant effects; a ­simple 96 hr toxicology trial does not tell the ­whole story. Exposure to mercury has serious detrimental effects to larval development regardless of ­whether the mercury comes from the ­mother or through the diet. Both maternal and dietary mercury significantly affected growth and development ­until the onset on metamorphosis in experimental ­trials. Both dietary mercury (10 µg/g dry weight) and maternal mercury reduced larval mass, but not the duration of the larval period. ­These sublethal effects, when combined, interact with lethal consequences, however. Larvae exposed to mercury from a ­mother living in a mercury-­contaminated environment and then fed diets spiked with mercury had 50% greater mortality compared with reference ­mothers fed a control diet (Bergeron et al., 2011a). ­These results suggest that the effects of contaminants ingested by the ­mother and

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transferred to offspring may be further exacerbated by ­later exposure to mercury as the larvae develop, although ­there ­were no interactive effects of exposure to maternal and dietary mercury (Todd et al., 2011b). Mercury exposure reduces the size of larvae at metamorphosis, and t­ hese effects persist for about a year postmetamorphosis. However, mercury exposure does not affect survivorship. Indeed, no new novel adverse effects developed a­ fter 1 yr in experimental ­trials, and ­there was no evidence of per­sis­tent effects of dietary mercury ingestion in terrestrial toads that could be attributed to ­earlier exposure (Todd et al., 2012b). Other ele­ments. Boron reduces hatching success in A. americanus. At low concentrations (50 mg L-1), 46.7% of eggs hatched, whereas at high concentrations (100 mg L-1), only 11.3% hatched in experimental ­trials in Pennsylvania. In control ­trials, 82% of eggs hatched (Laposata and Dunson, 1998). Since wastewater boron has been mea­sured at 169 mgL-1, such concentrations could adversely affect toads breeding at sites where wastewater effluent is disposed. Selenium is passed down from ­mothers to offspring via their eggs, and selenium is known to bioaccumulate in toads (Bergeron et al., 2010a). Mercury may facilitate the amount of selenium transferred from ­mothers to eggs. ­Under natu­ral conditions, American Toads do not selectively avoid sites contaminated by polychlorinated biphenyls (PCBs) (Gibbs et al., 2017). pH. Even very short-­term exposure of American Toad larvae to pHs of 3.0–3.5 ­causes 100% mortality (Leftwich and Lilly, 1992). In a study of 16 ponds in Ontario, survivorship of larval American Toads was significantly reduced (5–28%) in 4 of 7 ponds with a pH 80%), however (Freda and McDonald, 1993). In laboratory ­trials, ­there is a significant increase in embryo mortality at a pH of 4.2 and 0 µg/L of aluminum (Freda and McDonald, 1993). Freda (1986) and Freda et al. (1991) reviewed the lit­er­a­ture on the effects of pH on American Toad embryos, and suggested that lethal pHs ­were 3.8–4.2, with critical pHs at 4–4.2. Karns (1983) reported 100% egg fertilization in bog ­water with a pH of 4.2, although ­these embryos died of developmental abnormalities. Based on caged, transplant field experiments, Dale et al. (1985) recorded 100% mortality in A. americanus eggs at a pH of 4.1, and 10–58% mortality at a pH of 6.3. In laboratory ­trials, American Toad larvae increase their activity in the presence of acidified ­water, and ­were able to avoid ­water with lethal and sublethally low pH (Freda and Taylor, 1992). DOC. DOCs of 4–29 mg/L have no effect on larval survivorship of American Toad tadpoles (Freda and McDonald, 1993). Larvae can survive at low oxygen levels (20 years previously. Toads ­were found generally at higher elevations in the western and southern portions of the survey area, and ­were associated with open habitats

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38  Bufonidae

Anaxyrus americanus breeding habitat. ­Great Smoky Mountains National Park, Tennessee. Photo: C.K. Dodd, Jr.

(pastures), mixed deciduous forests, non-­developed areas, and low levels of acid and base deposition; toads tended to be absent from evergreen forest. In Illinois, no trends ­were discernable based on call counts from 1986–1989, and American Toads ­were heard on >85% of call survey routes (Florey and Mullin, 2005). However, Minton (2001) noted that they ­were not as abundant as they ­were 40 yrs ago in Indiana farmland and urban settings. In the Northeast, populations ­were considered declining in 4 states (Delaware, Mas­sa­chu­setts, Pennsylvania, West ­Virginia) based on 7 yrs of data using occupancy modeling (Weir et al., 2009). Meshaka et al. (2020a) noted a decline in a protected area in western Pennsylvania, a decline they attributed to forest succession as the area gradually changed from open habitats favorable to American Toads to a more closed forested habitat. Villena et al. (2016) suggested populations in the South ­were declining based on probability of occurrence through time. Like most amphibians, the primary threat to American Toads comes from habitat loss or alteration. As early as 1909, concern was being expressed about the effects of agriculture and wetland loss on the distribution of this species (Miller, 1909a). The presence of American Toads is negatively correlated with the extent of human-­based development as well as the extent of forested land, within 3,000 m of breeding sites around the ­Great Lakes (Price et al., 2004). In Iowa, a survey of collections made prior to 1950 compared with collections made ­after 1950 suggested a 29% decline in toads, possibly due to continuing loss of habitat (Christiansen, 1998). In contrast, ­there appeared to be no change in status of this species between past and recent surveys in Iowa (Christiansen, 1981; Lannoo et al., 1994); the differing results may be due to survey methodology, rather than changes in occupancy or abundance.

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Habitat fragmentation may be more of a threat to juvenile dispersal than to adults, inasmuch as juveniles tend to avoid open habitats and prefer deciduous leaf litter substrates to ­those of conifers or bare soil (Rothermel and Semlitsch, 2002; Smith and Schulte, 2008). The effects of fragmentation, such as occurs in agriculturally dominated landscapes, may be ameliorated by providing shrubby or wooded riparian strips allowing for dispersal (Maisonneuve and Rioux, 2001). Ponds in agricultural areas also can be enhanced as breeding sites by limiting nitrogen influx, limiting access by cows, and by not stocking fish (Knutson et al., 2004). In Ontario, toad abundance was negatively associated with mining activities, both in terms of metal contamination and in alteration of vegetative structure (Sasaki et al., 2015). While American Toads are affected by some forms of habitat disturbance, the response is not always negative. When habitats are opened up, such as when peat bogs are mined, toad abundance may actually increase around wetland margins. This might result from a greater tolerance of American Toads to dry conditions, or it could reflect an a­ ctual preference for the habitat conditions found in mined peat bogs (Mazerolle, 2003). The presence of ­cattle in breeding ponds had ­little effect on postmetamorphic American Toads, and Burton et al. (2009) suggested that toads may benefit from controlled grazing. In the Southern Appalachians, American Toad abundance did not differ among control forest sites and sites where ­there ­were gaps created by wind disturbance or salvage logging (Greenberg, 2001a). ­After 1–2 yrs post-­harvest, American Toad abundance actually increased on timber harvested sites in Maine, although toads ­were still more abundant in 11 and 23 m buffer strips along headwater streams than in the clearcuts per se (Perkins and Hunter, 2006). In Missouri, most American Toads ­were captured leaving an area that had been clearcut during 2 yrs following the clearcut (Semlitsch et al., 2008), although experimental evidence suggested that partial harvests actually increase survivorship (Earl and Semlitsch, 2013). In response to fire, toads burrow into the ground where they may suffer from superficial burns (Pilliod et al., 2003), but populations do not appear to be adversely affected by periodic fire (Greenberg et al., 2018b). They do survive in burned areas (Floyd et al., 2002; Keyser et al., 2004), and their terrestrial abundance ­later may actually increase as habitats are opened up (Kirkland et al., 1996). The ability to detect amphibian species at breeding sites is not constant from year to year. Skelly et al. (2003) used data from American Toads, in part, to suggest that resurvey results vary depending on the duration and spatial scale of the detection effort. American Toads had only a 0.3 probability of presence (naïve estimate), leading ­these authors to suggest that a population may shift breeding sites through time, despite the tendency to be site philopatric in breeding

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Anaxyrus americanus 39

site choice from one year to the next. Further, the importance of multiple site visits to estimate occupancy throughout a breeding season is emphasized by the results of MacKenzie et al. (2002). They computed detection probabilities for A. americanus at 29 Mary­land sites visited from 2–66 times (mean 9.6) throughout a breeding season, and determined an occupancy estimate of 0.49, 44% higher than the proportion of sites where toads ­were actually observed. Hecnar and M’Closkey (1996a) noted short-­term increases in occupancy over a 2 yr period which may suggest shifting breeding sites, but it is difficult to make trend statements based on short-­term data. The fact that American Toads may shift breeding sites if fish become established, and that they rapidly colonize newly created ponds, lends credence to the “shifting breeding site through time” hypothesis. The likelihood of changing breeding sites temporally, spatial scale, duration of surveys, and the number of visits per site all need to be incorporated into the design of monitoring programs for this species. Roads and other high-­volume transportation corridors do have significant effects on American Toads, as they do on many other frog species (e.g., Cunnington et al., 2014). For example, 433 A. americanus ­were killed on a 3.6 km stretch of highway in Ontario over a 4 yr period (Ashley and Robinson, 1996), and 111 A. americanus ­were counted dead on 4 Indiana survey routes over a 1 yr period (Glista et al., 2008). On impervious roads and at higher road densities, A. americanus populations are negatively impacted at small local scales, but not over a more widespread regional scale (Marsh et al., 2017). As might be expected, most mortality is seasonal, associated with spring immigration and juvenile emigration. American Toads may not use existing culverts to cross some roads (Patrick et al., 2010), so more experimental work is necessary to facilitate traverse across roads and highways. The spatial effects of highway mortality are most pronounced within 500 m of a breeding site. Near Ottawa, Ontario, for example, American Toad abundance was significantly negatively correlated with traffic density, at least within 2,000 m of breeding ponds (Eigenbrod et al., 2008). Since American Toads easily disperse as far as 1,000 m from wetlands (Forester et al., 2006), the negative effects of roads with high traffic volume extend considerable distances across a landscape. The area of immediate adverse impact (the “road-­zone” effect) extends 200–300 m from the pavement (Eigenbrod et al., 2009). Traffic noise does not appear to inhibit calling by this species. Vargas-­Salinas et al. (2014) found that American Toads, a species with a high call peak frequency, called randomly with regard to traffic noise intensity in eastern Ontario. Toads also do not alter their call characteristics in response to low or high volumes of traffic noise (Cunnington and Fahrig, 2010), and their calls are

Dodd_Canada_int_5pgs_B1&B2.indd 39

equally effective at attracting females regardless of traffic noise (Cunnington and Fahrig, 2013). On the plus side of transportation corridors, American Toads ­will use terrestrial habitats and wetlands located in power-­line rights-­of-­way as breeding sites if such corridors go through deciduous forest (Yahner et al., 2001; Fortin et al., 2004). A most unusual source of mortality of American Toads was reported by Miller (1909b). He noted that sewer traps caught large numbers of toads, and that sewer cleaners removed “piles of toads” each fall and spring. He went on to estimate that 24,000 toads ­were caught in the sewers of Worcester, Mas­sa­chu­setts, alone, and that mortality could approach 50,000 toads per year. It is likely that such human-­made traps have caught toads in virtually ­every municipality throughout the species’ range, even to the pre­sent day. Moore and Hecnar (2018) also reported American Toads killed by an electric fence. Inasmuch as A. americanus often breeds in fishless ponds, the introduction of fish may lead to the disappearance of breeding toads (Sexton and Phillips, 1986). Nonindigenous anurans also may impact American Toads. Howard (1988a) noted 10 males at an Indiana breeding site 1 year, where ­there had been 200 males and 60 females 2 years ­earlier; the decline coincided with the introduction of American Bullfrogs (L. catesbeianus) into the pond. In captivity at least, the Cane Toad (Rhinella marina) outcompetes American Toads (Boice and Boice, 1970), but since ­these species do not occur together, this result may be more pertinent to the status of Southern Toads (A. terrestris) in Florida than to American Toads in nature. American Toads readily and rapidly colonize newly constructed or restored breeding ponds (Briggler, 1998; Kline, 1998; Mierzwa, 2000; Stevens et al., 2002; Touré and Middendorf, 2002; Weyrauch and Amon, 2002; Brodman et al., 2006; Barry et al., 2008; Shulse et al., 2010; Denton and Richter, 2013; Bartelt and Klaver, 2017; Baecher et al., 2018). Bartelt and Klaver (2017) noted colonization within 2 yrs. Toads moved from 42–2,932 m to reach the restored sites, traversing row crop habitats to do so. In the Prairie Pothole Region, they may use restored conservation grasslands, but they are more common in native habitats (Ba­las et al., 2012). ­Water in retention ponds, however, may have chemicals, resulting in sublethal effects, such as reduced size at metamorphosis (Snodgrass et al., 2008). If golf courses can be managed so as to reduce effects from American Bullfrogs and fish, even ­these highly artificial wetlands can serve as breeding habitat for American Toads (Mifsud and Mifsud, 2008). However, if bullfrog larvae are allowed to overwinter, then toad survivorship to metamorphosis is reduced. One way to reduce negative effects is to manage ­these wetlands by creating variable hydroperiods that do not allow bullfrog overwintering or the presence of predatory fish (Boone et al., 2008).

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40  Bufonidae

Anaxyrus baxteri (Porter, 1968) Wyoming Toad ETYMOLOGY

baxteri: a patronym honoring George T. Baxter (1919– 2005), a University of Wyoming professor who discovered the species in 1946 while working on his master’s degree, and ­later called attention to its near extinction. NOMENCLATURE

Stebbins (2003): Bufo baxteri Fouquette and Dubois (2014): Bufo (Anaxyrus) baxteri Synonyms: Bufo hemiophrys baxteri IDENTIFICATION

Adults. The Wyoming Toad is a small (to 68 mm SUL), dark-­brown, gray, or greenish toad with dark blotches and an indistinct, narrow, mid-­dorsal stripe. The elongate boss between the eyes is prominent (narrow and high), and the parotoid gland is more distinct, elevated, smoother, and frequently separated from the indistinct postorbital crests than in the closely related Canadian Toad. Many warts cover the dorsum, and ­these warts form unique patterns, lasting from metamorphosis through adulthood, that can be used to uniquely identify individuals (Morrison et al., 2016). Tubercles on the rear feet are well developed for digging. Males have a dark throat patch. Males are slightly smaller than females, ranging from 46–57 mm SUL as opposed to 47–68 mm SUL (Withers, 1992). Smith et al. (1998) gave an adult size range of 47.0–59.5 mm SUL (mean 52.7 mm). Sanders (1987) and Smith et al. (1998) discussed additional aspects of morphology and cranial osteology. Larvae. The larvae are small and black, reaching 25– 27 mm TL just prior to metamorphosis. Eggs. The eggs are small (2–3 mm in dia­meter) and black. They are deposited in long gelatinous strings, 1 from each oviduct. ­There are about 10–12 eggs per cm of string (Withers, 1992). DISTRIBUTION

This species is endemic to the Laramie Basin of Wyoming and is currently known only from the vicinity of Mortenson Lake. Historically it occurred at Porter Lake and other nearby localities, but no longer does so. The range of this species is shown by Baxter and Stone (1985). FOSSIL REC­O RD

­There are no fossils reported for this species.

Dodd_Canada_int_5pgs_B1&B2.indd 40

Distribution of Anaxyrus baxteri

SYSTEMATICS AND GEOGRAPHIC VARIATION

Anaxyrus baxteri is a member of the Americanus clade of North American bufonids, a group which includes A. americanus, A. fowleri, A. hemiophrys, A. houstonensis, A. terrestris, and A. woodhousii. It is closely related to A. hemiophrys, and somewhat more distantly to A. fowleri, A. americanus, and A. houstonensis (Pauly et al., 2004). This species was originally described as a subspecies of the Canadian Toad based on differences in morphology, parotoid venoms, and advertisement calls (Porter, 1968). Relying on a variety of morphological and call characteristics, Smith et al. (1998) elevated it to specific status. High percentages of metamorphs ­were produced in laboratory crosses between A. baxteri and A. hemiophrys (Porter, 1968). ADULT HABITAT

The Wyoming Toad is known historically only from short-­grass communities near ponds, seepage lakes, and in adjacent floodplains of the Laramie Basin. The species was associated with wetlands in wind-­erosion basins that contained vegetated sand dunes used for burrowing. At Mortenson Lake, it is found in a mixed-­sedge (Eleocharis palustris–­Scirpus americanus) shoreline community. The species is associated with short (10 m from shore are rare, and the distance from calling site to overwintering site may only be 30 m. ­After breeding, the toads often disperse from the calling sites southward to a nearby ditch, the berms of which provide overwintering sites. Emergence occurs in May when daytime temperatures reach 22°C. Activity occurs throughout the warm season, at least to October, with preferred substrate temperatures reaching >20°C. Juveniles prefer slightly warmer substrates than adults. Wyoming Toads are diurnal and tend to become dormant at night. During the warmest part of the day, they may dig themselves into shallow depressions. They also spend a considerable amount of time down mammal burrows (Parker, 2000). Some toads go back and forth between the burrows and foraging areas, whereas ­others may remain in burrows for many consecutive days. Toads likely use ground squirrel or pocket gopher burrows for overwintering in addition to the berm near a ditch used by calling toads (Withers, 1992). Parker (2000) excavated 1 burrow and found a toad 36 cm below the surface. Dispersal by recent metamorphs appears rather ­limited, and does not occur even to the north shore of Mortenson Lake. CALLING ACTIVITY AND MATE SE­L ECTION

Calls are heard from late May to June, with a calling season lasting from 9 to 36 days (Withers, 1992). Calls appear to be initiated by rising temperatures and usually occur ­after the last spring frost, although a sudden cold spell can interrupt the breeding season. Males arrive before females at the breeding pond. Males call while sitting in shallow ­water (4–11 cm) or floating in leaf litter at the ­water’s surface. Calling occurs in the vicinity of emergent vegetation at ­water temperatures of 18–22°C. Peak calling air temperatures are 21–27°C, but calls have been heard as low as 10°C (Withers, 1992). At Mortenson Lake over a 4 yr period, males called from 1 to 6 calling centers with each center containing 2–15 males; occasional lone males also ­were heard calling

Dodd_Canada_int_5pgs_B1&B2.indd 41

(Withers, 1992). ­These calling centers may shift during the breeding season and annually. The call of the Wyoming Toad is a buzzing trill characterized by its longer duration (4–12 sec at 15°C), slower repetition rate (37–48 pulses/sec; mean 41.7), and a dominant frequency of 1,450–1,700 cps when compared with the closely related A. hemiophrys (Porter, 1968). ­There may be a 7–9 sec interval between calls. The calls can be heard by a ­human observer 200 m away. Calling occurs both diurnally and at night, but is strictly temperature-­dependent. When grasped, male Wyoming Toads have a release vibration and warning call. BREEDING SITES

Breeding occurs only in the shallow, littoral w ­ aters mostly on the eastern and southern sides of Mortenson Lake. Calling also occurs at a nearby ditch, but breeding has not been documented ­there. REPRODUCTION

Oviposition occurs from mid-­May to early June, depending upon environmental conditions. Eggs are deposited in long strings in shallow ­water (3.5–6.3 cm) among low vegetation. The strings may be placed in a helical clump or deposited linearly through the vegetation. The pH at Mortenson Lake is 6.6–8.6, and conductivity should be 900 captive-­reared toads ­were released in the hope of maintaining a wild population. ­There are a number of hypotheses regarding the decline of this species, chiefly the effects of land alteration and the lethal effects of pesticide (fenthion) spraying. The timing of anuran disappearances in the Laramie Basin coincided with the initiation of fenthion spraying for mosquitoes. Not only ­were Wyoming Toads affected, but Northern Leopard Frogs

Dodd_Canada_int_5pgs_B1&B2.indd 43

as well (Baxter et al., 1982). However, ­there is no direct evidence that this pesticide caused the population declines. ­Later, the amphibian chytrid fungus was identified from Wyoming Toads (see Diseases), and is now thought to have played a decisive role in the near extinction of this species (Odum and Corn, 2005). The species was federally listed as Endangered in 1984 ­after the toad was thought extinct in the wild, although listing was sought much ­earlier (CKD, memorandum, Office of Endangered Species, 1982). A Recovery Plan was approved in 1991, but a Recovery Team was not appointed ­until 2001, 17 years ­after the species was listed. The population has been monitored continuously, and some management recommendations have been implemented. ­Limited sight-­based count surveys ­were used to judge the results of the captive release program, but ­these efforts provided data only for the years 1990–1992 and ­were considered of ­limited value. ­Cattle ­were allowed to graze in the area in an attempt to control vegetation as per the recommendations of Withers (1992), but ­cattle may put dormant toads at risk and trample dense vegetation (Parker, 2000; Parker and Anderson, 2003; Linhoff and Donnelly, 2016). Withers (1992) recommended a number of additional management actions, including relocating a boat ramp that bisected the Wyoming Toad’s habitat and phasing out fishing within the lake. Recent research has centered on captive rearing conditions and the effects of habitat management on toadlet survival. Recent metamorphs are 11–15 cm, and the size at metamorphosis is positively related to forb cover up to 35%. Toadlets do better in low (0–10 cm) and high vegetation (>30 cm), rather than at mid-­level (10–30 cm) vegetation heights, although the experimental treatment effects ­were not significant. Polasik et al. (2016) recommended that in field mesh enclosures, time to metamorphosis would be decreased in warmer ­water. They further suggested that vegetation heights of 10–30 cm and up to 35% forb cover within ­these enclosures could increase postmetamorphic toadlet size, which in turn could increase overwinter survival rates. Other recommendations included using small-­scale vegetation manipulation (mechanical, prescribed burning, ­limited ­cattle grazing) and soft release re­introduction techniques. Despite successes in captive propagation, the re­ introduction program has met with repeated setbacks and unsatisfying results. In general, past recovery efforts ­were hampered by bureaucratic infighting, lack of rigorous habitat management, reliance on non-­peer-­reviewed interpretations of data, often questionable restrictions on research, and poor communication (Dreitz, 2006). Despite many past years of in­effec­tive management, the Wyoming Toad continues to survive in captivity and, hopefully, in at least a small wild population. Currently, the USFWS estimates that ­there are

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44  Bufonidae

about 500 Wyoming Toads in captivity. Large numbers of animals have been released at Mortenson and 2 other sites (Lake George and Rush Lake at Hutton Lake National Wildlife Refuge). The agency has proposed (as of 2020) establishing a conservation area along the Laramie and ­Little Laramie Rivers near the Hutton Lake and Mortenson Lake

National Wildlife Refuges in southeastern Wyoming. The area would comprise a proposed area of about 19,682 ha in Albany County, Wyoming to conserve this species and would be managed as part of the Arapaho National Wildlife Refuge Complex (see https://­www​.­fws​.­gov​/­mountain​-­prairie​/­refuges​ /­wtca​.­php).

Anaxyrus boreas (Baird and Girard, 1852) Western Toad Crapaud de l’Ouest ETYMOLOGY

boreas: from the Greek boreas meaning ‘north wind’ or ‘northern’; halophilus, from the Greek halos meaning ‘sea’ or ‘salt,’ and philos meaning ‘having an affinity for.’ NOMENCLATURE

Stebbins (2003): Bufo boreas Fouquette and Dubois (2014): Bufo (Anaxyrus) boreas Synonyms: Bufo boreas halophilus, Bufo canagicus, Bufo canagicus halophilus, Bufo columbiensis, Bufo columbiensis halophilus, Bufo halophila, Bufo halophilus, Bufo laminator, Bufo lentiginosus pictus, Bufo nestor, Bufo pictus, Bufo politus, Rana canagica IDENTIFICATION

Adults. Anaxyrus b. boreas (Western Toad) has a wide range of dorsal coloration, from light gray to greenish to a dull black; most are brownish gray. This medium-­sized toad has numerous warts (granular glands) covering its dorsum which are light brown in color. The parotoid glands are oval and smooth, and they and the warts may be tinged in red. The Western Toad is distinguished from most other toads within its range by its complete lack of supraorbital and postorbital cranial crests (but see accounts of the recently described A. monfontanus, A. nevadensis, and A. williamsi). Adults have a white vertebral stripe down the ­middle of the dorsum, and the venter is dull white with vari­ous amounts of spotting. ­There may be a light area ­under the eye and a con­spic­u­ous black blotch between the thighs ventrally. The limbs are barred or blotched with melanin. Tubercles and toes may be tipped with orange. Males have an enlarged nuptial pad on the thumb during the breeding season, but not a dark throat as do many other Anaxyrus; ­there is no vocal pouch (but see Awbrey, 1972). Anaxyrus b. halophilus (California Toad) is less blotched (reduced dorsal melanin) than A. b. boreas, has a wider head

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Head pattern of Anaxyrus boreas. Illustration by Camila Pizano

and larger eyes, smaller feet, and a weaker development of the margins along the dorsal stripe. In addition, ­there is considerable regional variation in the extent of dorsal and ventral melanin and in the relative width of the dorsal stripe (Karlstrom, 1962). Some of this morphological variation may stem from the dif­fer­ent evolutionary histories of the vari­ous toad populations currently recognized ­under the name A. boreas. Juveniles are patterned like adults but may have red warts dorsally and they lack the white mid-­dorsal stripe. They also have bright yellow or orange flecks on the bottoms of their feet and body. Burger and Bragg (1947) provided a detailed description of recently transformed toadlets. In general, Western Toads from higher elevations (mountains) are smaller than conspecifics from low elevations (along the coastal Pacific Northwest). Karlstrom (1962) noted that ­there was considerable variation among populations in terms of mean and maximum sizes. The largest male he found was 112 mm SUL, and the largest female 120 mm SUL. Mean sizes (in mm SUL) for males and females are as follows: Alaska (59.5, 68.7); British Columbia (84.5, 86.2); Oregon (78.3, 93.1); Butte and Tehama Cos., California (85.9, 87.7); Contra Costa and Alameda Cos., California (84.3, 93.3); Alpine, Placer and Plumas Cos., California (80.5, 95.6); Mariposa County, California (81.3, 90.6);

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Anaxyrus boreas 45

Mariposa, Madera, and Merced Cos., California (94.3, 99.1); Fresno and Tulare Cos., California (76.1, 86.2); Kern County, California (75.2, 83.1); Colorado (68.5, 80.2); Utah (83.4, 94.1); Mono and Mineral Cos., California (71.5, 81.6); Inyo County, California (67.3, 81.4); Los Angeles and San Bernardino Cos., California (78.4, 85.7); San Bernardino County, California (73.1, 75.7); and Baja California, Mexico (70.4, 77.4) (Karlstrom, 1962). Additional size rec­ords are available for southern Utah (males: mean 86.8 mm SUL, range 75–98 mm; females: mean 96.3 mm SUL, range 81–111 mm; Robinson et al., 1998), western Montana (adults 55–125 mm SUL; means 89–95 mm; Adams et al., 2005), Idaho (mean SULs of males 69.9 mm and females 78.4 mm; Bartelt et al., 2004), Oregon (depending on location, males a mean of 83–95 mm SUL, maximum 125 mm; females a mean of 93–119 mm SUL, maximum 130 mm; Bull, 2006), and the northern ­Great Basin (mean 82.3 mm SUL, range 53–133 mm; Gordon et al., 2020). Some individuals may reach 145 mm SUL (Jones et al., 2005). Larvae. The larvae are usually jet black and they have no iridescence on the body. However, they sometimes are lighter in coloration than the deep black color of other Anaxyrus, and they have a very transparent tail. The tail musculature is not bicolored in lateral view and the tail fin appears clouded. In Colorado, some larvae become olive brown just prior to metamorphosis and have pigmented tail fins. Livo (1999) suggested that this coloration was an environmentally induced phenotype in response to predation by Garter Snakes (Thamnophis). In profile, the snout is flattened in appearance, and the tail tip is pointed and deepest at one-­quarter of its length. When viewed from above, the body is broadest between the snout and the spiracle. The largest A. b. halophilus larvae mea­sure 56 mm TL (Storer, 1925), although larvae in Colorado mea­sured 34–37 mm TL just prior to metamorphosis (Burger and Bragg, 1947). Albino larvae ­were reported from Washington by Hensley (1959). A description of the larvae is in Burger and Bragg (1947). Eggs. The Western Toad oviposits 3,000–8,000 eggs per clutch in long, mostly double, strings. Jelly partitions do not separate individual eggs from one another within the continuous string tube as they do in A. canorus. In A. b. halophilus, the dia­meter of the outer envelope mea­sures 4.9–5.3 mm; the inner envelope mea­sures 3.5–3.8 mm; the dia­meter of the ovum mea­sures 1.65–1.75 mm. ­There are 13–52 eggs per 2.54 cm of string (Karlstrom, 1962); Savage and Schuierer (1961) counted 120 eggs per 10 cm for 2 strings. Werner et al. (2004) noted that the egg strings from a single female can mea­sure 20 m in total length. Egg strings oviposited near algal mats may incorporate algae into the gelatinous string, giving a green hue to the eggs. Additional

Dodd_Canada_int_5pgs_B1&B2.indd 45

details on the eggs of this species are in Karlstrom (1962). Savage and Schuierer (1961) illustrate the eggs. DISTRIBUTION

Western Toads are among the farthest northward-­distributed amphibians in North Amer­i­ca and occur from near sea level along the West Coast to 3,655 m in California and Colorado (elevation rec­ords in Stejneger, 1893; Karlstrom, 1962; Campbell, 1970d; Salt, 1979; Livo and Yeakley, 1997; Keinath and Bennett, 2000; Muths et al., 2008). In the Rockies, they occur at elevations >2,100 m. Anaxyrus b. boreas is found from southern Alaska and the southwestern Yukon Territory south to northern California and much of northern Nevada. The range includes much of southern British Columbia, west central Alberta, western Montana and Wyoming, and northern and central Utah. Isolated populations are in the southeastern Yukon Territory (Liard River drainage), northern Alberta, and discontinuous in Nevada and southwestern Utah; Thompson et al. (2004) reported populations in 12 geographic areas of Utah. Historically, the species occurred from south central Wyoming south through the Rocky Mountains to northern New Mexico. Many of ­these populations have dis­appeared, and the species likely no longer occurs throughout much of its former range in the central and southern Rockies. Anaxyrus b. halophilus occurs from northern California and western Nevada south into Baja California. Scattered populations are found in isolated areas of the Mojave Desert.

Distribution of Anaxyrus boreas

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46  Bufonidae

Western Toads are found on islands, including ­Middle (Barrier Islands group), Long, Prince of Wales, Mitkof, Wrangel (Alaska), Vancouver Island, the Queen Charlotte Islands, the Alexander Archipelago, and islands in Prince William Sound and Puget Sound (Bainbridge, Blakely, Camano, Cypress, Fidalgo, Harstine, Lopez, Orcas, San Juan, Shaw, Whidbey). Island locations are in Slevin (1928), Myers (1930a), Swarth (1936), Slater (Brown and Slater, 1939; Slater, 1941a), Andis (2018), and Ream et al. (2019). Impor­tant distributional references include: multiple states or provinces (Slevin, 1928; Logier and Toner, 1961; Jones et al., 2005; Corkran and Thoms, 2020), Alaska (Van Denburgh, 1898; Hock, 1957: Hodge, 1976; Andis, 2018; Ream et al., 2019), Alberta (Salt, 1979; Eaton et al., 1999; Russell and Bauer, 2000), British Columbia (Logier, 1932; Cowan, 1936, 1939; Carl, 1943; Carl and Hardy, 1943; Herreid, 1963; Cook, 1977; Matsuda et al., 2006; Ovaska and Govindarajulu, 2010; Slough, 2013), California (Storer, 1925; Karlstrom, 1958, 1962; Lemm, 2006; Flaxington, 2021), Colorado (Maslin, 1959; Smith et al., 1965; Hammerson, 1999), Idaho (Slater, 1941b; Tanner, 1941; Lucid et al., 2020), Mexico (López et al., 2009), Montana (Black, 1970, 1971; Franz, 1971; Marnell, 1997; Maxwell et al., 2003; Werner et al., 2004), Nevada (Ruthven and Gaige, 1915; Linsdale, 1940; Hovingh, 1997), New Mexico (Campbell and Degenhardt, 1971; Stuart and Painter, 1994; Degenhardt et al., 1996), Oregon (Gordon, 1939; Pearl et al., 2009a), Utah (Van Denburgh and Slevin, 1915; Tanner, 1931; Ross et al., 1995; Hovingh, 1997; Robinson et al., 1998; Thompson et al., 2004), Washington (Slater, 1955; Metter, 1960; McAllister, 1995), Wyoming (Baxter and Stone, 1985; Koch and Peterson, 1995), and the Yukon (Cook, 1977; Slough and Mennell, 2006). FOSSIL REC­O RD

The Western Toad is known from Pleistocene (Rancholabrean) deposits in California (at Rancho La Brea and Shasta Counties; Brattstrom, 1953, 1958), Colorado, and Nevada (Holman, 2003). Holman (2003) notes that a combination of characters is useful in identifying fossils of this toad, such as a lack of cranial crests on the fronto-­ parietal, a very narrow distil one-­third of the humerus, a moderately curved ilium, the anterior margin of the ventral acetabular expansion having a hemispherical curve with the shaft’s posterior, the apices being pointed rather than curved in the dorsal and ventral acetabular expansions, and the dorsal prominence being low with 2 or 3 small tubercles. Holman (2003) also contains illustrations of some of ­these features.

Dodd_Canada_int_5pgs_B1&B2.indd 46

SYSTEMATICS AND GEOGRAPHIC VARIATION

The Western Toad is a member of the Nearctic clade of New World toads. Two subspecies historically are generally recognized (A. b. boreas, A. b. halophilus), although systematic evaluation of toads referred to A. boreas is ongoing. Additional species may be described based on the results of molecular studies, and it seems certain that current taxonomy may not reflect the levels of ge­ne­tic divergence within Western Toads. Many Western Toad populations in the Southwest are found in isolated remnants of desert springs and riparian areas or located in wetter mountainous regions where they have differentiated as intervening habitat has become drier and unsuitable for toad survival (see accounts of A. monfontanus, A. nevadensis, and A. williamsi). Anaxyrus boreas has long been recognized as being related evolutionarily to a morphologically similar group of toads, including A. canorus, A. exsul, and A. nelsoni, together constituting the Boreas clade. Precise relationships have been evaluated differently depending on the type of data and analyses used (Tihen, 1962a; Blair, 1959, 1964; Graybeal, 1993, 1997; Shaffer et al., 2000; Stephens, 2001; Pauly et al., 2004; Goebel, 2005). At times, even Incilius alvarius has been considered a member of the group (Blair, 1959, 1963b, 1964), although it is not considered so at pre­sent. It seems, however, that ­there is a consensus that toads previously known as A. boreas and A. canorus are paraphyletic. Thus, the origin of the toads used in systematic research is impor­ tant for understanding phylogeny, and this may account for differing hypotheses of relationships. Goebel (2005) and Goebel et al. (2009) have identified 3 major lineages in the A. boreas group of toads, including a northwestern (NW), an eastern (E), and a southwestern (SW). Each of ­these major lineages contains minor lineages (but see Lucid et al., 2021). The toad currently recognized as A. b. boreas largely falls into the NW and E clades, with most A. b. halophilus allocated to the SW clade. However, it is clear that A. canorus is paraphyletic and related to both the NW and SW groups of A. boreas (also see Stephens, 2001), that A. nelsoni is related to A. b. halophilus, and that A. exsul is quite unique but also distantly related to A. b. halophilus and A. nelsoni. The Western Toads from Darwin Canyon in Inyo County, California, previously suspected of being a fully supported lineage as differentiated from NW A. boreas as are A. nelsoni and SW A. boreas, actually represent a release of individuals from Deep Springs (i.e., A. exsul) (C. Richard Tracy, personal communication, 2020). ­These lineages reflect differentiation among isolated populations of a much more contiguous Western Toad distribution that extended throughout western North Amer­i­ca prior to Pleistocene climate changes.

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Anaxyrus boreas 47

Hybridization is not uncommonly reported in the lit­er­a­ture where dif­fer­ent species or lineages come into contact. Natu­ral hybrids between A. boreas and A. canorus are known from Mono County, California (Morton and Sokolski, 1978). A suspected hybrid was reported from Alpine County, California (Mullally and Powell, 1958), although the identification has been questioned (Karlstrom, 1962). Natu­ral hybrids have been reported between A. boreas and A. punctatus in Darwin Canyon, California (Feder, 1979). Artificial hybrids have been produced between A. boreas and A. americanus, A. speciosus (as A. compactilis), A. punctatus, and Incilius nebulite (as Bufo valliceps); at least some development occurs in crosses with A. debilis and I. canaliferus (A.P. Blair, 1955; Blair, 1959, 1963b) and even South American and Eu­ro­pean bufonids (Blair, 1964). Intrapopulation ge­ne­tic structure has been studied in a few Western Toad populations. Gene frequencies among adults tend to be consistent from one year to the next within a population, but they may be dif­fer­ent from the larval cohorts inhabiting the same population (Samollow, 1980). This result might reflect se­lection acting for or against genotypes, especially since 90–95% of the larvae ­will never reach sexual maturity. Gene frequencies among cohorts also vary among individual ge­ne­tic loci. Ge­ne­tic structure was not detected among 16 populations of A. boreas in northern Idaho and northeastern Washington (Lucid et al., 2021). In British Columbia and Alberta, female Western Toads are dimorphic in color, with some yellowish and ­others reddish. Sympatric males are only yellowish in color. According to Schueler (1982b), the proportion of reddish females increases as one moves ­toward the coast. On Vancouver Island and in the lower Fraser Valley, however, this dimorphism is not apparent. The reddish-­yellowish coloration may reflect differences in habitat use during the short activity season (Bartelt et al., 2004), with reddish females inhabiting the forest floor where reddish conifer ­needles are predominant, and males remaining mostly in the vicinity of breeding sites characterized by yellowish muddy bottoms (Schueler, 1982b). A canorus-­like phenotype has been reported for the higher elevations of Glacier National Park (Black, 1971). ­These toads are smaller than typical A. boreas, have large dorsal warts set in blotches, and have a vertebral stripe that is broken rather than continuous. Toads from the southern Rocky Mountains, however, do not have any dichromatic color or pattern variation between the sexes. Western Toads from Fish Lake Valley, Esmeralda County, Nevada have a very spotted throat (Linsdale, 1940).

Dodd_Canada_int_5pgs_B1&B2.indd 47

ADULT HABITAT

This species occurs at higher elevations in the southern Rockies and other high mountains of the West in wet shrublands, crop and hayfields, and spruce-­fir, mountain fir, aspen, corkbark fir, Englemann spruce, bog birch, and willow and lodgepole pine forests interspersed with wet or moist open meadows. They are found from the mountain foothills to sub-­alpine meadows. In coastal areas, they are found among sand dunes, shrubs, and moist Sitka spruce climax forests (Pimentel, 1955). Throughout much of their range, Western Toads prefer open habitats (>50% open canopy), minimal ground cover, and areas with a more southern exposure (Bull, 2006; Browne and Paszkowski, 2014). However, Browne et al. (2009) found that abundance was associated with closed deciduous and mixed-­boreal forests or tall (but not short) shrubs at least at the 50–100 m scale in Alberta. They also prefer sites with high densities of refugia nearby. In a study of female movement patterns, for example, Bartelt et al. (2004) and Browne and Paszkowski (2014) found that females spent far more time in open-­ canopied habitats than in canopies where cover was >50%. Females also preferred forest edges, avoided clearcuts, and tended to seek refuge ­under shrubs that provide protection from dehydration. Males are more associated with ­water than are females (Browne and Paszkowski, 2014). Historically, a mosaic of open forests and meadows existed throughout the range of this species as overgrown areas ­were reopened due to periodic wildfires or other disturbances. ­After a fire, ground cover decreases, although coarse woody debris may actually increase due to tree fall. ­After an area is burned, A. boreas quickly colonizes the burned or partially burned areas (Guscio et al., 2007), even as unburned habitats go unoccupied, and populations may increase substantially for up to 3 yrs post-­burn (Hossack et al., 2013b). Such colonization may occur even when the closest breeding ponds are at a considerable distance away. The terrestrial wandering tendencies of some Western Toads may facilitate recolonization of burned or disturbed areas and allow them to respond quickly to disturbance events as long as cover sites are available to protect them against predators and dehydration. Hossack and Corn (2007b) and Rochester et al. (2010) noted, however, that populations of Western Toads declined ­after initial colonization of burned sites and 2 yrs post-­burn, respectively (also see Hossack et al., 2013b). Thus, it remains to be determined ­whether toad colonization is a short-­term response to fire disturbance, or ­whether ­there ­will be long-­term benefits to the population. Adult A. boreas have been found in saline Pyramid Lake, Nevada, where they take refuge from the heat and dry conditions in an area other­wise nearly devoid of shade or

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48  Bufonidae

vegetation. They also take refuge in pockets of tufa washed by wave action. Brues (1932) stated that “due to the conditions prevailing ­here they appear to have become as highly aquatic as frogs.” Tolerance of somewhat saline conditions allows them to inhabit an other­wise uninhabitable environment. TERRESTRIAL ECOLOGY

Western Toads walk rather than hop. When not breeding, adults are found commonly in shallow depressions called forms or in tunnels. The animals dig the form in a concealed location into moist soil that allows them to maintain a favorable thermal microenvironment as well as to absorb moisture directly from the soil. In addition, toads in forms are difficult to locate, presumably lessening predation chances and enabling them to be good sit-­and-­wait ambush predators. Campbell (1976) found that females ­were more likely to dig forms than males, and that as many as 25% of the females and 13% of the males could be found in forms. Other microhabitats favorable to adults are ­those that provide moist substrates and shelter with elevated relative humidity compared to surrounding habitats, such as woody debris and shrub cover, as well as ­those in proximity to breeding and overwintering sites (Long and Prepas, 2012; Browne and Paszkowski, 2014, 2018). In Alberta, se­lection for higher or lower percentages of canopy cover varied by site, sex, and season, and toads generally selected sites with taller vegetation, a lower percentage of herbaceous ground cover, and a higher percentage of shrubs; the type of vegetation was not impor­tant. Females selected disturbed grasslands (e.g., parklands and pasture), but males avoided or did not use ­these habitats very significantly. Toads avoided mineral soils, with preferences ­toward organic substrates and substrates with accumulated woody debris; such microenvironments had a greater moisture content. Linear corridor habitats ­were avoided by males for migration and foraging, with both breeding females and foraging males preferring pond edges (Browne and Paszkowski, 2018). Anaxyrus boreas normally is active from spring through November, depending on weather conditions and elevation, but activity begins in January along the coast (e.g., Nussbaum et al., 1983; Lemm, 2006). Activity may be somewhat bimodal especially at higher elevations, with breeding and feeding occurring from spring ­until the dry conditions of late summer inhibit daily activity. With cooler temperatures in the autumn, toads reemerge from shelters and feed ­until weather conditions turn cold. In Yosemite Valley, for example, A. b. halophilus become active from mid-­April to early May, and at Big Bear Lake, California, activity occurs from March to October (Mullally, 1952). ­After breeding, they depart the breeding sites and assume a more terrestrial

Dodd_Canada_int_5pgs_B1&B2.indd 48

existence, although many remain in the vicinity (within 100–300 m) of streams and other moist and sheltered locations. Most retire for the season in September and early October, although warm autumns and the lack of storms may prolong the activity season into November. Year-­round activity has been reported at a warm spring (15°C) in northwestern Utah (Thompson, 2004) and in lowland parts of their range in California (Storer, 1925). Activity may occur at any time of the day, with peaks at midday and ­after dark (­Sullivan et al., 2008; Long and Prepas, 2012). Toads frequently move short distances (e.g., within 15 m of refugia at night; Long and Prepas, 2012), and ­will orient themselves to bask in the warm sun. Adults have a somewhat biphasic daily activity period in early spring at high elevations. On cold spring mornings, Western Toads ­will sit at the entrances to cover sites and warm themselves by basking (Karlstrom, 1962). They remain active ­until the warmest part of the day (12:00–14:00), when they return to cover ­under loose soil, in mammal burrows, or ­under surface debris. Toads ­will reemerge in the late after­noon and remain active ­until the dry mountain air cools. This pattern is repeated in the autumn as daily temperatures become cooler. Activity occurs at temperatures as low as 3°C as toads seek shelter for the night, and toads routinely are active at temperatures 200 m/day (Bull, 2006) and travel >400 m in a single day (Bartelt et al., 2004); Schmetterling and Young (2008) recorded median movements of 152–162 m/day, although most movements ­were 1 km within a stream channel over a 42 day period (Griffin, 1999). Griffin (1999) also noted upstream dispersal prior to the breeding season by females. The lit­er­a­ture suggests that Arroyo Toads rarely disperse away from the stream margins farther than adjacent upland terraces in narrow canyon habitats, but this may not be so everywhere. For example, Holland and Sisk (in Lovich, 2009) noted that Arroyo Toads have been found in uplands 1,175 m from riparian habitats along a tributary of San Mateo Creek in San Diego County. In coastal localities, they may move more extensively overland (to 1.2 km) into upland grasslands and sage scrub (in Sweet and ­Sullivan, 2005; but possibly the same movement reported by Lovich, 2009). Except during the breeding season, activity is largely nocturnal from January to early August, when body temperatures (100 m distant from summer activity areas (Griffin, 1999). During the day and in winter, adults seek refuge in root channels, rodent burrows, along stream terraces, or in moist areas associated with the under­ground passage of ­water; burrows may be horizontal or vertical (Griffin, 1999). Many individuals can occupy a single refugium. Cunningham (1955c) reported individuals buried in sand up to 45.7 cm below the surface. Thunderstorms may bring them rapidly to the surface. Locomotion is by hopping rather than walking, as in many toads.

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Anaxyrus californicus 63

­After metamorphosis, froglets remain very near ­water for a week, then move to dryer sand bars for about 3–8 weeks; ­there, they hide in vegetation and ­under surface debris. Juveniles and newly metamorphosed toadlets are diurnal but ­will take refuge in existing burrows during the hottest part of the day. Their body temperatures can reach 26–37°C. Small juveniles often sit in the direct sun even on extremely hot substrates without apparent distress; evaporative cooling may assist in keeping body temperatures lower than the surrounding environmental temperatures. At about 30 mm SUL, dispersal begins to areas dominated by willows; it is at this size that they become capable of burrowing themselves. They ­will burrow 10–18 cm below the surface and remain inactive ­there for 6–8 months (Griffin, 1999; Sweet and ­Sullivan, 2005). By August, juveniles assume a nocturnal activity pattern. CALLING ACTIVITY AND MATE SE­L ECTION

Males emerge from overwintering sites prior to females. Calling may begin in late March and extend to late July. The call of A. californicus has been described as a “sweet trill” (Myers, 1930b) or as a clear, prolonged musical trill (Stebbins, 1962). The call has a dominant frequency of ca. 1.42 kHz, a mean pulse rate of 41.5 pulses/sec and a mean call duration of 6.62 (range 2–14) sec (Stebbins, 1962; ­Sullivan, 1992b). Males also have a warning call that is somewhat like the musical trill of the advertisement call. Males position themselves along pools and call from exposed locations where they display a high level of site fidelity. From 1 to 3 males may call from a pool on any one night (S. Sweet, in Jennings et al., 1994b). Calling ceases when temperatures are ca. 13°C and avoid ambient temperatures >28–30°C. They are able to survive a mild freeze, presumably by converting body ­water to ice crystals. According to Karlstrom (1962), they show no par­tic­u­lar thermal preferences, and are likely to be active anytime during normal ambient temperature conditions for this region and season (2–30°C). Yosemite Toads are diurnal. During terrestrial movements and at night during cold temperatures, they shelter in bur­ nder rocks, logs, and other coarse woody debris, in rows, u tree stumps, or in rodent (Marmota flaviventris, Microtus montanus, Spermophilus beldingi, Thomomys monticola) burrows several cm below the surface (Liang, 2013). Such sites often have less canopy cover and fewer woody species than other potential sites. Overwintering also occurs in such sites, as well as in cracks and crevices in rocks beneath the soil or among the roots of vegetation. When their body temperature reaches 8–10°C, the toads become active. Toads are commonly seen sitting at the entrance to their shelters, where they may bask for 30–60 minutes before leaving, especially at lower temperatures (Mullally and Cunningham, 1956a). Basking serves to warm the toad as it begins its daily activity, and toads may achieve body temperatures of 18°C despite the morning chill. Karlstrom (1962) also noted crepuscular and nocturnal activity, especially during the breeding season. Males tend to remain in the vicinity of the breeding pools, whereas females disperse more widely ­after their eggs are deposited. Mullally (1953) also noted that many Yosemite Toads stay near watercourses where they feed as they move several meters up and down the shoreline. Still, movements away from ­water can be extensive. For example, Morton and Pereyra (2010) noted that females tended to move to the vicinity of a talus slope 800 m from the nearest breeding pond, but that males did not. Liang (2013) tracked adults to a mean of 270 m away from aquatic breeding sites; females (maximum distance traveled 1.261 km) moved farther than males (to 865 m) and had larger home ranges. ­Others (Karlstrom, 1962; Kagarise Sherman, 1980; Kagarise Sherman and Morton, 1984; Morton and Pereyra, 2010) noted movement distances of 150–850 m away from ­water, and Martin (2008) recorded a maximum distance from ­water of 657 m (mean distance 279 m) in the northern part of the species’ range. This variation may reflect differences in surrounding habitats and available microclimates. Activity continues throughout the summer as late as September–­early October. As with A. boreas, Yosemite Toads likely are photopositive and sensitive to light in the blue spectrum (‘blue-­mode response’). As such, they prob­ably have true color vision.

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68  Bufonidae

CALLING ACTIVITY AND MATE SE­L ECTION

Toads emerge from dormancy in late spring to early summer depending on weather conditions and when the pond ice melts. Kagarise Sherman (1980) reported first emergences from 29 April to 4 June, depending upon year. They do not dig out from ­under snow, but must wait for it to melt from the entrance to their winter dormancy site. Emergence likely occurs ­later at higher elevations as snow and ice melts allowing access to the breeding ponds. However, even during the breeding season, temperatures may fall below freezing, leaving a thin crust of ice on the breeding ponds. ­There are 4–8 days between emergence and arrival at breeding sites and the first deposition of eggs (Karlstrom, 1962; Kagarise Sherman, 1980). Calling may begin sporadically at temperatures of 1–2°C, with full choruses trilling at temperatures of 11–12°C and especially >14°C. Chorusing peaks at midday and in the after­noon, but Karlstrom (1962) recorded calling even at near-­freezing temperatures at 21:50 hrs. Calling usually ends by mid-­June, but in years of ­later emergence, calling continues to mid-­July (Kagarise Sherman, 1980). Unlike the closely related A. boreas, the male’s voice is a long melodious trill. The trill consists of a series of 26–51 (mean 38) evenly spaced notes lasting from 0.8 to 3.8 sec (mean 2.6). ­There are 6 note harmonics, with a frequency range to 8,000 cps and a dominant frequency of 1,550 cps at 24–25°C (Karlstrom, 1962). ­There is the potential for sexual se­lection based on size, inasmuch as larger males have a voice with lower tones, and larger males do seem to be more successful at mating than smaller males. Males may call in close proximity to one another, or space themselves at considerable distances along a wetland margin. At least during the day, they tend to remain at the same calling station, although they may move about searching for

Amplexing adults and egg strings of Anaxyrus canorus. Photo: Earl Gonsolin

Dodd_Canada_int_5pgs_B1&B2.indd 68

females. Indeed, ­silent males tend to search near calling males and thus attempt to intercept a female approaching the caller. Such satellite male be­hav­ior is not uncommon in the Anura. ­There is some support for the hypothesis that in low density situations, males tend to call more than search. As the density of calling males increases to a certain level, some males may alter their tactics by ceasing to call and trying to intercept a female moving ­toward a calling male (Kagarise Sherman and Morton, 1984). Calling occurs from the shallow ­water along the margins of a pond or stream. Male–­male aggression (fighting, wrestling, making a low-­volume clucking sound) occurs should a nearby male approach a calling male. Male territoriality seems directed ­toward the protection of adequate space in which to intercept a female rather than a favorable oviposition site (Kagarise Sherman and Morton, 1984). Aggressive bouts may only last 20 sec, but aggressive encounters may last 5 min. Large males tend to win more fights than smaller males, which usually give up and return to from whence they came. The calling male rears up in a nearly vertical position, and the vocal sac is prominent as he trills. Chorusing by 1 male tends to trigger additional males to begin calling. Calls last for several seconds, and intervals between calls can last several minutes. Calls have good carry­ing capacity and may be heard >30 m away. According to Karlstrom (1962), chorusing may be more or less continuous among toads occupying a large meadow. Calling males are easily silenced if they are approached at night or as a result of loud noises, such as traffic. During the day, however, they are less easily disturbed. Males recognize females on the basis of their girth (full of eggs) and the fact that they are ­silent if contacted. The dichromatic coloration does not appear to play a part in sex recognition. Amplexus is pectoral. Males ­will attempt to amplex any moving object in their vicinity; if another male is contacted, he ­will emit his “warning vibration” and utter a release cluck. This alerts the amplexing male as to the sex of the animal he has clasped. Males also actively physically discourage amplexus by other males through pushing and kicking. Despite ­these efforts, balls of toads have been observed during peak mating periods as excited males vie for females and desperately grasp any object or multiple objects within reach (Kagarise Sherman and Morton, 1984). Females may have some choice in terms of males. A female may approach a calling male directly and actually touch him, and initiate amplexus. Females are known to move among calling males as if seeking a specific caller. Female choice seems to be effective in about 50% of the observed cases, with the remainder subject to random male interception. The males do not seem to be particularly selective in their choice

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Anaxyrus canorus 69

of females. Females also have been observed attacking amplectant pairs trying to dislodge the male and so obtain a mate (Maier, 2018b). BREEDING SITES

Optimal breeding sites include alpine lakes and pools of clear and cold shallow ­water surrounded by meadows (Fellers et al., 2015). Yosemite Toads also breed in shallow, slow-­moving runoff streams. They initially prefer breeding sites near the forest, as forested habitat provides cover during the still cold spring nights experienced immediately ­after emergence. As the temperatures increase, they move farther into the meadow and occupy the borders of wetlands and streams as the forested areas become drier. In meadow pools, occupancy is associated with several microclimate variables, specifically mean ­water depth (4.35 cm), mean ­water temperature (24.8°C), the percentage of surface ­water in pools (60%), mean detritus depth (1.62 cm), and mean vegetative height (13.2 cm) (Liang et al., 2017). The presence of nonnative trout does not appear to have an effect on ­whether A. canorus occupies a breeding site (Knapp, 2005; Fellers et al., 2015). In a comprehensive monitoring study, Brown et al. (2012) found Yosemite Toads breeding in an estimated 84% of recently occupied watersheds, but in only 13% of historically occupied watersheds. Of all 134 watersheds sampled, toads bred in only 22% of them. REPRODUCTION

As might be expected, the date of breeding is dependent upon elevation and local environmental conditions, with breeding commencing ­later at higher elevations. Using museum specimens, Goldberg (2021b) found spermiogenesis occurring from July to October, with mature ovarian oocytes

Egg strings of Anaxyrus canorus. Photo: Marty Morton and Maria Pereyra

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in July and October (no animals ­were available from other months). This suggests that some vitellogenesis occurs in autumn, in contrast to Morton and Pereyra (2010) who observed that ovaries of some postbreeding females ­were only partially developed before overwintering. Breeding normally occurs from early May to mid-­June in the high Sierras as soon as snow and ice melt at the nearly alpine breeding pools. Timing is impor­tant, since freezing severity and ice duration may cause significant embryonic mortality (Sadinski et al., 2020). Interestingly, ice slush ­causes increased mortality, although some embryos may survive 30–60 min encased in ice. Kagarise Sherman and Morton (1984) noted that breeding may not commence ­until as late as 20 June following winters with exceptional snowfall, and Wiggins (1943) and Mullally (1953) recorded gravid females on 11 August and 18 August, respectively. Sadinski et al. (2020) noted breeding from May to July soon ­after snow melted, which varied, as might be expected, by elevation. Even spring storms can interrupt a breeding season for weeks ­after it has started. Yosemite Toads routinely move to breeding sites when snow still covers much of the ground, as it does for 8–9 months of the year, often tiptoeing across snow patches (Kagarise Sherman and Morton, 1984). Egg deposition occurs in shallow ­water (7–8 cm) along a wetland or stream margin (Sadinski et al., 2020). Preferred sites have a silt bottom that allows the mated pair to swim about leaving egg strings anchored among sedges. Eggs are deposited over the sediment or detritus surface over an area of 1–2 m2, and a single egg clutch produces 1,500–2,000 larvae (Karlstrom, 1962). Hatching occurs at 5–12 days ­after oviposition, as the sun speeds up development of the eggs despite the cold ­water temperatures. A female only produces 1 clutch per year and may skip years between reproductive bouts in order to obtain sufficient energy to provide yolk for a clutch (Kagarise Sherman, 1980; Kagarise Sherman and Morton, 1984). Most oviposition occurs during the after­noon and lasts only long enough for females to extrude their eggs and males to fertilize them. Amplexed pairs may be seen moving about a shallow wetland (perhaps for a few meters) seeking an oviposition site, and several mated pairs may deposit their eggs in close proximity to one another. ­After amplexus, the adults seek shelter for the night. ­There appear to be many more males than females in attendance at any one time at a chorus. For example, Karlstrom (1962) stated that ­there ­were 10 males per female at the chorus he studied at Tioga Pass, California. However, Kagarise Sherman and Morton (1984) noted that the sex ratio may change through time at a site (6.3 males per female to 1 male per 2 females at their site over a period of

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6 yrs), and that the degree of polygyny varies accordingly. Females presumably come into a breeding site, are immediately amplexed, deposit their eggs, and leave. Males, on the other hand, likely remain chorusing at the breeding site for 7–17 days in order to intercept as many females as pos­si­ble. Male success appears to be tied to the duration of his stay and when he arrives; thus, males arriving ­earlier and staying longer have a better chance of successful reproduction than do ­later arrivals (Kagarise Sherman and Morton, 1984). Males do not remain throughout the breeding period, inasmuch as the short activity season necessitates maximum foraging in order to survive the long winter. Males may mate with more than 1 female (to 3), or not be successful at all. Male mating success was zero 58.7–88.3% of the time in Kagarise Sherman’s (1980) 4 yr study, whereas 10.5–32% of males mated with 1 female, 1.2–9.3% mated with 2 females, and only 0.5% mated with 3 females and that in only 1 yr. Females, on the other hand, mate with only a single male annually and some may skip breeding seasons. ­Because of the variance in mating success, the operational sex ratio is actually 10–38 males per female (Kagarise Sherman, 1980). LARVAL ECOLOGY

Eggs are deposited in shallow w ­ ater that provides a positive thermal environment for embryonic development. Karlstrom (1962) considered the primary high-­elevation reproductive adaptation of A. canorus to be the placement of eggs where they could be warmed through shallow-­water heat insolation. The sun’s rays warm the eggs and accelerate development despite the overall cold ­water and air temperatures early in the season. Larvae tend to remain in the shallow warm ­waters during the day, but retreat to deeper ­water ­after dark (Mullally, 1953). Presumably the deeper ­water holds heat longer than the shallow ­waters along the shoreline. Hatching success varies annually and by site, with most embryonic mortality occurring early in development (Sadinski

et al., 2020). Significant larval mortality occurs if w ­ ater temperatures reach 31°C. The longer larvae remain within the breeding pond, the greater the temperatures to which they are able to acclimate. Thus, tadpoles easily survive ­waters at 32°C during the late summer stages of development. The larval CTmax is 36–38°C (Karlstrom, 1962). Still, ambient temperatures rarely get this high, especially in the generally cool high elevations inhabited by the Yosemite Toad. The large amount of yolk in A. canorus eggs allows the larvae to grow to 9–10 mm body length (26 mm TL) prior to feeding. Thus, larvae are able to attain one-­third of their final body size even before feeding begins. Such parental investment ensures the likelihood of successful metamorphosis despite the extreme high-­elevation mountain conditions. In laboratory ­trials, reducing the hydroperiod results in an acceleration of time to metamorphosis, but tadpole size (snout-­urostyle length, body mass) did not differ between slow-­drying and fast-­drying mesocosms (Lindauer and Voyles, 2019). Metamorphosis occurs in 40–50 days in the laboratory (e.g., Jennings and Hayes, 1994b) and 56–110 days in nature, depending upon thermal conditions. Metamorphs are 8.4–13.2 mm SUL. ­Under laboratory conditions, metamorphs had a mean SUL of 12.9 mm and a body mass of 0.37 g (Lindauer and Voyles, 2019). Aside from insect predators, the greatest threat to larval Yosemite Toads is desiccation as ponds and wetlands dry during the summer (Jennings and Hayes, 1994b). Ponds are more likely to dry when the snowpack is shallow and no rain falls during the spring. If the previous winter’s snowfall was heavy, ponds are likely to be deeper and hold ­water longer, which allows for metamorphosis to occur. Exceptionally dry springs and summers cause ponds to draw down quickly. Larval A. canorus do not exhibit any changes in be­hav­ior when exposed to chemical stimuli from the nonnative brook trout Salvelinus fontinalis (Grasso et al., 2010). POPULATION BIOLOGY

Tadpole of Anaxyrus canorus. Photo: Marty Morton and Maria Pereyra

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­ fter metamorphosis, toadlets must grow quickly in order to A obtain sufficient body reserves to last their first very long winter. Sexual maturity is reached in about 3–5 yrs in males and 4–6 yrs in females (Kagarise Sherman, 1980; Kagarise Sherman and Morton, 1984; note that Karlstrom, 1962, was incorrect in his speculation that maturity was reached in 2–3 yrs). Goldberg (2021b) noted mature males at 38 mm SUL and females at 44 mm SUL. Sex ratios are unknown, but Morton and Pereyra (2010) recorded 237 females and 225 males over a 7 yr period at Yosemite National Park. Morton and Pereyra (2010) further suggested that the cold weather and short activity period results in an irregular breeding cycle for this species, since females must acquire energy in a rather short period of time

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(Salvelinus fontinalis) (Grasso et al., 2010). Egg masses are preyed upon by an as yet unidentified flatworm (Turbellaria sp.; Sadinski et al., 2020). Larvae are preyed upon by dragonfly naiads and predaceous diving beetles. Birds eat larvae that are stranded by lowering ­water levels. Adult and subadult Rana sierrae and R. muscosa also feed on larval Yosemite Toads (Mullally, 1953; Pope and Matthews, 2002). Adults are preyed upon by California gulls (Larus californicus) and Clark’s nutcrackers (Nucifraga columbiana). Adults are particularly vulnerable as they cross snowfields. Adults have a noxious secretion produced by the parotoid and granular glands on the dorsum that may aid in their protection. Female adult Anaxyrus canorus. Photo: Ceal Klingler

for egg production and to survive dormancy. Yosemite Toads may have long lives; Kagarise Sherman and Morton (1984) recorded maximum female longevity of at least 15 yrs, and males of 12 yrs. In a comprehensive assessment of the status of this species in multiple watersheds in the Sierra Nevada, Brown et al. (2012) found that Yosemite Toad populations ­were small within watersheds, with the largest 2 populations having only 16–21 and 18–19 breeding males per year. Male survivorship was 0.49–0.72. Few egg masses ­were oviposited per year, and the proportion of breeding areas producing successful metamorphs varied from 0.14 to 0.73. Within the watersheds Brown et al. (2012) examined, Yosemite Toads tended to breed at only 2 sites ­every year, and only occasionally at other sites. In addition to avian predators, Kagarise Sherman (1980) reported mortality from multiple amplexus, exposure, unidentified illness or pathogens (possibly Aeromonas), and from trampling by large mammals. She also found dead toads with no apparent injury or cause of death. DIET

Yosemite Toads are ambush predators, eating a variety of invertebrates, including ants, bees, wasps, beetles, millipedes, flies, mosquitoes, lepidopteran larvae, dragonfly larvae, and spiders (Mullally, 1953; Wood, 1977). Males tend to reduce feeding activities during the breeding season (Wood, 1977). Larvae have been observed scavenging on dead conspecific and heterospecific tadpoles and adults, fish, beetle larvae, and mammals (Maier, 2019). PREDATION AND DEFENSE

Eggs, larvae, and postmetamorphic A. canorus are unpalatable to some predators, such as the nonnative brook trout

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DISEASES, PARASITES, AND MALFORMATIONS

Egg masses and embryos experience significant mortality from the fungus Saprolegnia diclina (Sadinski et al., 2020). Kagarise Sherman (1980) reported mortality from unspecified illness or pathogens, and speculated that red-­leg (the bacterium Aeromonas) might be responsible. However, amphibian chytrid (Bd) was detected in 23–30% of toads sampled in Yosemite National Park (Fellers et al., 2011) and is now thought to be a major ­factor in this species’ decline in distribution and abundance. Recently metamorphosed A. canorus are highly susceptible to chytrid infection, but ­there does not appear to be a residual effect of disease risk due to drought (Lindauer and Voyles, 2019). Yosemite Toads are parasitized by the cestode Cylindrotaenia americana and the nematode Cosmocercoides variabilis (Walton, 1941). SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. Survival of embryos and hatchling Yosemite Toads is affected by low pH. In experimental ­trials, Bradford et al. (1992) estimated the LC50(7 day) for embryos as 4.7 and for hatchlings as 4.3. Survival was normal at a pH of 5.0. However, a pH of 5.0 caused ­earlier hatching than normal. Metals. Exposure of embryos and larvae of aluminum at 39–80 μg/L had no effect on survivorship (Bradford et al., 1992). However, sublethal effects included a reduced body size in tadpoles and an ­earlier hatching by embryos. UVB radiation. UVB radiation does not affect hatching success or developmental rates of embryos (Vredenburg et al., 2010a; Sadinski et al., 2020). STATUS AND CONSERVATION

The Yosemite Toad has declined dramatically (by 50%) over the last 30 years, despite protection of habitat within large national parks (Martin, 1991; Kagarise Sherman and Morton, 1993; Drost and Fellers, 1996; Stebbins and Cohen, 1995; Corn, 2003; Morton and Pereyra, 2010). Even in surviving populations, abundance has decreased

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Habitat of Anaxyrus canorus. Photo: Gary Fellers

substantially. In times of drought, egg deposition sites may not be available, or larvae may not have sufficient time to complete metamorphosis (Myers, 1930a; Karlstrom, 1962). At such times, substantial mortality may occur and set back recruitment considerably. Another potential killing agent is winterkill, especially in years where snowpack—­which serves to insulate the subsurface retreat sites—is low. Drought and winterkill over several seasons could selectively target early arriving males or their offspring, and thus alter the breeding

Anaxyrus cognatus (Say, 1822) ­Great Plains Toad Crapaud des Grandes Plaines ETYMOLOGY

cognatus: from the Latin word cognatus, meaning ‘related by birth.’ NOMENCLATURE

Conant and Collins (1998) and Stebbins (2003): Bufo cognatus Fouquette and Dubois (2014): Bufo (Anaxyrus) cognatus Synonyms: Bufo dipternus, Bufo lentiginosus cognatus, Chilophryne cognata, Incilius cognatus The species Anaxyrus californicus was originally described as Bufo cognatus californicus (Camp, 1915).

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sex ratio in subsequent years (Kagarise Sherman and Morton, 1984). The potential for climate change (drought, alteration of rainfall patterns) to lead to reductions or even the extinction of this species is a real possibility. Given the extensive ge­ne­tic structuring among populations based on breeding sites and drainages (Shaffer et al., 2000; Maier, 2018a), this species must be conserved on a population-­by-­population basis. ­Every population becomes impor­tant, and the survival of the species ­will not be ensured ­unless each population receives specific attention. Maier (2018a) identified 3 “Germinate Evolutionary Units” in Yosemite National Park that ­will likely become refugia for this species as the effects of climate change drastically alter the environment of this high-­ elevation species. ­These are areas of critical importance to the survival of the species. Although acidification has been suggested as a pos­si­ble ­factor influencing population decline, ­there is no empirical support for the hypothesis (Bradford et al., 1994a). Grasso et al. (2010) suggested that removal of nonnative trout from Yosemite Toad habitats would have no effect on this species ­because the toad is unpalatable. Likewise, Matchett et al. (2015) could not find evidence of a negative effect of pack animals on breeding by this species, although this could be a minor prob­lem in Yosemite. The Yosemite Toad is protected by the National Park Ser­vice and the State of California as Endangered. The toad is protected as Endangered ­under provisions of the US Endangered Species Act of 1973.

IDENTIFICATION

Adults. ­Great Plains Toads are medium to large in size. The ground color is olive to pale gray brown, and ­there are large, well-­defined blotches of olive, dark green or dusky coloration over the dorsum. ­These blotches are bordered with a light tan, brown, or green ring of color. Darker individuals have an obvious, light mid-­dorsal stripe that may not be apparent in light individuals. The entire dorsum is covered with small, warty bumps. The prominent cranial crests and parotoids are large and together form an “L” shape that comes together to form a “V” ­shaped osseous boss (a bump) between or ­behind the nares; the boss is not well developed ­until the toad is ca. 37 mm SUL. Parotoids are large and oval ­shaped and are contacted by the postorbital crests. The bellies are white, usually with no blotches or spots, but rarely spotted. Juveniles, but not adults, have small, brick-­ red tubercles. The rear feet have prominent metatarsal tubercles that aid in digging. Males have a dark, loose-­ skinned throat that forms an apron in appearance, and

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tion of larvae during development. They usually reach 29–30 mm TL prior to transformation. Eggs. The small eggs (about 1.2 mm in dia­meter) are black dorsally and lighter ventrally, although the distribution of the pigment makes the eggs appear dark all over. They are enclosed in a gelatinous capsule within an extended string of tough, elastic gelatin, and each egg is separated from its neighbor by a partition. Rarely are ­there 2 rows of eggs within a single string (Bragg, 1936). Egg strings may be single or double. Two scalloped envelopes surround the vitellus, the outer envelope averaging 1.7–2.1 mm and the inner envelope averaging 1.6 mm (Bragg, 1937b; Livezey and Wright, 1947). DISTRIBUTION Head pattern of Anaxyrus cognatus. Illustration by Camila Pizano

cornified thumbs. When the vocal sac is extended, it is sausage ­shaped and juts up over the snout. Melanistic juveniles ­were reported by Bragg (1958a). Although occasional individuals are >100 mm SUL, most ­Great Plains Toads are smaller than this. Males are on average smaller than females. In Oklahoma, calling males ­were 72–90 mm SUL (mean 80 mm) (Bragg, 1940c); Bragg (1950e) ­later reported Oklahoma males at 62–103 mm SUL (mean 79.4 mm) and females at 49–112 mm SUL (mean 85.8 mm). In Krupa’s (1994) study, the mean size of males was 69.5–79.8 mm SUL (range 56–98 mm), and females ­were 71.8–85 mm SUL (range 60–115 mm). Nebraska adults averaged 65.1 mm SUL (range 52–78 mm) (Ballinger et al., 2010). In Arizona, males ­were 68.8 mm in mean SUL, whereas females ­were 72.8 mm SUL (­Sullivan and Fernandez, 1999). Based on museum specimens, Goldberg (2018e) recorded Arizona males from 50 to 73 mm SUL (mean 65.6 mm) and females from 61 to 84 mm SUL (mean 73.2 mm). Wright and Wright (1949) reported males as 47–95 mm SUL and females as 60–99 mm SUL. Smith (1934) recorded a 114 mm SUL female in Kansas. Length is directly correlated with body mass. Larvae. At hatching, the tadpoles are black with the ventral coloration lighter than the dorsal coloration. As the tadpoles grow larger, they become grayer and mottled brown with silver chromatophores that may be pre­sent throughout the dorsum. Venters are gray, and ­there are golden chromatophores scattered throughout the belly. Viscera may be seen through the belly early in development but become obscure as the tadpole grows. Eyes are large with a golden iris. Tail fins are highly arched and contain scattered dendritic pigment patterns on the dorsal fins. Ventral tail fins are clear. Bragg (1936) provided detailed illustrations and a descrip-

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As its common name implies, this is a grassland and semi-­ arid species that occurs throughout the ­Great Plains from southeastern Alberta, southern Saskatchewan, and Manitoba, southward through north and west Texas, then south to Aguascalientes and San Luis Potosi, México. It occurs in the semi-­arid west through intermediate elevations in southeastern Utah (Green River drainage) and northeastern Arizona (in 3 isolated areas), the Sonoran Desert in southern Arizona, the Mojave Desert and Imperial and Coachella Valleys of southeastern California, and much of New Mexico (excluding the southern Rockies), especially along the Rio Grande. The species is absent from the Rocky Mountains, the Black Hills, and from the high mountains of

Distribution of Anaxyrus cognatus

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central Arizona and west and central New Mexico. Populations occur in southern Colorado, but rec­ords north of the ­Great Salt Lake in Utah are likely in error (Mulcahy et al., 2002). ­There may be other populations in the grasslands south of the ­Grand Canyon and perhaps elsewhere in northeastern Arizona and southeast Utah, but fieldwork has yet to be carried out in ­these regions. The range extends to its farthest point east along the Missouri River into central Missouri. Impor­tant distributional references include: Alberta (Russell and Bauer, 2000; Pearson, 2009), Arizona (Brennan and Holycross, 2006; Murphy, 2019; Holycross et al., 2021), California (Vitt and Ohmart, 1978; Goodward and Wilcox, 2019; Flaxington, 2021), Colorado (Hammerson, 1999; Whalen, 2021), Kansas (Smith, 1934; Collins, 1993; Collins et al., 2010), México (López et al., 2009), Montana (Black, 1970, 1971), Minnesota (Oldfield and Moriarty, 1994), Missouri (Johnson, 2000; Daniel and Edmond, 2006), Nebraska (Lynch, 1985; Ballinger et al., 2010; Fogell, 2010), Nevada (Linsdale, 1940), New Mexico (Degenhardt et al., 1996), North Dakota (Wheeler and Wheeler, 1966; Hoberg and Gause, 1992), Oklahoma (Sievert and Sievert, 2006), Saskatchewan (Cook, 1960; Heisler et al., 2013), South Dakota (Fischer, 1998; Ballinger et al., 2000; Kiesow, 2006), Texas (Dixon, 2000; Tipton et al., 2012; Davis and LaDuc, 2018), Utah (Tanner, 1931; Mulcahy et al., 2002), and Wyoming (Baxter and Stone, 1985). FOSSIL REC­O RD

This species is known from many fossil deposits from the Miocene (Kansas, Nebraska), Pliocene (Kansas, Nebraska, Texas) and Pleistocene (Colorado, Kansas, Sonora, Texas) (Rogers et al., 1985; Rogers, 1987; Holman, 2003). In the fossil lit­er­a­ture, bones of this species are often referred to as Bufo cf. B. cognatus. Tihen (1962b) and Holman (2003) gave identifying characters, such as its distinctive frontoparietals, but noted that the ilia may be confused with ­those of other species. SYSTEMATICS AND GEOGRAPHIC VARIATION

The ­Great Plains Toad is a member of the Cognatus clade of North American bufonids. This clade also includes A. compactilis of México and A. speciosus, but Masta et al. (2002) have shown that the clade is paraphyletic. The Cognatus clade is a ­sister group to the larger Americanus clade of toads. Rogers (1972, 1973) provided information on morphological and ge­ne­tic variation in the species. A single hybrid A. cognatus x A. hemiophrys was found in Minnesota (Brown and Ewert, 1971), and a single A. cognatus x A. woodhousii hybrid was reported from Arizona (Gergus et al., 1999). Hybridization has been achieved in

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laboratory crosses between A. cognatus and A. americanus, A. boreas, A. debilis, A. punctatus, A. terrestris, A. woodhousii, Incilius alvarius, and I. nebulifer (Moore, 1955; Blair, 1959, 1972). Artificial crosses may produce few larvae, however, with A. americanus, A. quercicus, or A. punctatus (Blair, 1972). Chihuahuan Desert populations of A. cognatus exhibit ­little between-­population ge­ne­tic variation, and most variation occurs within populations (Jungels et al., 2010). It appears that A. cognatus uses river corridors for dispersal thus reducing ge­ne­tic differentiation among distant populations. ADULT HABITAT

Anaxyrus cognatus is a species mostly of the dry short-­grass or mixed-­grass prairies of the ­Great Plains and the arid and semi-­arid deserts and grasslands of the American Southwest (Bragg, 1940c, 1950b; Bragg and Smith, 1943; Boeing et al., 2014). Both tall-­grass and short-­grass prairies are occupied, as are mesquite (Prosopis sp.) grasslands, riparian areas in deserts, desert scrub, creosote bush (Larrea tridentata) desert, sagebrush (Artemisia tridentata) plains, pine forests, and mesquite woodlands. As one proceeds westward in the short-­grass prairies, the species becomes more restricted to riparian habitats. In the eastern part of its range, it ­favors prairie uplands where it is associated with loess hills and heavy floodplain soils (Timken and Dunlap, 1965). In the ­Great Plains, it is especially associated with sand plains and sandhill habitats that are necessary so that the toads can dig below the frost line. This species is frequently encountered in cultivated and agricultural fields, where deep-­water wetlands imbedded in croplands are vital to its survival (Mushet et al., 2012). ­Great Plains Toads are found to 1,900 m in New Mexico (Degenhardt et al., 1996). Within this extensive region, the species is subject to extreme variation in temperature, from very hot summers to very cold winters. Precipitation is often scant and spottily distributed and can vary greatly from year to year. This species occupies about 65% of potential breeding sites in west Texas, regardless of ­whether the adjacent land is in crops or open range (Anderson et al., 1999). Occupancy of a site is positively correlated with the presence of adjacent nearby wetlands, but landscape structure per se does not have much influence on this species (Gray et al., 2004). This species occasionally may be found in or near cave entrances (Black, 1973a; Collins, 1993). TERRESTRIAL ECOLOGY

Activity occurs throughout the warm season. In Alberta, this extends from April through September (Pearson, 2009), whereas in Oklahoma it extends from March through September. Most activity occurs at night or during overcast

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weather, but they may be active diurnally ­after heavy rain. During the day, they remain dormant in shallow (131 m, with the longest being 808 m. Movements occur only sporadically during the late summer hot and dry season when environmental conditions are not optimal. Then, toads go into a period of dormancy in protected refugia or emerge only at night, and many do not emerge at all even during periods of favorable rainfall (Bragg, 1945a). They burrow backward into the soil and rest alert below the surface, or they may occupy cracks in mud. A light rain may bring them back to the surface, and they can readily rehydrate from soil moisture alone (Walker and Whitford, 1970). Long-­distance movements frequently occur between breeding ponds, foraging areas, and overwintering sites, usually at night. ­Great Plains Toads can travel from summer foraging areas to winter dormancy sites over a considerable distance. For example, Ewert (1969) noted a mean movement distance of 229 m (range 100–1,036 m) between release points at foraging areas and winter dormancy sites. During long-­distance movements, toads may use man-­made landmarks, such as roads or the borders of agricultural fields, as dispersal corridors. Juveniles can disperse over considerable distances ­after metamorphosis. For example, Ewert (1969) found toadlets >900 m from the nearest breeding pond. Juvenile dispersal occurs during mass movement events when literally tens of thousands of small toads (30–52 mm SUL) may be observed moving unidirectionally, often north (Breckenridge, 1944; Bragg and Brooks, 1958). During dispersal, toadlets often stop to feed for a short time, but movements are steady and occur even during the heat of the day. ­These mass movements do not appear correlated with humidity, moonlight, or the opportunistic presence of smooth roads offering few obstacles to travel (Smith and Bragg, 1949). Dense cover is preferred as a microhabitat in choice tests, although the species does not avoid open habitats (Tester et al., 1965). Foraging mode may change in relation to vegetative cover, with juvenile toads in dense vegetation adopting a sit-­and-­wait foraging strategy, while ­those in open vegetation actively pursue prey (Flowers, 1994). Anaxyrus cognatus does not appear to regulate its body temperature to any extent by selecting a par­tic­u­lar density of

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vegetation inasmuch as its body temperature is very similar to ambient temperature in experimental ­trials. It also does not seem to actively avoid high temperatures (Tester et al., 1965). Indeed, this species prefers high temperatures (31°C), even more so than some tropical toads (Sievert, 1991). Postmetamorphic juveniles often aggregate among conspecifics in terrestrial situations; aggregations consist of 4–7 toadlets. This tendency occurs during daylight hours but not at night, and therefore suggests a role for vision in the formation of juvenile aggregations. In addition, toadlets aggregate in areas previously occupied by conspecifics, suggesting a role for chemoreception in their spatial distribution. The aggregations are not formed in response to optimal feeding areas or to certain preferred habitat structure, and perhaps they serve as an antipredator function (Graves et al., 1993). Anaxyrus cognatus is often found in very cold environments, but it is not tolerant of freezing temperatures and possesses no cryoprotectants (Swanson et al., 1996). During freezing, liver glucose levels may be elevated, but not enough to serve as a cryoprotectant. Swanson et al. (1996) suggested that ­Great Plains Toads might be able to supercool within protected burrows, but empirical support has not been forthcoming. Most activity occurs at temperatures >15°C (Hammerson, 1999), with mortality at 41–42°C (Paulson and Hutchison, 1987). ­Great Plains Toads take refuge from both hot and cold temperature extremes by burrowing deep within sandy soils. When first emerging ­after winter, toads remain in the winter burrow about a day before exiting, presumably resting, and then depart from the site of the winter burrow within 6 days. Summer retreats may be up to 55 cm below the surface during extended periods of hot and dry weather (Ewert, 1969). The summer burrow may be envisioned as an inverted question mark with the toad situated in the upper part of the short end (C.E. Burt in Tihen, 1937). Toads enter winter burrows from late summer to fall (e.g., August–­September in Minnesota; Ewert, 1969), usually at night. Summer burrows may be converted to winter dormancy burrows. In Minnesota, overwintering sites ­were located in raised areas along roads or in soil banks in the prairie; such areas frequently had a minimum of snow cover compared with A. americanus sites in the same general area. Winter retreats extend from 74 to 104 cm below the surface to ensure the toad is below the frost line (Graves and Krupa, 2005), but toads ­will move both horizontally and vertically within the burrow depending on conditions. A rising groundwater level in spring may kill A. cognatus. Anaxyrus cognatus is frequently found in mammal burrows, such as ­those of pocket gophers (Geomys bursarias, Thomomys talpoides) and prairie dogs (Cynomys ludovicianus), and in

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76  Bufonidae

old badger (Taxidea taxus) burrows (Ewert, 1969; Lomolino and Smith, 2004). CALLING ACTIVITY AND MATE SE­L ECTION

­ fter emergence, males move t­ oward the breeding sites. A Ewert (1969) noted that toads traveled >805 m to reach a par­tic­u­lar breeding site, and that a male might bypass an apparently suitable site with other calling males in order to reach a breeding pool. The toads appear to know where they are ­going, as movements are not random. Adults may alter their direction once movements have begun. Calling is closely tied to precipitation. Males arrive at breeding sites prior to females both seasonally and daily. Calling can begin from late after­noon to early eve­ning, but most calling begins ca. 30 min ­after dark. Choruses function both to stimulate females and to orient males to a breeding site. A few males begin to call, then more and more males arrive ­until a large number of toads is calling in full chorus. Breeding aggregations can be quite large, and males often return to the same breeding site from one year to the next. No breeding may take place during years of drought. If one pool has a large number of chorusing males and a nearby pool has a small number of males, the latter males ­will move to the larger chorus. By ca. 21:30, females arrive at the breeding site, and most breeding occurs within 1–3 hrs ­after sunset (Krupa, 1994). Calling starts to taper off ­after 1:00. Choruses have been heard at air temperatures of 8°C, but chorusing is initiated only at temperatures >12°C (Bragg, 1940c) with most favorable temperatures between 16.5 and 21°C (Ballinger et al., 2010). The mean body size of breeding adults increases at a pool as the season progresses, indicating that small toads arrive at breeding sites prior to larger toads (Krupa, 1994). The advertisement call of A. cognatus is an ear-­splitting trill that can be heard at least 2 km away. Standing next to a chorus of calling males is deafening. In Minnesota, the call has a dominant frequency of 2,250–2,725 cps, a duration of 5.0–42.7 sec, and a pulse rate of 16–18 pulses/sec (Blair, 1957). In Texas, the call has a dominant frequency of 1,775–2,075 cps, a duration of 10.4–53.8 sec, and a pulse rate of 13–15 pulses/sec (Blair, 1956b). The dominant frequency of the male’s call is negatively influenced by body length, but is unrelated to ­water temperature (Krupa, 1990); pulse rate (positively) and call duration (weakly negative), however, are influenced by temperature but not body length. The dominant frequency of the call may vary among individual males even over the course of a single night. Males call from many types of sites, although females prefer larger breeding sites and ­will bypass males calling from small or unsuitable locations. Even if a male from a small breeding site amplexes her, she ­will attempt escape or

Dodd_Canada_int_5pgs_B1&B2.indd 76

move to a larger site; eggs are rarely deposited in small, inappropriate locations. Calling occurs from around the borders of pools and from within the ­water of shallow (0.53 mg/L (LC50 2.45 mg/L) (Birge et al., 1980). Some of ­these chemicals had ­little effect on hatching, however. The LC50(4 day) for methylene chloride is >32 mg/L. The herbicide paraquat has an LC50 (96 hr) of 15 mg/L affecting Fowler’s Toad tadpoles (Mayer and Ellersieck,

Dodd_Canada_int_5pgs_B1&B2.indd 103

1986). Residues of the insecticide fenvalerate ­were found in A. fowleri at concentrations of 0.02 ppm in Arkansas, although the sample size was very low (Bennett et al., 1983). The insecticide had been applied 5 days previous to sampling. Sanders (1970) provided TL50 values for DDT administered to A. fowleri larvae from 24 to 96 hrs ­after exposure at ages of 1–7 weeks. ­These values decreased with age and time ­after exposure. For example, the TL50 for 1 week old larvae ­after 24 hrs of exposure was 5.3 mg/L. However, the TL50 for 7 week old larvae 96 hrs ­after exposure was only 0.03 mg/L. Exposure of larvae to DDT clearly was lethal to this species in small doses. Sanders (1970) provided additional TL50 values for a host of pesticides, including trifluralin, endrin, toxaphene, guthion, TDE, methoxychlor, heptachlor, dieldrin, DEF, malathion, aldrin, hydrothol 191, benzene hexachloride, lindane, silvex 2-(2,4,5-­T), molinate, and paraquat for 4–5-­week-­old larvae at 24–96 hrs ­after exposure. Lindane was the least toxic pesticide, whereas endrin was the most toxic. In another comparative study, Ferguson and Gilbert (1968) found that aldrin and dieldrin ­were less toxic to adult Fowler’s Toads ­after 36 hrs of exposure than ­were DDT and toxaphene in treated cotton fields. Toads residing in areas previously treated by the pesticides showed up to a 200-­fold increase in re­sis­tance compared with toads from untreated sites. STATUS AND CONSERVATION

Roads have helped in monitoring Fowler’s Toads populations, but they also have detrimental impacts, especially if they are located near breeding sites or on migratory pathways. On Cape Cod, for example, Fowler’s Toads ­were active on park roads ­whether or not it was raining during the postbreeding season (Timm and McGarigal, 2014).

Breeding habitat of Anaxyrus fowleri. Gastrophryne carolinensis also bred in this pond. ­Great Smoky Mountains National Park, Tennessee. Photo: C.K. Dodd, Jr.

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104  Bufonidae

Individual toads spent extended periods of time on the road and used the same portion of the road on multiple nights. Undoubtedly, traffic takes a considerable toll on toad populations throughout the toad’s range. Fowler’s Toads are easily monitored by listening for their calls on prescribed survey routes. Their loud call is easily detected at a considerable distance, and 5 min call listening protocols have proven nearly as effective as 10 min protocols in allowing researchers to detect this species (Burton et al., 2006). On the negative side, toads are frequent victims of highway-­related mortality and perhaps tens of thousands are killed annually (Ferguson, 1960; Campbell, 1969; Sutherland et al., 2010). Throughout much of its range, A. fowleri is a very common species. However, ­there is no doubt that populations of Fowler’s Toads have declined or been extirpated at many locations. This is not surprising, since much of its range occurs in the heavi­ly urbanized eastern United States and Canada. For example, Lazell (1976) notes its disappearance from Penikese, Muskeget, Cuttyhunk, Nantucket, and Tuckernuck islands in Mas­sa­chu­setts ­because of pesticide spraying from the 1940s to the 1960s. Davis and Menze (2000) suggested populations might be declining in Ohio, and Weir et al. (2014) did the same for the Northeast in general. In contrast, populations may be increasing in the South (Villena et al., 2016) and in Illinois (Florey and Mullin, 2005), and the toads are considered common in Indiana (Brodman and Kilmurry, 1998), Georgia, (Jensen et al., 2008) and Arkansas (Trauth et al., 2004). Klemens (1993) considered them secure in eastern New ­England but vulnerable in western New ­England due to ­human activity. Fowler’s Toad is considered rare in Ontario ­because of its restricted distribution (Green, 1989) and its loss of breeding habitat. It has a discontinuous distribution along the north shore of Lake Erie, and several populations have dis­appeared since the 1950s (e.g., Choquette and Jolin, 2018). Populations at Point Pelee Park dis­appeared ostensibly ­because of ­human disturbance, and populations on Pelee Island dis­appeared due to intensive agriculture associated with pesticide use. Other populations may be stable, however, except at Long Point. Using a 23 yr dataset, Greenberg and Green (2013) showed that population declines at Long Point ­were associated with the invasion of the nonindigenous common reed, Phragmites australis, which led to a loss of breeding habitat. In turn, this reduced recruitment and population growth, despite the maintenance of adult habitat. At least ­until 2002, the population at Long Point appeared to be large but with occasional fluctuations (Logier, 1931; Evans and Roecker, 1951; Green, 1989, 1992). From 2002 to 2011,

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however, the population underwent a sustained decline. The age structure of the population was similar between the sexes, but was highly variable both before and ­after 2002. During the decline, the average age among 469 toads examined increased, but ­there was no change in adult survivorship. ­These results suggested an aging population with reduced recruitment, likely related to the loss of breeding habitat caused by the aforementioned Phragmites (Middleton and Green, 2015). The conservation status of this species in Canada and its potential critical habitat was summarized by Green (1989) and Weller and Green (1997) and updated by Green et al. (2019). Like many other toads, Fowler’s Toad appears resilient to certain types of habitat disturbances, such as agriculture, silviculture, and suburbanization (Ferguson, 1960; Foster et al., 2004; Jensen et al., 2008; Barrett et al., 2016), as long as breeding sites are maintained and suitable overwintering sites are available. The presence of ­cattle in breeding ponds had ­little effect on postmetamorphic Fowler’s Toads, and Burton et al. (2009) suggested that toads may benefit from controlled grazing. This may be due to the toad’s preference for open habitats, in contrast to the American Toad that prefers forested woodlands. For example, Fox et al. (2004) found few Fowler’s Toads compared to American Toads in diversely managed forested watersheds in Arkansas regardless of type of management used. Still, clearcutting may be detrimental to the species, at least over a short-­term period (McLeod, 1995; McLeod and Gates, 1998). Fowler’s Toads also appear to tolerate prescribed burning inasmuch as this opens up the habitat making it more favorable from the toad’s perspective (McLeod, 1995; McLeod and Gates, 1998; Floyd et al., 2002). Fowler’s Toads rapidly colonize newly constructed or restored wetlands (Briggler, 1998; Mierzwa, 1998; Merovich and Howard, 2000; Brodman et al., 2006; Palis, 2007; Denton and Richter, 2013; Walls et al., 2014a; Mitchell, 2016), and have been found in reclaimed surface mine sites 19–29 yrs ­after mining had ceased (Myers and Klimstra, 1963). However, Stiles et al. (2017a) observed only 6 individuals at 2 ponds over a 2 yr period at a mine reclaimed site in Indiana. Fowler’s Toads have been successfully repatriated to several locations owned by the National Park Ser­vice in the New York City metropolitan area using mostly larvae for release (Cook, 2008). The only unsuccessful release site was impacted by saltwater overwash. This species is considered Endangered in Ontario and Vermont, Threatened in New Hampshire, of Special Concern in Michigan, and has been proposed as of Special Concern in Connecticut (Klemens et al., 2021). The species is considered Endangered by COSEWIC.

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Anaxyrus hemiophrys 105

Anaxyrus hemiophrys (Cope, 1886) Canadian Toad Crapaud du Canada ETYMOLOGY

hemiophrys: from the Greek words hemi meaning ‘half ’ and ophrys meaning brow or eyebrow. The name might refer to the eye shape. NOMENCLATURE

Conant and Collins (1998) and Stebbins (2003): Bufo hemiophrys Fouquette and Dubois (2014): Bufo (Anaxyrus) hemiophrys Synonyms: Bufo woodhousii hemiophrys

Males are generally smaller than females. In contrast, Underhill (1961) found no statistical difference in the size of South Dakota adults, with a male mean of 51.5 mm SUL and a female mean of 52.6 mm SUL. Other specific rec­ords include: 58–68 mm SUL for males and 56–80 mm SUL for females (Wright and Wright, 1949); in Minnesota, the largest male mea­sured by Breckenridge and Tester (1961) was 60 mm SUL, whereas the largest female was 68 mm SUL; Cook (1964b) reported a maximum size of 85 mm SUL for males and 91 mm SUL for females. Larvae. The tadpole is darkly pigmented dorsally. The tail musculature is bicolored, the tail fins are unpigmented, and the ventral tail fin is narrow. The throat area is clear, and the venter is light, extending to the gut. TL is ca. 30 mm. Eggs. The eggs have not been described. Presumably they are similar to ­those of the closely related A. americanus. DISTRIBUTION

IDENTIFICATION

Adults. Canadian Toads are brown to brownish green to gray, but occasional individuals may be reddish to reddish brown in color (Cook, 1964b). The cranial crests fuse to form a con­spic­u­ous boss (bump) between the eyes, and postorbital crests are weak or absent. A vertebral stripe is pre­sent down the midline of the dorsum. Parotoids are wide and long and are not greatly elevated. The species has numerous (10–14) dark spots dorsally, with 1–2 warts per spot. Warts are darker than the ground color, although they are lighter than the dark patch that surrounds the warts. Venters are dirty white or slightly yellowish with variable dark flecking or spotting. Males have a swollen thumb and are darker in coloration than females all year-­round; during the breeding season, they also have a dark throat. Underhill (1961) provided additional information on morphology.

The range of the Canadian Toad extends from the extreme southern portion of the Northwest Territories south throughout eastern Alberta to just south of the border with Montana. The range then extends eastward through western North Dakota, then south across eastern North Dakota to northeastern South Dakota and western Minnesota. From ­there, the species occurs northwest across southwestern Manitoba and much of Saskatchewan. The species barely enters the extreme southwestern corner of Ontario. Harper (1963) corrected some previously erroneous rec­ords from the northern portion of the range. Impor­tant distributional references include: Alberta (Russell and Bauer, 2000), Manitoba (Harper, 1963), Minnesota (Oldfield and Moriarty, 1994; Moriarty and Hall, 2014), Montana (Black, 1970, 1971), North Dakota (Wheeler and Wheeler, 1966; Hoberg and Gause, 1992), Northwest Territories (Preble, 1908; Harper, 1931; Timoney, 1996; Fournier, 1997, undated), Saskatchewan (Heisler et al., 2013), and South Dakota (Peterson, 1974; Fischer, 1998; Ballinger et al., 2000; Kiesow, 2006). FOSSIL REC­O RD

Fossils of the Canadian Toad have been reported from Pleistocene Irvingtonian deposits from Kansas and Texas, and from Rancholabrean deposits in Alberta (Tihen, 1962b; Holman, 2003). Bones of this species are often referred to as Bufo cf. B. hemiophrys. SYSTEMATICS AND GEOGRAPHIC VARIATION

Head pattern of Anaxyrus hemiophrys. Illustration by Camila Pizano

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Anaxyrus hemiophrys is a member of the Americanus clade of North American bufonids, a group that includes A. americanus, A. baxteri, A. fowleri, A. houstonensis, A. terrestris, and A. woodhousii. It is closely related to A. baxteri and somewhat

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106  Bufonidae

ADULT HABITAT

Distribution of Anaxyrus hemiophrys

more distantly to A. fowleri, A. americanus, and A. houstonensis (Pauly et al., 2004). Suggestions that A. hemiophrys is more closely related to A. microscaphus than the Americanus clade (Blair, 1963b) are not supported. The Canadian Toad hybridizes in nature with a number of other toads in narrowly defined contact zones. ­These include A. americanus (Cook, 1983; Henrich, 1968; Green, 1983, 1996; Green and Pustowka, 1997), A. boreas (Cook, 1983), A. cognatus (Brown and Ewert, 1971), and A. woodhousii (Meacham, 1962). Interspecific amplexus has been reported ­under natu­ral conditions (Cook, 1983; Eaton et al., 1999). Hybridization in the laboratory has been reported with A. americanus, A. houstonensis, A. microscaphus (small % reach metamorphosis), A. terrestris, and A. woodhousii, but not with A. punctatus (Moore, 1955; Blair, 1961a, 1963a, 1972). Even hybrids backcrossed with the parental species frequently produce larvae that survive through metamorphosis. The rusty color phase of A. hemiophrys has been described in detail and mapped by Cook (1964b). The frequency of toads with rusty coloration varies among populations, but most populations with rusty toads are found in west central Manitoba and adjacent east central Saskatchewan. In populations containing rusty toads, from 6–35% of the individuals observed are reddish. The rusty color phase appears absent from Alberta and much of southern Saskatchewan and southwestern Manitoba. What­ever its origin, the rusty coloration is not associated with substrate color.

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Anaxyrus hemiophrys is usually found in damp open areas adjacent to more permanent ­water bodies such as lakes and rivers. This species is usually found in open sites close to ­water (e.g., 80% within 8 m; Breckenridge and Tester, 1961), but some ­limited dispersal occurs into adjacent mixed upland forests. Dense cover is preferred as microhabitat in experimental choice tests, although the species does not avoid open substrates (Tester et al., 1965). Preferred habitats include grass (e.g., Andropogon sp.) meadows and willow (Salix sp.) bogs near ­water and in the surrounding poplar (Populus tremuloides) and willow forests and aspen parklands of the Transition and Canadian (Boreal) life zones (Stebbins, 1954; Breckenridge and Tester, 1961; Roberts and Lewin, 1979; Cook, 1983). In other areas, it is found associated with prairie ponds and lakes (Underhill, 1961; Henrich, 1968; Williams, 1969), and fen and open-­water wetlands near upland habitat (Annich et al., 2019). Canadian Toads avoid the Jack Pine (Pinus banksiana) forests of Alberta (Roberts and Lewin, 1979), as they do not like dry substrates. The open habitats preferred by this species experience more rapid temperature fluctuations than closed-in forest habitats. TERRESTRIAL ECOLOGY

This is definitely a cold-­adapted species. The activity season may be very short, especially at the northern portion of the species’ range. In the Northwest Territories, for example, toads exit their hibernacula around the first week of May and return the first week of September, making the activity season 3.5–4 months (Kuyt, 1991). Emergence in Manitoba occurs in May, although full chorusing does not occur ­until June (Tamsitt, 1962). In Minnesota, emergence occurs from late April to mid-­June over a 5–6 week period, with the peak of emergence in mid-­May (Tester and Breckenridge, 1964a; Kelleher and Tester, 1969). First emergence dates can vary by as much as 20 days, depending on environmental conditions, and emergence only occurs when all traces of soil frost have dis­appeared (Kelleher and Tester, 1969). Emergence may be triggered by a period of warming weather (>15–21°C) or precipitation. Adults emerge prior to juveniles, but juveniles are active ­later in the season than adults. As with many amphibians, Canadian Toads do not occupy ­every wetland that appears suitable. In Alberta, for example, they ­were found at 9 of 25 sites sampled, bred at only 7 sites, and ­were common at only 3 sites (Roberts and Lewin, 1979). Canadian Toads tend to be found near ­water and appear less purely terrestrial than many other bufonids. However, ­there is some disagreement in the lit­er­a­ture as to the extent of summer nonbreeding activity in relation to ­water. For example, Breckenridge and Tester (1961) reported most

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Anaxyrus hemiophrys 107

activity within close proximity to the margins of prairie ponds and lakes in Minnesota, whereas ­others (Cook, 1983; Constible et al., 2010) have noted that in boreal forest populations, this does not appear to be the case. Toads wander extensively in upland forest, and combined short-­term movements can translate into long-­distance directional movements that vary in timing and tortuosity of the routes taken. In 1 study, toads moved from 4.7–116.6 m/ day and traveled up to 4.7 km with a maximum displacement of 1,836 m (Constible et al., 2010). In a study of toads in a prairie region, the maximum distance between captures was 341 m (Breckenridge and Tester, 1961). Movements are not stimulated by rainfall, and movements among individuals do not vary synchronously; they occur both by day and by night. Toads frequently stay in one place for several days, then suddenly move to a new location. Differences in nonbreeding habitat use and activity may reflect variation in the habitats occupied by the toads and perhaps other environmental differences. Daily movements are generally 38°C for only 30 min or less (Williams, 1969). When tadpoles encounter high temperatures, they immediately move ­toward cooler portions of the wetland.

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The diet of adult A. hemiophrys consists mostly of arthropods, almost all of them insects. Beetles and ants are particularly favored; other prey include grasshoppers, locusts, mayfly and damselfly nymphs, cicadas, true bugs, vari­ous types of flies, ichneumonids, spiders, and rarely snails (Moore and Strickland, 1954). However, ­there is 1 report of a large Canadian Toad attempting to eat a young Red-­ winged Blackbird (Agelaius phoeniceus) (Cook and Cook, 1981). Juveniles consume smaller prey, such as beetles, midges and other small flies, springtails, mites, and ants. Flies are very impor­tant in the juvenile diet. The types of prey reflect the terrestrial habits of the toad. PREDATION AND DEFENSE

Like other toads, Canadian Toads have noxious skin secretions concentrated in the granular glands of the warts and parotoids. The toads are well-­camouflaged, especially when buried in the sand with only the eyes showing. When approached, they may leave their subsurface retreats and move rapidly into the surrounding vegetation. They readily take to ­water and run quickly through vegetation to make their escape. However, 2 cases of death-­feigning have been reported (Nero, 1967; McNicholl, 1972), and toads often remain immobile and hunkered ­toward the ground when approached. Juveniles are eaten by a wide range of predators, including Garter Snakes (Thamnophis sp.), birds (Red-­tailed Hawk),

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Anaxyrus hemiophrys 109

and mammals (raccoons, badgers) (Breckenridge and Tester, 1961; Tamsitt, 1962; Tester and Breckenridge, 1964a). ­There are many reports of pos­si­ble predators for Canadian Toads, but few ­actual observations. POPULATION BIOLOGY

­ here is a certain degree of regional variation in the life T history characteristics of this species. The largest Canadian Toads are generally found in the south, whereas areas with the smallest toads are located in 3 populations in the ­middle of the latitudinal range (Eaton et al., 2005b). ­These latter populations also contained the oldest individuals. Despite regional variation, size and age are positively correlated within a population. Populations may be male or female biased. For example, toads caught emerging from overwintering sites at a drift fence in Minnesota had a sex ratio of 1.13:1, whereas hand-­caught toads had a ratio of 4.1:1. Hand catching toads could yield a biased sex ratio ­because of male visibility and activity, and Breckenridge and Tester (1961) suggested that females actually left the pond margins and ­were therefore less likely to be captured. In other data presented by ­these authors, the overall sex ratio was 1:1 in 1 year and 1.3:1 a second year. Additional observations suggested that males made up from 33–38% of the population, but that sex ratios varied considerably among sites and years (Kelleher and Tester, 1969). Growth occurs rapidly, but growth rates also vary among populations (Eaton et al., 2005b). Toadlets metamorphosing at 11–12 mm SUL reach 19.3–28 mm SUL in Alberta (Roberts and Lewin, 1979) and 22.3–28.4 in Manitoba (Tamsitt, 1962) by August, and a mean of 31 mm by September in Minnesota (Breckenridge and Tester, 1961). Even small individuals can overwinter successfully, however, as Roberts and Lewin (1979) found 22 mm SUL juveniles in June that must have metamorphosed the previous year. In

Minnesota, males reach maturity by 38–45 mm SUL, as indicated by the presence of motile sperm; females are mature by 43–45 mm SUL, as indicated by the presence of pigmented oocytes (Tester and Breckenridge, 1964a). Thus, it is pos­si­ble that males could reach a minimum breeding size midway to late in the first breeding season following their metamorphosis. Female Canadian Toads reach maturity by the second summer ­after metamorphosis (Eaton et al., 2005b). The age at maturity may vary, however, depending on when tadpoles metamorphose. If early in the season, a long growing season allows them to reach maturity by the end of the first summer; ­those metamorphosing late in the season would be delayed by the necessity of overwintering and resuming growth the following spring. In this case, maturity may not be reached ­until ­after the breeding season, so the first opportunity for breeding may occur well ­after maturity is reached. At 1 population in Alberta, however, females did not reach sexual maturity ­until 4 yrs, although the sample size was very small; in another population, both males and females did not reach maturity ­until 3 yrs of age. ­These data suggest the potential for considerable variation among populations in life history traits, perhaps depending on environmental conditions. Maximum longevity is 7–12 yrs, but again, ­there is much variation among populations (Eaton et al., 2005b). Simply put, toads in some populations live much longer than ­those in other populations. The density of Canadian Toads can be rather high. Roberts and Lewin (1979) recorded 12/1,000 m2 in Alberta, with maximum densities found within 50 m of ­water. Densities become lower as the toads disperse into mixed upland forests. Other reports suggest that populations are not as large as ­those of other toad species, with adults being more widely dispersed. In Minnesota, the ratio of juveniles to adults was 1.24–14.25:1, depending upon year sampled (Tester and Breckenridge, 1964a; Kelleher and Tester, 1969). Mortality of the small size classes is high, especially due to ­water loss, temperature extremes, and predation. Mortality of overwintering toads occurs when frost is deep, especially as a result of lower snowfall amounts than normal. In such conditions, juveniles might not be as able as adults to burrow beneath the deepening frost level. Over 1 winter sampled, however, survival rates ­were about equal for males, females, and juveniles. Kelleher and Tester (1969) estimated annual survivorship at 24–44% depending upon year and location. COMMUNITY ECOLOGY

Adult Anaxyrus hemiophrys. Photo: Jeffrey LeClere

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The presence of fish may or may not influence survivorship of A. hemiophrys larvae. In Alberta, for example, larval recruitment is not affected following winters in which severe fish kills occur. Reduced fish abundance may even decrease the abundance of developing toad tadpoles, perhaps ­because of

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110  Bufonidae

complex trophic interactions among dif­fer­ent types of fish and insect abundance (Eaton et al., 2005a). Experiments have not demonstrated that small-­bodied fish can have significant predatory effects on toad larvae in boreal habitats, perhaps ­because the toad tadpoles are distasteful and not an impor­tant part of fish diets ­under normal circumstances. The range of this species may be allopatric or sympatric to that of other toads. In Minnesota, for example, A. hemiophrys is allopatric with A. americanus. Whereas A. hemiophrys is found in prairie habitats, A. americanus is confined to forested habitats. Neither species breeds in ponds in the ecotone between ­these habitats. Thus, ­there appears to be habitat segregation between them in contact zones in the Itasca and Waubun region (Williams, 1969). Not surprisingly, larval prairie-­dwelling A. hemiophrys have a higher thermal tolerance than the forest-­dwelling A. americanus since they are more likely to be exposed to sunlight in the open prairie habitat. In other regions, ecotones may form an impor­tant area for hybridization between closely related species. In southeastern Manitoba, for example, a 3 km wide zone of hybridization occurs between the prairie species A. hemiophrys and the eastern forest species A. americanus (Green, 1983). Unlike the area in Minnesota, hybridization occurs extensively within this narrow band, where most toads are intermediate between the parental species. The hybrid zone prob­ably formed about 8,000 years ago ­after the retreat of glacial Lake Agassiz, which brought ­these species into secondary contact. Although ­there is a degree of stability in the hybrid zone, it appears to be shifting westward. DISEASES, PARASITES AND MALFORMATIONS

The amphibian pathogen Batrachochytrium dendrobatidis (misidentified as Basidiobolus ranarum) was reported from Canadian Toads originally captured in North Dakota (Taylor et al., 1999b). Canadian Toads have been used as a surrogate species for understanding the effects of the chytrid pathogen on A. baxteri, a critically endangered species. For example, transmission of the fungus from infected to healthy toads was confirmed via clinical experiments using this species (Taylor et al., 1999e). Myiasis from green blowfly (Lucilla silvarum) larvae has been reported from Canadian Toads in Alberta (Eaton et al., 2008). A single Canadian Toad with an unspecified malformation was reported by Converse et al. (2000). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

­ here are few data on the effects of habitat alteration on this T species. Canadian Toads breed in open areas, including

Dodd_Canada_int_5pgs_B1&B2.indd 110

Breeding habitat of Anaxyrus hemiophrys. Photo: Cindy Paszkowski

recent clearcuts. One study on the effects of clearcutting around ponds could not demonstrate that the width of a buffer strip was impor­tant for conserving Canadian Toads. The authors (Hannon et al., 2002) therefore suggested that in some habitat types, a buffer zone around wetlands may be small and yet be adequate to protect this species. Anaxyrus hemiophrys ­will colonize restored wetlands, although the wetlands must be within the relatively short dispersal distance of the species (Lehtinen and Galatowitsch, 2001). In the Prairie Pothole Region, they occupy restored conservation grasslands (Ba­las et al., 2012). Anaxyrus hemiophrys is uncommon and not protected in the Northwest Territories, but most populations occur in Wood Buffalo National Park (Fournier, 1997). Timoney (1996) recommended population monitoring and complete protection of this northernmost overwintering site from ­human disturbance. Primary concerns involved road modification, vehicular traffic, and vegetation intrusion. ­There is no evidence of declines in Saskatchewan (Didiuk, 1997), but declines have been reported in central and southern Alberta and in parts of Manitoba, perhaps due to drought and the destruction of wetlands (Weller and Green, 1997; Russell and Bauer, 2000). ­These latter authors noted declines in Elk Island National Park, Alberta, from 1971 to the mid-1980s, suggesting that habitat alteration alone is not responsible for the species’ decline. Drought, on the other hand, decreases the amount of snowpack and thus may expose dormant toads to excessively cold temperatures normally mediated by the snow’s insulation capacity. This species is included in a management plan for amphibians in the Northwest Territories (Government of the Northwest Territories, 2017). The species is considered secure in Manitoba and Saskatchewan, but data deficient in Alberta.

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Anaxyrus houstonensis 111

Anaxyrus houstonensis (Sanders, 1953) Houston Toad ETYMOLOGY

houstonensis: named ­after Houston, Texas. NOMENCLATURE

Conant and Collins (1998): Bufo houstonensis Fouquette and Dubois (2014): Bufo (Anaxyrus) houstonensis Synonyms: Bufo americanus houstonensis IDENTIFICATION

Adults. The Houston Toad is a small, brown, gray, or purplish-­gray toad with a somewhat herringbone pattern of dark brown or greenish blotches separated by light areas between them. The postorbital crest is large. Parotoids are elongated, about twice as long as broad. ­There are from 1 to 5 warts per spot, but rarely >3. Warts are smooth and rounded. A mid-­dorsal light stripe may be pre­sent. Venters have some midventral spotting anteriorly but not posteriorly, and are other­wise pale. The dorsal surface of the femur is dark striped. Males have dark throats during the breeding season. Sanders (1953) described the osteology in detail. Males are slightly smaller than females. Males ­were ca. 52–64 mm SUL (mean 57 mm) in 1 Bastrop County study and ­were 50–66 mm SUL (mean 57 mm) in the Houston area (Brown, 1971b). In another study in Bastrop County, males averaged 61 mm SUL (range 53–77 mm) and females 66 mm SUL (range 54–84 mm) (Jacobson, 1989). The smallest female from Bastrop County producing eggs was

Head pattern of Anaxyrus houstonensis. Illustration by Camila Pizano

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59.5 mm SUL, and the largest was 81 mm SUL; males w ­ ere 53–61.8 mm SUL (Quinn and Mengden, 1984). A fourth study in Bastrop found that males averaged 57.1 mm SUL and females 63 mm SUL (Hillis et al., 1984). Larvae. The body of the tadpole is dark without contrasting markings. Venters are evenly pigmented. The throat is largely pigmented, and ­there are no gaps in the dark pigment on the dorsal part of the tail musculature. The light ventral part of the tail musculature is narrow. Tail fins, however, are mostly unpigmented. The snout is sloping in lateral view. TL is 7 g/m3), relative humidity (>37%), a barometric pressure >752 mmHg, and an average daily wind speed 13°C (Brown et al., 2013a). Houston Toads are somewhat of an explosive breeder, with most males appearing over a short number of nights (3–5) early in the season. Jacobson (1989) found that males ­were at a breeding pond for only 24 nights over a 4 month period. Some amplectant males stayed as long as 10 nights (median 5 nights), but non-­amplectant males only stayed for a median of 3.5 nights (Jacobson, 1989). About one-­third of the males ­were at breeding ponds only once during a season, with ­others showing up multiple times depending upon rainfall (Kennedy, 1962; Hillis et al., 1984; Price, 2003). Males begin calling just before sunset, often initially from terrestrial burrows 1–40 m from the breeding site (Hillis et al., 1984). The burrows are not constructed by the toads, but consist of rodent burrows, old root burrows, or spaces ­under downed logs. ­These retreats are frequently located in gullies leading to the breeding pond. ­After a night of calling, the toad ­will retreat back to a burrow ­until the next night. Movement to breeding sites occurs at just about sunset and continues ­until ­after midnight. Females arrive several hours ­after sunset and are quickly amplexed. By 02:00, females have ceased moving to ponds. Males mate 1–3 times per season, but only a very few mate more than once. In Jacobson’s (1989) study, 56% of the males ­were never observed in amplexus. Males that are successful in amplexus and fertilization may not be dif­fer­ent

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in size than unsuccessful males in some years (Jacobson, 1989), although in other years successful males are larger than non-­successful males (Hillis et al., 1984). ­These latter authors suggested that male experience and/or dominance ­were more impor­tant in mating success than female preference for large males. A few females also ­were observed in amplexus with more than 1 male during the breeding season. Males space themselves randomly around breeding pools and call from the shoreline, shallow ­water, or from on top of debris well above the ­water’s surface. The call of A. houstonensis is a long musical trill. The dominant frequency in 1 study was 2,300 cps with a mean duration of 7.3 sec (range 3.8–11.2) and a mean of 32 trills/sec at 19.5°C (Blair, 1956b). Brown (1971b) recorded calls with a dominant frequency of 1,915–2,089 cps with a duration of 9.5– 16.6 sec and 18.7–32.3 trills/sec at 14.5–22.1°C. Amplexus may last 24 hrs, and Jacobson (1989) recorded 1 pair that stayed together 34 hrs. Amplexus can take place for up to 6 hrs prior to oviposition (Hillis et al., 1984). Once a female begins oviposition, she does not stop ­until the entire clutch is laid, even if this means continuing well into daylight. Males do not release females for as long as 0.5 hrs ­after oviposition has ceased. Males grasp any other toad within their range in their attempts at amplexus, including already amplectant pairs. Males are rarely (only about 8% of the time) successful in attempts to displace other males. Females also have been observed displacing unwanted males. Interspecific amplexus has been observed between Houston Toads and I. nebulifer (Brown, 1971b). Previous reports that Houston Toads do not use the same breeding ponds from one year to the next are inaccurate. Using surveys, telemetry, movement observations, and juvenile dispersal data, Vandewege et al. (2013) showed that ­there is a high degree of breeding site fidelity in this species. Fidelity is evident both within and among years, and most toads remain within 75 m of the pond of initial capture. BREEDING SITES

The Houston Toad breeds in temporary rain pools and ditches located within its forest habitat. Pools must persist at least 30 days to allow for larval development. Forstner and Ahlbrandt (2003) noted that pH (range 5.33–6.7) is not a ­factor in breeding site se­lection, but that ponds with an embankment slope of 10° or less had a higher associated reproductive success than ­others. Still, pool levels fluctuated substantially. The best ponds for reproduction have multiple shallow zones across all pool height levels. REPRODUCTION

Houston Toads breed in the spring and early summer, with rec­ords extending from February to June, although most

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Egg strings of Anaxyrus houstonensis. Photo: Michael Forstner

breeding occurs from mid-­February to early April (Kennedy, 1962; Brown, 1971b; Hillis et al., 1984; Price, 2003). Breeding aggregations may form a few days before ­actual oviposition takes place, but it is normal to have first calling and oviposition occur on the same night. Despite the potentially long season, oviposition occurs on only a very few nights and may be delayed ­until late in the season if spring rains do not arrive. Jacobson (1989) found females at a breeding site on only 15 nights in 4 months, and oviposition occurred on only 7 nights. Nearly all females ­were pre­sent on only 1 or 2 consecutive nights, and virtually all females visited a pond only once during the breeding season. Nearly all eggs are oviposited during the second and third night of the main chorusing. Oviposition occurs from 19:00 ­until noon the following day, with most occurring ­after 03:00. Clutch sizes in captivity ranged from 513–6,199 (Quinn and Mengden, 1984). Kennedy (1962) reported a single clutch size of 728 from a hybrid cross with I. nebulifer. Greuter (2004) obtained estimates of a mean of 1,279–2,098 eggs, depending upon which counting technique was used, with 2 exact counts of 2,807 and 4,211. It has been suggested that females ­will oviposit more than1 clutch per season, but it has not been demonstrated. Hatching occurs in 2–8 days and is temperature dependent (Hillis et al., 1984; Quinn and Mengden, 1984). The warmer the temperature, the sooner the eggs hatch. At hatching, larvae are 6.1–6.7 mm TL. LARVAL ECOLOGY

Larvae consume algae and pine pollen and have been observed eating the jelly envelopes of previously hatched eggs. During the day, tadpoles may form large aggregations, but ­these aggregations tend to disperse at night. Tadpoles tend to remain on the substrate by day, but at night they forage on detritus and algae that is attached to aquatic

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114  Bufonidae

corridors, such as gullies, ravines, and water-­filled drainages to facilitate dispersal (Hillis et al., 1984; Thomas and Allen, 1997). The maximum dispersal distance is reported from a head-­started juvenile caught 1.34 km from its release point (Vandewege et al., 2012). Juveniles prefer areas with moisture and shade, are found on both clay and sandy substrates, and occupy both pine and mixed oak-­juniper vegetation communities (Greuter, 2004). Sunny dry habitats are avoided. DIET

Tadpole of Anaxyrus houstonensis undergoing metamorphosis. Photo: Jim Godwin

vegetation, along the shoreline, or at the pond surface. In captivity, the duration of the larval period ranges from 15 to 100 days (Quinn and Mengden, 1984), whereas the larval period lasts about 60–65 days in a wild population (Hillis et al., 1984). Larvae reach a maximum size of 20–22 mm. Newly metamorphosed toadlets are 7–12.5 mm SUL in the lit­er­a­ture, but Sirsi et al. (2020) noted toads “immediately ­after metamorphosis” at 17–18 mm SUL. Larvae do not respond to chemical cues from potential invertebrate predators in the absence of conspecific or heterospecific alarm cues. However, they reduce their activity levels significantly when exposed to chemical cues resulting from attacks on conspecifics or other bufonid (e.g., I. nebulifer) species (Preston and Forstner, 2015b). ­These authors suggested that the time to metamorphosis could be extended should tadpoles develop in dense aggregations with other species that might be attacked by predators.

Houston Toads feed on many invertebrates, including beetles, flies, lacewings, and small moths (Bragg, 1960b). They appear to exhibit some degree of selectivity by avoiding certain types of scarab beetles. Juveniles are both sit-­and-­wait and active foragers, particularly feeding on small ants (Thomas and Allen, 1997). Juveniles use shallow burrows—­which they create by digging backward into the sand—as ambush sites. PREDATION AND DEFENSE

Tadpoles are eaten by the snakes Nerodia erythrogaster and Thamnophis proximus; newly metamorphosed toadlets are killed by nonindigenous fire ants (Solenopsis invicta) (Freed and Neitman, 1988; Greuter, 2004; Sirsi et al., 2020). Nerodia erythrogaster also is a predator of adults (Zughaiyir et al., 2021). McHenry et al. (2010) reported an American Bullfrog consuming an adult Houston Toad. As with other toads, the skin contains noxious or poisonous secretions that deter a variety of predators. POPULATION BIOLOGY

In a study at breeding ponds in Bastrop County, Texas, the sex ratio of A. houstonensis was male biased (11 males per female), but in traps set away from the ponds, the sex ratio was quite dif­fer­ent (2.8 males per female). The probability of

JUVENILE DISPERSAL

Immediately ­after metamorphosis, toadlets may return to ­water briefly before dispersing. Larvae metamorphose si­mul­ta­neously in very large numbers. Initially they remain near the ­water’s edge (ca. 12 m) for about 2 weeks (Vandewege et al., 2013), then gradually disperse to adjacent forested uplands. Dispersal occurs both day and night. ­After 48 hrs, toadlets disperse about 3.2 m from ­water (range 0.7–5.13 m) and seek refuge buried ­under grass or sedge tussocks (Swannack et al., 2006). By the 13th–19th day, they can travel at least 8 m, and by day 30 they can be up to 35 m away (Greuter, 2004; Vandewege et al., 2013). In contrast to Swannack et al.’s (2006) observations, however, Greuter (2004) noted that juveniles often stayed in the vicinity of ­water for up to 3 weeks prior to beginning to disperse. Recent metamorphs are capable of dispersing as far as 100 m or more from the breeding site, and may use physical

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Adult Anaxyrus houstonensis. Photo: Robert A. Thomas

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capturing females in traps was 3.5 times greater than when capturing them at ponds, whereas the disparity was only 0.28 times for males ­after adjusting for sampling effort. A male-­biased sex ratio at ponds is not surprising since males likely remain at ponds longer than females, who deposit their eggs and leave soon ­after oviposition. Still, the overall sex ratio was highly male biased and changed from one year to the next. This led Swannack and Forstner (2007) to suggest that populations of Houston Toads are male biased ­because of differences in the age at first reproduction between males and females. Quinn and Mengden (1984) suggested females reach maturity in 14–15 months and prob­ably first breed the second spring following transformation; males likely breed the first spring following metamorphosis. Interestingly, Houston Toads raised in captivity have a 1:1 sex ratio (Jones et al., 2017). As might be expected, larval and early stage metamorph survival is quite low. Greuter (2004) found a larval survivorship of only 4.73%. From metamorphosis to 13 weeks post-­metamorphosis, the population estimate further declined by 15%. In arrays around natu­ral and artificial ponds, Greuter (2004) captured from 15 to 332 metamorphosed juveniles; numbers captured varied by year and location. At hatching, juveniles are 400 mg/L (Birge et al. 1980). STATUS AND CONSERVATION

Historically, A. quercicus was considered an extremely abundant species, especially in Florida. Hamilton (1955) described it as extraordinarily abundant, and Carr (1940a) stated that it was pos­si­ble to drive across the central Florida peninsula and never be out of the sound of breeding choruses. With extensive landscape alteration and the destruction of unique communities and breeding ponds, ­those days are gone. The Oak Toad is particularly susceptible to habitat loss, as it depends on small temporary pools in which to breed. Such habitats are easily overlooked or ignored, and most are unprotected. Likewise, the species cannot tolerate fish, so the introduction of fish into formerly fish-­free wetlands is detrimental to this species. ­Because of its small size, the species also may be sensitive to the presence of red imported fire ants (Solenopsis victa) that have invaded its habitat. Oak Toads do not tolerate urbanization well, and abun-

Breeding habitat of Anaxyrus quercicus. Other species breeding at this site included A. gryllus, A. terrestris, G. carolinensis, D. femoralis, D. squirella, L. capito, L. sphenocephalus, P. ocularis and S. holbrookii. Photo: C.K. Dodd, Jr.

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dance may be reduced in silvicultural and agricultural areas (Delis et al., 1996; Surdick, 2005). For example, Oak Toads ­were common in northwest Gainesville, Florida, in the early 1980s, but have virtually dis­appeared as small wetlands have been destroyed. Guzy et al. (2012) indicated that although this species was a reliable indicator of wetland health, it was effectively excluded from urbanized wetlands in the Tampa Bay region. Fi­nally, alteration of precipitation patterns leading to extended periods of drought make this essentially annual species vulnerable to regional climate change. Oak Toads may be found in cypress flatwoods ponds ­after ditching (Vickers et al., 1985), although ­whether reproduction takes place has not been documented. They also colonize areas previously clearcut 1.5–4 yrs ­earlier (Enge and Marion, 1986; Russell et al., 2002b). It should be noted, however, that ­there was no replication in some of ­these studies, and that conclusions about short-­term tolerance to clearcutting ­were based on presence, not reproduction or demographic analy­sis. The presence of A. quercicus is indicative of low-­intensity land use (Surdick, 2005). The Oak Toad inhabits fire-­maintained ecosystems throughout its range, and prescribed fire regimes seem to enhance the habitat quality for this species (Langford et al., 2007). Native grasses increase following prescribed fire, and clumps of grasses provide ideal microhabitat for Oak Toads (Baxley and Qualls, 2009). When fire is excluded from breeding sites for ca. >8 yrs, Oak Toads dis­appear (Klaus and Noss, 2016). The species readily re-­colonizes wetlands that experience saltwater overwash during tropical storms (Gunzburger et al., 2010), but does not seem to take to newly constructed ponds (Pechmann et al., 2001). The species is considered a Species of Greatest Conservation Need in ­Virginia’s Wildlife Action Plan (Pague and Mitchell, 1987; Mitchell and Reay, 1999; https://­dwr​.­virginia​.­gov​ /­wildlife​/­information​/­oak​-­toad​/­). In North Carolina, Dorcas et al. (2007) noted that they have under­gone dramatic declines in recent years and attributed declines to habitat loss, acidification of breeding ponds, and the potential impacts of disease and red imported fire ants. Enge (2005a) reported 2,131 A. quercicus ­were harvested for the pet trade from 1990 to 1994 in Florida.

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Anaxyrus retiformis 141

Anaxyrus retiformis (Sanders and Smith, 1951) Sonoran Green Toad

mea­sures ca. 1.15 mm in dia­meter. Eggs are deposited singly or in short strands. The outer envelope of the eggs has an agglutinating property that ­causes them to clump together, giving the appearance of a small egg mass rather than single eggs or a small string.

ETYMOLOGY

DISTRIBUTION

retiformis: from the Latin retis meaning ‘of a net’ and forma meaning ‘shape.’ The name is in reference to the reticulated dorsal pattern of black lines. NOMENCLATURE

Stebbins (2003): Bufo retiformis Fouquette and Dubois (2014): Bufo (Anaxyrus) retiformis Synonyms: Bufo debilis retiformis

Anaxyrus retiformis is known only from south central Arizona and west central Sonora, México. Impor­tant distributional references include Nickerson and Mays (1968), ­Sullivan et al. (1996b), Blomquist (2005), Brennan and Holycross (2006), López et al. (2009), Murphy (2019), and Holycross et al. (2021). FOSSIL REC­O RD

No fossils have been identified.

IDENTIFICATION

Adults. This is an attractive, small, yellow-­green toad with a dark reticulated pattern covering the dorsum and upper portions of the limbs. The dorsum is not spotted as in A. debilis. The head is flat. Parotoid glands are large in relation to the size of the toad and are patterned like the dorsum. Venters are unspotted. Males are smaller than females. Savage (1954) reported a mean of 43.8 mm SUL (range 40–47 mm) for males and 47 mm SUL (range 45–49 mm) for females but sample sizes ­were very small. ­Sullivan et al. (2000) reported means of 48.1–52.1 for calling males at 3 populations. In museum specimens from Arizona, Goldberg (2020h) reported males at 42–50 mm SUL (mean 46.1 mm) and females 48–62 mm SUL (mean 54.2 mm). Maximum size is 64 mm SUL. Larvae. At hatching, larvae are yellow with scattered dark pigment. Larger tadpoles are pale yellow in coloration and stippled with black and golden patches on the dorsum and tail musculature. They are rounded in appearance with the eyes situated dorsally, and are much paler than most other toad tadpoles. The dorsal tail musculature has more abundant melanophores than the ventral musculature giving a somewhat striped appearance when viewed laterally. Dorsal tail fins may have scattered melanophores, but ventral tail fins are clear. The lateral sides of the body may be darkly pigmented ­behind the eyes, and the belly has golden flecking. ­There is no coloration on the throat and midbelly. Zweifel (1970b) noted that tadpoles are relatively transparent even when full grown, and provides illustrations of larvae at vari­ous stages of development. Eggs. The eggs are light cream to yellow in color with a dark band located one-­third the distance from the top of the animal pole (Ferguson and Lowe, 1969; Zweifel, 1970b). Scattered melanin occurs over the entire egg. ­There are 2 gelatinous membranes surrounding the vitellus, which

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SYSTEMATICS AND GEOGRAPHIC VARIATION

The Sonoran Green Toad was described originally as a subspecies of A. debilis (Sanders and Smith, 1951). It is indeed most closely related to A. debilis within the Nearctic clade of North American bufonids (Pauly et al., 2004), and together with A. kelloggi, ­these species form the Debilis clade within the Punctatus group. ­Under laboratory conditions, this species ­will hybridize and produce larvae that complete metamorphosis with A. punctatus, A. debilis, and A. kelloggi (Ferguson and Lowe, 1969). Hybridization in nature with A. punctatus also has been reported (Bowker and ­Sullivan, 1991; ­Sullivan et al., 1996b). Calls of hybrids between A. retiformis and A. punctatus are aberrant and similar to A. punctatus (­Sullivan et al., 2000).

Distribution of Anaxyrus retiformis

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142  Bufonidae

ADULT HABITAT

The Sonoran Green Toad is a species of the desert flats, valleys, and ­gently sloping bajadas of the lower Sonoran Desert. It occurs in Colorado River Desert Scrub, semidesert grassland, and in Arizona Upland Desert Scrub. TERRESTRIAL ECOLOGY

Sonoran Desert Toads are rarely observed and are nocturnal in be­hav­ior. They remain hidden during the day, presumably in rodent burrows, root channels, and in under­ground crevices. Very ­little is known of the species’ life history. Sonoran Green Toads are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Sonoran Green Toads likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Breeding is opportunistic and associated with desert rainfall. Choruses form rapidly and are often small. In the Sonoran Desert, for example, the chorus size ranged from 3 to 50 and the chorus only lasted a mean of 2 days (­Sullivan, 1989). ­Sullivan et al. (1996b) ­later reported sizeable breeding aggregations consisting of >200 adults. The advertisement call is a short insect-­like buzz with a dominant frequency of 2,714–3,376 cps, a duration of 2.0–4.3 sec, with 193–238 trills/sec (Ferguson and Lowe, 1969; ­Sullivan et al., 1996b, 2000). Savage (1954) described the call as “a rising crescendo of a single drawn-­out note, not unlike the buzz­er of an electric alarm clock, with a slight trill giving the effect of a vibrating police whistle.” The call duration is not correlated with temperature, but the pulse rate and dominant frequency are correlated with temperature (­Sullivan et al.,

Adult female Anaxyrus retiformis. Photo: David Dennis

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2000). Males call at air temperatures of 26–30°C. The call duration is not correlated with SUL, but the pulse rate and dominant frequency are negatively correlated with SUL (­Sullivan et al., 2000). Nocturnal calling occurs on land 1–5 m from the ­water’s edge, and amplexus occurs on dry, damp, or wet substrates. Ferguson and Lowe (1969) recorded amplexus up to 18 m from ­water. Males call from ­under vegetation such as shrubs and grasses, and ­will try to amplex any female moving in their vicinity. Females then carry the smaller male to the breeding site. Satellite males are common at breeding sites, with up to 3 satellites for a single calling male (­Sullivan, 1996b). BREEDING SITES

Breeding occurs in temporary pools and ditches that form ­after summer thunderstorms. ­These include roadside pools in desert washes and ­cattle tanks. REPRODUCTION

Breeding occurs from July to September associated with the summer monsoon rains (Goldberg, 2020h). The smallest reproductively active female examined by Goldberg (2020h) was 48 mm SUL and the smallest male was 42 mm SUL, both collected in August. The clutch size of this species is unknown. Hatching occurs in 2–3 days ­after oviposition. LARVAL ECOLOGY

Laval diets are unknown. The duration of the larval period also is unknown but prob­ably very short; Savage (1954) noted metamorphs 13 days ­after previously observing breeding in the area. At hatching, larvae are 3.1–3.4 mm TL (Zweifel, 1970b).

Larval Anaxyrus retiformis. Photo: Cecil Schwalbe

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Anaxyrus speciosus 143

POPULATION BIOLOGY

Goldberg (2020h) stated that juveniles 16–18 mm SUL collected in August ­were presumed to be young of the year. DISEASES, PARASITES, AND MALFORMATIONS

Sonoran Green Toads are parasitized by the cestode Distoichometra bufonis and the nematodes Aplectana incerta, A. itzocanensis, Oswaldocruzia pipiens, Physaloptera sp., and Rhabdias americanus (Goldberg et al., 1996a). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. Adult male Anaxyrus retiformis. Photo: Rob Schell STATUS AND CONSERVATION DIET

Nothing specific is known of the diet of A. retiformis. ­These toads presumably eat very small invertebrates such as ants and beetles. PREDATION AND DEFENSE

The Sonoran Green Toad presumably has noxious or toxic skin secretions as do other bufonids. Nothing is known concerning defensive be­hav­iors or predators.

Anaxyrus speciosus (Girard, 1854) Texas Toad ETYMOLOGY

speciosus: from the Latin speciosus meaning ‘showy or beautiful.’ Toads, indeed, are beautiful creatures. NOMENCLATURE

Conant and Collins (1998) and Stebbins (2003): Bufo speciosus Fouquette and Dubois (2014): Bufo (Anaxyrus) speciosus Synonyms: Bufo compactilis speciosus, Bufo lentiginosus speciosus, Bufo pliocompactilis, Bufo spectabilis IDENTIFICATION

Adults. The ground coloration of A. speciosus is olive, gray, or gray brown dorsally. ­There is a well-­defined pattern of dusky spots or somewhat elongated blotches dorsally. Cranial crests are low and poorly defined; warts are numerous and low-­to-­rounded without spines. The parotoid glands are oval and less than twice as long as wide. No

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Anaxyrus retiformis presently inhabits most of the same area as reported from historical collections. S­ ullivan et al. (1996b) suggested that hybridization with A. punctatus might be facilitated by the construction of ­cattle tanks that draw ­these normally habitat-­separated species to the same breeding sites. No information is available on status and population trends (Blomquist, 2005), although Bury et al. (1980) noted the need for a status survey >40 yrs ago ­because so ­little was known concerning this species.

mid-­dorsal stripe is pre­sent. Venters almost always lack spots, but Sievert and Sievert (2006) reported a black spot on the chest. This toad has 2 distinctive black spades on each hind foot that are used for digging; the inner spade is sickle ­shaped. Females are larger than males, with adults 65–100 mm SUL. Based on museum specimens, Goldberg (2018b) reported males from 50–73 mm SUL (mean 62.3 mm) and females from 57–98 mm SUL (mean 74.6 mm). Larvae. The dorsal part of the tadpole is lightly pigmented, and the tail musculature is light with dark lateral blotches that tend to form a stripe, giving a bicolored appearance. The body is an elongate oval. Tail fins are low with the greatest height midway down the tail. Eyes are situated dorsally. The maximum size is ca. 30 mm TL. Eggs. The eggs are brown or dark gray dorsally and yellow ventrally. The eggs are oviposited as fine tightly coiled strings in a gelatinous casing. One slightly scalloped jelly envelope surrounds the vitellus. This envelope is 1.8–2.4 mm in dia­meter, with the vitellus 1.2–1.6 mm in dia­meter. According to Livezey and Wright (1947), ­there are 11–17 eggs per 25 mm of string.

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144  Bufonidae

DISTRIBUTION

The Texas Toad occurs from southwestern Oklahoma south through Texas to the Gulf Coast. It occurs north into New Mexico along the Pecos River drainage, and southward in Chihuahua and northeastern México. This species is found on Mustang Island, Texas (Moore, 1976). Impor­tant distributional references include Smith and Sanders (1952), Degenhardt et al. (1996), Dixon (2000, 2013), Sievert and Sievert (2006), López et al. (2009), Tipton et al. (2012), and Davis and LaDuc (2018). FOSSIL REC­O RD

Fossils of this species have been reported from Miocene deposits in Nebraska, Pliocene deposits in Kansas, and Pleistocene deposits in Texas (Tihen, 1962b; Holman, 2003). According to Holman (1969, 2003), some of the fossil rec­ords now referring to this species ­were originally identified as belonging to A. cognatus. ­These species are closely related, and the slight osteological differences between them make specific identification tentative. SYSTEMATICS AND GEOGRAPHIC VARIATION

Anaxyrus speciosus is most closely related to A. cognatus within the Nearctic clade of North American bufonids (Pauly et al., 2004). ­These species form the Cognatus clade together with A. compactilis of México, but this clade is considered paraphyletic by Pauly et al. (2004). Cope (1889) and Smith (1947) regarded A. speciosus as a subspecies of A. compactilis, but evidence of their specific distinctiveness is based on calls (Bogert, 1960), morphol-

ogy (Rogers, 1972), and protein polymorphism (Rogers, 1973). Laboratory crosses between A. speciosus and A. californicus, A. cognatus, A. woodhousii, A. microscaphus, or Incilius nebulifer can produce successful metamorphs (A.P. Blair, 1955; Blair, 1959), but not always (Moore, 1955). The color and pattern of the hybrids is intermediate between ­those of the parents, but developmental abnormalities are common among larvae that do not survive. Crosses between A. terrestris or A. punctatus and A. speciosus ­were not successful (Moore, 1955; Blair, 1959). The sex of the parents is impor­tant in determining ­whether hybrid crosses ­will be successful. Some hybridization experiments also appear to rely on very small sample sizes. ADULT HABITAT

The Texas Toad is a species of the short-­grass plains, where it prefers sandy soils. In the Sierra Vieja Range of southwestern Texas, Jameson and Flury (1949) found it associated with the salt cedar–­mesquite and catclaw–­creosote bush plant associations in low, sandy washes and along the banks and floodplain of the Rio Grande. In Big Bend National Park, A. speciosus is a toad of mesquite scrub vegetation; vegetation communities include lechuguilla, creosote, mesquite thickets, and mixed scrub (Dayton et al., 2004). In New Mexico, it occurs from 900 to 1,300 m (Degenhardt et al., 1996). TERRESTRIAL ECOLOGY

Most activity occurs at night when air temperatures are >17°C. Activity can occur any time of the year, at least in south Texas, temperature and precipitation permitting (Moore, 1976). For example, Minton (1958) recorded juvenile activity on 26 February in the Big Bend region. Rainfall is necessary to stimulate emergence in the spring. ­After metamorphosis, young toads remain within the vicinity of breeding pools for up to several months as long as the pools retain ­water. CALLING ACTIVITY AND MATE SE­L ECTION

Distribution of Anaxyrus speciosus

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Breeding choruses are often very large, with males sometimes moving about continuously calling and searching for females (Bragg, 1940b), or calling from prominent positions around the shoreline (Moore, 1976). Males precede females to the breeding sites and ­will amplex any toad within their vicinity. Sometimes this be­hav­ior results in a “seething mass” of struggling toads with up to 5 males piled on top of one another (Bragg, 1950b). According to Bragg (1945a), females ­will bypass males calling from very small pools in ­favor of

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t­ hose calling from nearby larger pools. The small pools undoubtedly have a much shorter hydroperiod for tadpole development than the larger pools, but how the female makes this discrimination is unknown. Calling occurs both night and day but is most intense at night. The call is a very short, loud, explosive trill with a dominant frequency of 2,400–2,800 cps, a duration of 0.4–0.6 sec, and a mean of 39–57 trills/sec at 23–27°C (Blair, 1956b). As in A. cognatus, amplexed male A. speciosus have a slow series of warning vibrations to alert a conspecific as to the sex of the grasped animal (Blair, 1947b). Wiest (1982) reported calling at air temperatures of 18.5–23.1°C. Females ­will move ­toward calling males via triangulation and ­will initiate amplexus by touching them (Axtell, 1958). At this time, the male stops calling and grabs the female. Satellite males in the vicinity ­will also attempt to amplex the female, but the largest male usually wins the strug­gle. If grasped by a small male, a female ­will attempt to dislodge him, although not always successfully. BREEDING SITES

Breeding occurs in both clear and muddy, shallow, temporary ponds and pools. Sites may be located in open fields or near streams, irrigation ditches, in ­cattle tanks, and even in buffalo wallows. Ponds should be open-­ canopied, and may be ­free of vegetation or contain a considerable amount of aquatic vegetation (Bragg and Smith, 1942). REPRODUCTION

Based on histology, Goldberg (2018b) noted that males and females are capable of breeding over an extended period from spring through late summer. The smallest mature male was 50 mm SUL in his sample; females may reach maturity at ca. 60 mm SUL. Breeding is opportunistic and occurs from spring throughout the summer when stimulated by heavy rainfall. For example, breeding has been observed in early April in Texas and as late as August in Oklahoma and September in Texas (Bragg, 1940b; Bragg and Smith, 1942; Moore, 1976; Wiest, 1982); most reproduction prob­ably occurs from May to July. The eggs are deposited in long strings in shallow pools. No data are available on clutch size, but ­there are 14–20 eggs per 30 mm of string (Wright and Wright, 1949). Degenhardt et al. (1996) reported additional data on reproduction in A. speciosus, but ­these observations ­were based on Bragg’s (1955) reports on “B. com­ pactilis” from southwestern Utah, which is actually A. microscaphus.

LARVAL ECOLOGY

Nothing is known of the larval ecol­ogy of this species except that larvae eat algae scraped from rocks. Metamorphosis occurs in as quickly as 18 days (Moore, 1976). DIET

The adult diet consists almost entirely of arthropods, particularly ants, lepidopteran adults and larvae, and many types of beetles; other prey include spiders, millipedes, centipedes, true bugs, weevils, flies, crickets, and homopterans (e.g., cicadas, aphids) (Smith and Bragg, 1949). Specific dietary items change somewhat between spring and summer, presumably in response to availability rather than se­lection. Juveniles eat mostly ants, beetles, and flies (Smith and Bragg, 1949). Texas Toads are reported to feed commonly ­under streetlights on insects drawn to the light. PREDATION AND DEFENSE

Tadpoles of Spea multiplicata (as S. hammondii) avoid attacking live or dead larvae of A. speciosus (Bragg, 1960a). Like many other toads, the Texas Toad is cryptically colored. When approached by a predator, it may crouch low against the substrate and only slowly seek cover ­under vegetation (Bragg, 1945a). No information is available on predators, but they likely include snakes, birds, and carnivorous mammals. POPULATION BIOLOGY

Growth is rapid during the first 3 months following metamorphosis, averaging 10 mm per month in Texas (Moore, 1976). Growth rates then decrease to almost zero during the cold winter months. It seems likely that sexual maturity is reached by the second spring following metamorphosis. The large disparity in size between males and females suggests that females reach maturity ­later than males, assuming growth rates are equal.

Adult Anaxyrus speciosus. Photo: Robert Hansen

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COMMUNITY ECOLOGY

Anaxyrus speciosus and A. cognatus are sympatric in the ecotone between short-­grass and mixed-­grass prairies, with A. speciosus replacing A. cognatus to the west (Bragg and Smith, 1943). The presence of A. speciosus larvae tends to inhibit the growth and survival of larvae of Gastrophryne olivacea and Gray Treefrogs (Dryophytes versicolor/​ D. chrysoscelis complex), but not A. houstonensis ­under laboratory conditions. The presence of A. woodhousii larvae, however, had no effect on A. speciosus (Licht, 1967). Decreases in body mass occur in larval A. speciosus when reared with Scaphiopus couchii larvae (Dayton and Fitzgerald, 2001). Data suggest that S. couchii are able to outcompete A. speciosus larvae and may exclude them from shallow ­water desert pools in some areas, such as Big Bend National Park. Breeding habitat of Anaxyrus speciosus. Photo: David Hillis DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus has been reported from this species in New Mexico (Suriyamongkol et al., 2019). The myxozoan Myxidium serotinum parasitizes A. speciosus (McAllister and Trauth, 1995) as do 2 species of cestodes and 3 species of nematodes (Kuntz, 1941). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available.

Anaxyrus terrestris (Bonnaterre, 1789) Southern Toad ETYMOLOGY

terrestris: from the Latin terrestris meaning ‘pertaining to the Earth.’ NOMENCLATURE

Conant and Collins (1998): Bufo terrestris Fouquette and Dubois (2014): Bufo (Anaxyrus) terrestris Synonyms: Bufo clamorus, Buffo clamosus, Bufo erythronotus, Bufo lentiginosa, Bufo lentiginosus, Bufo lentiginosus pachycephalus, Bufo musicus, Bufo rufus, Chilophryne lentiginosa, Incilius lentiginosus, Rana lentiginosa, Rana musica, Rana terrestris, Telmatobius lentiginosus IDENTIFICATION

Adults. This is a medium-­sized toad with coloration ranging from reddish to olive green to gray to nearly

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STATUS AND CONSERVATION

The Texas Toad appears to be widespread and abundant throughout its range, and no large-­scale population declines have been reported (Dayton and Painter, 2005). Some populations in the agricultural regions of south Texas may be declining due to pesticide use (Dixon, 2000). Considering its abundance, it is surprising that so ­little information is available on its life history.

black. Red toads in the South are usually this species. The head is relatively broad with a sharp snout; females have broader heads than males. Like the American Toad, ­there is usually 1 wart per spot, and the cranial crests are well developed and quite obvious. The cranial crests have a large knob at the posterior ends of the parallel interorbital crests that increases in prominence with age, and this feature is usually more prominent in females. The posterior portion of the crest has a relatively long extension called the preparotoid ridge that extends at right ­angles and touches the parotoid gland. Parotoids are of vari­ous shapes, from oval to elongate to kidney ­shaped. The white mid-­dorsal line is not well developed. The Southern Toad is usually larger and darker than the sympatric Fowler’s Toad. Dorsal warts are prominent. Venters may be spotted in the pectoral region or unmarked. Limbs are relatively short, with the dorsal surface of the rear legs barred. The rear toes are webbed, except for the end of the fourth toe that extends beyond the webbing. Males normally have a dark subgular vocal sac, but this is not always darkly colored. An albino was reported from Duval County, Florida (Dyrkacz, 1981).

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deposited in continuous, gelatinous strings with a singular tubular membrane and no intercellular partition between adjacent eggs. Strings may be singular or paired, with 8–12 eggs per 25 cm in length. ­There are 2 jelly envelopes surrounding the egg (outer envelope 2.6–4.6 mm; inner envelope 2.2–3.4 mm in dia­meter) (Brown, 1956), although upon deposition the inner envelope may be difficult to see. The vitellus mea­sures 1.0–1.4 mm in dia­meter and may be elliptical. The normal clutch size is from 2,500 to 3,000 eggs per female. DISTRIBUTION

Head pattern of Anaxyrus terrestris. Illustration by Camila Pizano

Males are smaller than females. On the Isle of Hope, Georgia, males averaged 46.9 mm SUL (range 27–74 mm) and females 47.2 mm SUL (range 23–80 mm), but on the adjacent mainland, males averaged 54.9 mm SUL (range 46–61 mm) and females 61.6 mm SUL (range 50–84 mm) (O’Hare and Madden, 2018). Males ­were 51–85 mm SUL (mean 64.5 mm) and females 55–123 mm SUL (mean 81.1 mm) in 1 south Florida population, and 50–70 mm SUL (mean 59.5 mm) and 55–90 mm SUL (mean 69.6 mm), respectively, in another population (Meshaka and Layne, 2015). In central Florida, males averaged 58.5 mm SUL (range 48–65 mm), whereas females averaged 73.8 mm SUL (Bancroft et al., 1983). In north central Florida, males ­were 43–70 mm SUL (mean 53 mm) whereas females ­were 43–79 mm SUL (mean 58 mm) (Dodd, 1994). Brown (1956) recorded an adult mean of 48 mm SUL (range 56.3– 75.4 mm) in Alabama. Means and Richter (2007) recorded a ­giant male of 150 mm SUL. Larvae. The larvae are small, ovoid in appearance, and black. Small, purplish dots may be pre­sent dorsally. Venters are black with somewhat purplish spots scattered freely and not forming a continuous mass. The upper tail fin is somewhat spotted, but the lower tail fin is unspotted and clear to yellowish; tail fins are about equal in depth but wider than the tail musculature. The lower edge of the tail musculature is light cream or light yellow. The tail is short with a rounded tip. The eyes are positioned dorsally and they are close together. According to Dundee and Rossman (1989), ­there is a light oblique mark ­behind each eye. The body of the tadpole is 5–10 mm in length; total lengths approach 24–28 mm. Descriptions of larvae are in Wright (1929) and Siekmann (1949). Eggs. The eggs of A. terrestris are black dorsally and grayish white ventrally (Livezey and Wright, 1947). Eggs are

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The Southern Toad occurs on the Atlantic and Gulf Coast Coastal Plain from southeastern ­Virginia through eastern Mississippi, and southward through the Florida Parishes of Louisiana. An isolated population occurs in the upper Piedmont on the border of Georgia and South Carolina (e.g., Chamberlain, 1939). Most rec­ords of Southern Toads, however, are below the Fall Line. Populations of Southern Toads are found in the Florida Keys (Big Pine, Cudjoe, Sugarloaf: Duellman and Schwartz, 1958; Lazell, 1989) and on vari­ous Atlantic and Gulf Coast barrier islands. ­These include Hatteras, Bodie, and Smith islands, North Carolina (Lewis, 1946; Gaul and Mitchell, 2007; Parlin et al., 2019), Kiawah Island, South Carolina (Gibbons and Coker, 1978; Hanson and McElroy, 2015), Isle of Hope, Sapelo, ­Little Cumberland, and Cumberland islands, Georgia (Martof, 1963; Gibbons and Coker, 1978; Shoop and Ruckdeschel, 2006; O’Hare and Madden, 2018), St. George and St. Vin-

Distribution of Anaxyrus terrestris

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cent islands, Florida (Irwin et al., 2001), and Cat Island, Mississippi (M.J. Allen, 1932). Impor­tant distributional references include: Alabama (Brown, 1956; Mount, 1975), Florida (Lazell, 1989; Bartlett and Bartlett, 1999; Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007), South Carolina (Dodd and Barichivich, 2017; Fields, 2019), and ­Virginia (Tobey, 1985; Mitchell and Reay, 1999).

Laboratory experiments have demonstrated hybridization capability between ♀ A. terrestris and ♂ A. hemiophrys, A. houstonensis, A. punctatus, A. microscaphus, A. woodhousii, and Incilius nebulifer (Blair, 1959, 1961a, 1963a). Crosses between ♀ A. punctatus or A. woodhousii and ♂ A. terrestris are generally infertile, and ­those between ♀ I. coccifer and ♂ A. terrestris may produce larvae but few if any metamorphs (Blair, 1959, 1961a). A cross between a ♀ A. terrestris and a ♂ A. speciosus (as A. compactilis) was unsuccessful (Blair, 1959). Some hybrids are fertile and capable of backcrossing with parental species, such as the F1 progeny of ♀ A. terrestris and ♂ A. hemiophrys or A. woodhousii (Blair, 1963a).

FOSSIL REC­O RD

Fossil Southern Toads are known from Pleistocene Irvingtonian sites in Florida and Pleistocene Rancholabrean sites in Florida, Georgia, and Tennessee. The species is particularly common in Florida fossil deposits (Holman, 2003). Large knobs on the fronto-­parietal bone distinguish this species from other fossil Anaxyrus within its range. The dorsal pro­cess of the ilium is lower than that of A. americanus. Tihen (1962 a, b) and Holman (2003) note other distinguishing osteological characteristics among A. fowleri, A. americanus, and A. terrestris. Meylan (2005) named a new species of late Pliocene toad that appears to be related to A. terrestris as Bufo defensor (= Anaxyrus defensor). Anaxyrus defensor has the largest supraorbital crest on the fronto-­parietals of all New World bufonids. SYSTEMATICS AND GEOGRAPHIC VARIATION

Anaxyrus terrestris is a member of the Americanus group of toads, which also includes A. americanus, A. houstonensis, A. microscaphus, A. fowleri, and A. woodhousii. This group in turn is a member of the Nearctic clade of North American bufonids (Pauly et al., 2004). Based on molecular analy­sis, the species is most closely related to the southern clade of A. fowleri (Masta et al., 2002), a relationship that is dif­fer­ent from ­earlier phyloge­ne­tic hypotheses based on morphology and call similarity (e.g., Blair, 1963b). Divergence prob­ably took place within the last 2 my during the Pleistocene. Natu­ral and experimental hybridization has been reported with A. fowleri (Gosner and Black, 1956; Volpe, 1959b; Brown, 1969) and A. americanus (Wilbur et al., 1978). In southeastern Louisiana and adjacent Mississippi, Volpe (1959b) demonstrated extensive geo­graph­i­cal areas of hybridization between A. terrestris and A. fowleri. In some populations, “hybrid swarms” (individuals with an extensive diversity of character combinations) of toads ­were observed, whereas in other populations repeated backcrossing of hybrids with parental species was apparent. Thus, the extent of hybridization and its effects on phenotype vary a ­great deal in some areas, making identification difficult.

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ADULT HABITAT

Southern Toads occur in a wide variety of terrestrial and semiaquatic habitats, including seashore scrub, seashore dunes, pine flatwoods, scrub, sandhills, xeric hammock, dome swamps, coastal hydric hammock, inland hydric hammock, basin swamps, depression marshes, forested wetlands, basin marshes, steephead ravines, wet prairies, upland hardwood forests, and mixed-­upland forests (Carr, 1940a; Harima, 1969; Buhlmann et al., 1993; Enge et al., 1996; Enge, 1998a, 1998b; Enge and Wood, 1998; Smith et al., 2006; Baxley and Qualls, 2009; Meshaka and Layne, 2015; Chandler et al., 2015a; Erwin et al., 2016). They occur in ruderal and disturbed areas, such as sand pine plantations, other types of silviculture, agricultural areas, and urban and suburban habitats (Carr, 1940a; Neill, 1950a; Surdick, 2005; Alix et al., 2014b). According to Dundee and Rossman (1989), Southern Toads occupy dryer and higher sites when they are sympatric with A. fowleri. TERRESTRIAL ECOLOGY

Southern Toads are active year-­round, weather permitting, especially in the southern portion of the species’ range. Rainfall is not necessary to stimulate terrestrial activity, and in extreme drought toads may remain in terrestrial refugia rather than chance movements to what may be a dry breeding site (Dodd, 1994). During cold or dry weather, they take refuge in mammal burrows and holes, ­under surface debris and leaf litter, in rock piles, or buried into the soil. They seek shelter during the hottest part of the day in leaf litter or burrowed into the substrate. They ­will also occupy gopher tortoise (Gopherus polyphemus) burrows and stump holes in the South (Dziadzio and Smith, 2016; Murphy et al., 2021). Sometimes they are found with just the eyes and top of the head protruding above the substrate surface. Most activity occurs at night, with breeding migrations occurring almost exclusively at night (Todd and Winne, 2006). However, they are not immune to diurnal activity and are sometimes found hopping around during the ­middle of the day.

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Southern Toads are familiar with their environment, and some individuals are able to return to a release point even ­after being displaced as far as 1,609 m from the point of collection (Bogert, 1947). The direction of displacement does not appear to influence homing ability, with an estimated 50% of the toads eventually returning from a 640 m displacement over a period of several months. However, Bogert (1947) reported 1 individual traveling 790 m in a 24 hr period during favorable rainy conditions. Southern Toads are frequently found far from the nearest breeding ponds. In north central Florida, for example, toads ­were captured at a mean distance of 515 m from ­water (range 46–914 m) (Dodd, 1996). Southern Toads are monotonically photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). As such, Southern Toads likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Male A. terrestris have a loud and melodic (“musical,” Dundee and Rossman, 1989) trill; a chorus can be deafening. The calling period is extended throughout the spring and summer, but calls can be heard from February to October in southern Florida (Meshaka and Layne, 2015). Call characteristics in Texas include a mean pulse rate of 51.8–66 pulses/sec, a call duration of 6.9–8.8 sec, and a dominant frequency of 1,878–2,042 cps (Brown, 1969). In Florida, call characteristics include a pulse rate of 68–78 pulses/sec, a call duration of 1.5–8.3 sec, and a dominant frequency of

Amplexing Anaxyrus terrestris. Photo: Matt Greene

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2,000–2,300 cps (Blair, 1956b). Whereas pulse rates are lower and call duration is much longer than sympatric A. fowleri, the dominant frequencies are about the same. The advertisement call serves to attract females, and call differences help maintain species isolation. Anaxyrus terrestris has a release call that serves to alert both conspecific and heterospecific males that the amplexed animal is a male. Indeed, the release calls may be convergent in call structure so as to broaden their applicability even as mate advertisement calls diverge in call structure to allow for species isolation (Leary, 2001). The convergence centers on the periodicity of release vocalizations, which are not significantly dif­fer­ent among some toad species. However, character displacement in the periodicity of the advertisement calls of A. fowleri is pronounced both in sympatry and allopatry with A. terrestris. Advertisement periodicity is much higher in A. terrestris (mean 16.8–18.3 ms) than in A. fowleri (mean 7.3 ms). Males call from a variety of open locations, usually along the shoreline of a breeding site. They may face ­either the wetland or the shoreline, depending upon the positioning of aquatic vegetation. However, Wright (1932) recorded them calling perched on cypress logs, cypress knees and stumps, resting on aquatic plant stems, and sitting in shallow ­water along a pond margin. They do not prefer sites with obstructing vegetation, such as thick cattail. Males generally do not float while calling. Rainfall is not necessary to stimulate calling, but particularly large choruses call loudly during and ­after heavy rains. As the season progresses, rainfall may be more of a requirement to stimulate calling. Calling occurs by night or day. Wright (1932) recorded calling at temperatures of 14–31°C, and Dundee and Rossman (1989) noted that temperatures had to be >18°C in Louisiana for calling to occur. Mate se­lection and oviposition sequence are essentially the same as in A. americanus. Amplexus is axillary or supra-­ axillary, depending on the size of the female. Females are attracted by the calls of the males and move ­toward them, although they may be amplexed at any time by a satellite male or indeed by any male that intercepts them on the way to a breeding site. The oviposition sequence and the be­hav­ior of the courting pair have been described by Aronson (1944) and consist of amplexus, pre-­ovulatory be­hav­ior (restlessness and abdominal muscular contractions lasting from a few minutes to several hours), and oviposition (back arching, extrusion of eggs with simultaneous fertilization, movement along the wetland bottom as egg strings are extruded). If the egg strings break during deposition, amplexus ends, and the mating sequence is ­either reinitiated with the same or a dif­fer­ent male. A recently spent female ­will arch her back in a very pronounced manner to indicate she has finished

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spawning, although it may take several tries before the male releases her and moves elsewhere. Mating male Southern Toads may try to amplex virtually any species in their vicinity, from Southern Leopard Frogs and Cane Toads to large hylids (e.g., Schuman and Bartoszek, 2019b). Occasionally, males may be able to displace other males already amplexing a female, although the extent to which this occurs in nature and its significance remains unquantified (Lamb, 1984a). BREEDING SITES

Southern Toads breed in a variety of wetlands, from small, isolated, temporary pools and ponds to the shorelines of large permanent lakes. They breed in old field ponds, stream overflow basins, marshes, cypress savannas and cypress/gum ponds, and even in the quiet ­waters of shallow Coastal Plain streams (Wright, 1932; Liner et al., 2008). They readily breed in man-­made habitats, such as suburban retention ponds, roadside ditches, farm ponds, road ruts, borrow pits, and golf course ponds (Scott et al., 2008). Isolated ponds may be particularly impor­tant to reproduction and perhaps as areas of refuge and feeding. In South Carolina, for example, Russell et al. (2002a) captured nearly 2,000 A. terrestris at 5 wetlands 2,000 Southern Toads at 2 ponds over a 2 yr period, and Dodd (1992) captured 331 dif­fer­ent toads over a 5+ yr period in a very small depression marsh in central Florida. Mass metamorphosis can result in hundreds of thousands of recent metamorphs si­mul­ta­neously moving across a landscape (referred to in Florida as a “jubilee”). However, this does not necessarily mean that females produce large numbers of juveniles ­every year or at ­every breeding site. For example, Semlitsch et al. (1996) reported 816 females captured over a 16 yr period at a pond in South Carolina, but only 693 metamorphs. In Florida, Greenberg and Tanner (2005b)

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captured 1,321 adult Southern Toads at 8 ponds over a 7 yr period, and 1,359 metamorphs. However, in both studies, most of the metamorphs resulted from a single breeding year. ­There was no correlation between the number of adults using a breeding site and the number of juvenile recruits to the population. The total number of breeding adults is positively associated with juvenile recruitment (Greenberg et al., 2017a). Average air temperature does not influence juvenile recruitment. Populations of this species vary widely in abundance among years and breeding ponds, making ­simple trend analyses difficult at the landscape level (Greenberg et al., 2018a). For example, ­these authors found that recruitment was not correlated with the abundance of adult A. terrestris breeding populations in subsequent years. Low statistical power hampered an ability to detect overall trends, even with 24 yrs of monitoring data. COMMUNITY ECOLOGY

Anaxyrus fowleri and A. terrestris may occasionally be found breeding in the same pond (e.g., Gosner and Black, 1956). Experiments have shown that competition occurs between the larvae of ­these species at high and medium densities, but not at low densities (Wilbur et al., 1983). Competition occurs ­because of the similarities between the species in feeding ecol­ogy and habitat preference. High densities also affected the size differential between the larvae of the 2 species and the length of the larval period (Wilbur et al., 1983). Thus, the amount of habitat available and the timing of breeding become impor­tant to larval ecol­ogy if the 2 species occupy the same breeding pond. The presence of a predator, such as Notophthalmus, can actually minimize the effects of competition among anuran larvae by eating many small larvae and thus reducing larval densities. Larval A. terrestris are intense competitors with larval Pseudacris crucifer and Dryophytes gratiosus (Morin, 1983). On the other hand, A. terrestris larvae are sensitive to the density of larvae of other anurans, not just toad larvae. This sensitivity is manifested in slower growth rates depending on density, although most individuals ­will survive and metamorphose. Plasticity in the size at metamorphosis also helps mediate conditions within the breeding site. If competition becomes too intense, larvae can metamorphose at a smaller size than they might normally. Southern Toads also may be impacted by nonindigenous species, such as the Cuban Treefrog (Osteopilus septentrionalis). The presence of larval O. septentrionalis ­causes reduced growth rates and delayed metamorphosis in A. terrestris, at least ­under experimental ­trials (Smith, 2005a). In addition, the body mass of A. terrestris larvae at metamorphosis is decreased in the presence of Cuban Treefrog

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larvae. ­These species are syntopic in peninsular Florida, often using the same temporary, small breeding sites. DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytridiomycosis was not found on A. terrestris from individuals collected in South Carolina from 1940 to 1970 (Daszak et al., 2005), nor was it known from elsewhere in the Southeast through 2008 (Rothermel et al., 2008). It is now known from individuals in Alabama, North Carolina, and Florida (Rizkalla, 2010; Chiari et al., 2017; Lentz et al., 2021). Based on experimental ­trials, this species is very prone to mortality via chytrid infection (Searle et al., 2011). The fungus Basidiobolus sp. is known from Southern Toads from Florida (Okafor et al., 1984). Perkinsea, an alveolate pathogen, has been reported from Florida (Karwacki et al., 2018). Ranavirus is reported from North Carolina (Lentz et al., 2021). Sauer et al. (2019) found significant individual-­level variation in temperature preference and evidence for behavioral fever in both metamorphic and adult A. terrestris during the first 2 days ­after exposure. Individual-­level change in temperature preference was negatively correlated with ranaviral load and was a better predictor of load than average temperature preference or maximum temperature reached by an individual. Their results suggested that behavioral fever is an effective mechanism for resisting ranaviral infections in this species. Southern Toads are parasitized by the cestodes Cylindrotaenia americana, Distoichometra bufonis, and Mesocestoides sp., the trematodes Brachycoelium hospitale and Megalodiscus temperatus, and the nematodes Cosmocercoides variabilis, Gyrinicola batrachiensis, Oswaldocruzia pipiens, and Rhabdias americanus (Dickey, 1921; Manter, 1938; Pryor and Greiner, 2004; McAllister et al., 2015b). Biting midges of the genus Corethrella feed upon A. terrestris and may be attracted by the frog’s advertisement call (McKeever and French, 1991). The mite Hannemania hegeneri also is reported from this species (McAllister et al., 2015b). Polyphalangy and missing eyes have been reported from A. terrestris in Mississippi (Kreiser et al., 2016). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Nitrate. Nitrates in ­water can affect the duration of the larval period. High levels of nitrates (30 mg/L NO3-­N, or even fluctuations between 0 and 30 mg/L NO3-­N) act as a stressor, speeding up developmental rates of Southern Toads and causing larvae to metamorphose at smaller sizes than they would if not subjected to nitrates (Edwards et al., 2006). However, other chemicals in natu­ral ­water also may act to decrease growth rates and thyroxine concentrations, and nitrates may help modify a tadpole’s response to

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environmental stressors. Thus, the type of ­water (e.g., spring ­ ater versus reverse osmosis filtered ­water) in which w laboratory studies are conducted is impor­tant when assessing the effects of nitrates on larval development. Metals. Coal ash settling basins may have high concentrations of toxic chemicals, particularly arsenic, cadmium, chromium, copper, selenium, and other substances. In a series of transplant experiments, Rowe et al. (2001) demonstrated that Southern Toad larvae raised in such ­waters have a high rate of mortality compared to reference sites. This result involved both embryos and larvae, and stemmed from the toxic effects of the chemicals and the depauperate algal food sources at the coal ash site. If embryos survived to hatching (about 34%), then they had nothing to eat and ­were increasingly exposed to the toxic mix. Rowe et al. (2001) suggested that such basins could be population sinks for ­those species attempting to use them for breeding. Toxic ele­ments at coal ash settling basins also concentrate in adult toads frequenting ­these sites. Increased levels of selenium (17.4 ppm), arsenic (1.58 ppm), and vanadium (1.24 ppm) are observed in Southern Toads collected from coal ash basins, and transplant experiments suggested that ­these chemicals can be concentrated in significant amounts ­after only 7–12 weeks of exposure (Hopkins et al., 1998). ­These results indicate that Southern Toads bioaccumulate toxic substances and that bioaccumulation can occur rather rapidly. Elevated levels of toxic substances in coal ash settling basins can lead to increased levels of corticosterone B in resident toads, ­whether they are calling or not. Testosterone levels ­were also higher in all months than at control sites, suggesting altered androgen production, utilization, and clearance. Males called into August at ash basins but not at control sites, suggesting that the toxic substances affect hormone levels that in turn extends calling be­hav­ior to ­later in the season. Elevated levels of corticosterone B ­were evident in as ­little as 10 days when toads ­were transplanted from control sites to coal ash settling basins (Hopkins et al., 1997). Byproducts of coal ash waste sites can be transferred from adult females to their eggs, resulting in a 27% reduction in reproductive success (Metts et al., 2013). Metts et al. (2013) found maternal transfer of the toxic ele­ments lead, copper, selenium, and strontium to the eggs of contaminated females. Reproductive success was negatively correlated with selenium and copper concentrations in females, and selenium concentrations in eggs. Tadpoles can also accumulate significant levels of trace ele­ments from the substrate of contaminated sites, which in turn reduces tadpole survivorship and growth rates, extends larval periods, and reduces mass at metamorphosis (Metts et al., 2012). The interactive effects of maternal transfer and substrate-­based toxic ele­ment accumulation reduce tadpole survival to metamor-

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phosis by as much as 85%. Breeding sites affected by coal combustion wastes are thus ecological traps for this species and other pond breeding amphibians. Weir et al. (2016a) found no significant effect on A. terrestris embryo mortality from sublethal levels of zinc, and also that the presence of zinc had no interactive effect with copper on embryo mortality. They suggested that perhaps their findings differed from previous experiments ­because they exposed the metals to embryos rather than larvae, and that they used sublethal levels of zinc. Copper toxicity to embryos and larvae alone had no significant effects on extinction risk ­unless toxicity was very high. However, copper toxicity coupled with catastrophic reproductive failure (>50%) resulted in high extinction risk, although ­there ­were no interactive effects (Weir et al., 2016b). The models developed by Weir et al. (2016b) ­were most sensitive to juvenile and adult survivorship, and highlighted the importance of coupling the effects from multiple stressors with long-­term species demographic differences in response to ­those stressors. Heat. Southern Toads oviposit eggs in thermally heated reservoirs associated with nuclear power plants. Most such eggs die ­because of thermal loading, and thus ­these “cooling” reservoirs become population sinks. However, some eggs and tadpoles may survive in shallow areas receiving cooling inflows from surrounding springs, seeps, or creeks. In ­these areas, temperatures may be sufficiently cool to allow development. However, resulting larvae tend to grow faster than conspecifics in unaffected surrounding areas, and the larvae transform at a smaller size than normal (Nelson, 1974). Salinity. In experimental ­trials, larval A. terrestris had 100% survival at salinities ≤ 5 ppt. However, no larvae survived at salinities of 14 or 16 ppt over a 72 hr period; ca. 80% survived at 10 ppt (Brown and Walls, 2013). STATUS AND CONSERVATION

Villena et al. (2016) suggested populations in the South ­were declining based on probability of occurrence through time, although ­there was much variation from state to state. Undoubtedly, many populations of Southern Toads have been lost as wetlands and adjacent uplands have been destroyed or modified over the last 500 yrs. Mortality also occurs from natu­ral events, such as tornados (Smith, 2013). This species appears to be somewhat resilient, however, and is still frequently observed in suburban and agricultural regions throughout the Southeast (Delis, 1993). Highway-­ related mortality must be very high, as ­these toads are frequently found on paved roads on warm rainy nights (M.J. Allen, 1932; Sutherland et al., 2010). Even in suburban settings, mortality can be high where roads are located next to retention ponds and toads are drawn to nearby streetlights where they congregate to feed. Southern Toads readily

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Anaxyrus terrestris 155

Anaxyrus terrestris breeding habitat, McIntosh County, Georgia. Photo: C.K. Dodd, Jr.

use culverts, when available, to traverse ­under highways. The placement of tunnels is crucial to their effectiveness in toad conservation inasmuch as toads may use topographical features such as lake basin rims to facilitate movement (Dodd et al., 2004). ­There are a variety of other ­factors that likely influence Southern Toad populations, including climate change. ­There is growing evidence that changes in rainfall patterns in the Southeast may have affected reproduction. Despite regularly stable numbers of immigrating females, for example, some populations produced no (Dodd, 1994, over a 5 yr period) or few metamorphs as hydroperiods decreased or ponds did not fill (Semlitsch et al., 1996; Daszak et al., 2005; Greenberg and Tanner, 2005b). More data are needed from long-­term studies to determine what level of reproduction is “normal” (i.e., in terms of frequency and recruitment) for a toad population to maintain itself through time. Silvicultural activities impact much of the habitat occupied by Southern Toads on the southeastern coastal plains. Long-­term effects in a variety of habitat types from vari­ous silvicultural treatments are few as long as suitable nearby habitat is available; A. terrestris may be one of the most common amphibians within its environment even in human-­disturbed habitats (Dodd et al., 2007). Results of studies that mea­sured short-­term effects on the numbers of toads captured within 1.5–2 yrs of cutting also have suggested ­little impact on the species at wetlands surrounded by vari­ous levels of treatment (Clawson et al., 1997; Russell et al., 2002b). The latter authors used counts rather than abundance to assess treatment effects, and the treatments ­were conducted in such a manner as to cause minimal disturbances to wetlands and to minimize surface disturbances even in cutover treatments. Toads are frequently found in open habitats created during forestry operations,

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such as in canopy gaps and along skidder trails (Cromer et al., 2002). Such areas may contain high insect abundance. Likewise, a study that followed the effects of clearcutting in a Florida pine flatwood also could not detect short-­term effects in the numbers of Southern Toads using the site (Enge and Marion, 1986). Southern Toads may even be found more often in previously ditched cypress ponds than around cypress ponds holding ­water (Vickers et al., 1985). In contrast, Hanlin et al. (2000) resurveyed an area studied by Bennett et al. (1980) and found many fewer toads than in the previous study (700 vs. 3,312) despite a more extended sampling period. Habitat changes associated with pond restoration may have influenced subsequent use by toads. As with previous assessments, relative abundance was assumed based solely on count data. Simply counting toads, however, provides an inexact picture of the short-­term effects of clearcutting. Southern Toads appear to tolerate prescribed burning and are usually found soon ­after burning (Floyd et al., 2002; Moseley et al., 2003; Klaus and Noss, 2016). Studies that provide data on habitat use and demographics of toad populations give a somewhat dif­fer­ent picture of the short-­term effects of silviculture, and Todd and Rothermel (2006) have argued for the importance of examining metrics other than abundance in determining site impacts on amphibians resulting from silviculture. For example, Southern Toads ­will use breeding ponds adjacent to or in clearcut habitats. Although juveniles are readily produced from such ponds, they have lower survivorship than conspecifics reared in forested habitats (Todd and Rothermel, 2006). In addition, growth rates are less in clearcuts than in forested areas and, as a result, juveniles found in clearcuts are smaller than ­those in adjacent forests. Southern Toads also use habitat differently in clearcuts than they do in forested habitats. They move farther in forests than in adjacent clearcuts, but movement patterns are rather similar regardless of sex, body size, or environmental cues (humidity, wind, air temperature, precipitation, soil moisture, sky conditions) (Graeter, 2005; Graeter et al., 2008). Movements occur in straighter paths in forested habitats than in clearcuts, where toads wander in no par­tic­u­lar direction. Southern Toads may treat large clearcuts as filters to extensive movements, but they do not avoid them entirely, and they readily cross the forest-­clearcut edge. In an experimental choice test, toads moved from 3.5 to 324 m (mean 46 m) over a 5 night period when released into a managed forest in South Carolina. Toads traversed small clearcuts easily within a 24 hr period (Graeter, 2005; Graeter et al., 2008). In large clearcuts, however, the increased searching be­hav­ior by Southern Toads could expose them to predation or desiccation, and discourage them from traversing the area.

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Anaxyrus terrestris readily colonizes restored or newly created mitigation wetlands (Pechmann et al., 2001). However, not all such ponds are successful in producing juveniles. Of the 4 ponds studied by Pechmann et al. (2001), only 1 pond produced substantial numbers of metamorphs, and then in only 3 of 8 years during which the pond was studied. Populations can recover quickly in some coastal areas that receive hurricane overwash with resulting temporary increases in salinity (Gunzburger et al., 2010).

Traffic noise may affect tadpole be­hav­ior since tadpoles detect sound through vibrations in the ­water. In a series of experiments, larvae increased their activity when exposed to real-­world simulated traffic noise, but ­there was no effect on subsequent size at metamorphosis (Castaneda et al., 2020). Other potential effects of loud anthropogenic noise are unknown. A total of 16,005 A. terrestris ­were exported from Florida from 1990 to 1994 for the pet trade (Enge, 2005a).

Anaxyrus williamsi (Gordon, Simandle, and Tracy, 2017) ­Dixie Valley Toad

DISTRIBUTION

Known only from the western edge of ­Dixie Valley Playa in ­Dixie Valley, Nevada. See map in A. monfontanus account. FOSSIL REC­O RD

ETYMOLOGY

williamsi: named for Robert Williams, US Fish and Wildlife Ser­vice field supervisor who has championed the conservation of fauna and flora of the ­Great Basin in Nevada and California. The common name refers to the location where it occurs. NOMENCLATURE

Synonyms: Anaxyrus boreas [in part], Bufo [Anaxyrus] williamsi IDENTIFICATION

Adults. The dorsal ground color is olive brown with irregular black flecks. Parotoids are tan. The dorsum is covered by many round and irregular glands giving a somewhat pebble-­like appearance, and ­there is a distinct dorsal stripe that is cream in color. The forearms are olive to medium to dark brown. The hind legs have rust-­colored tubercles with dark brown bands on an olive ground color. The throat is white, and the venter is heavi­ly mottled in black. The tubercles of the hands, feet, fin­gers, and toes are bright orange. Adults are superficially similar to A. boreas, but are characterized by a smaller body size, larger closely set eyes, a smaller head, a larger tympanum, and shorter hind limbs when compared with Western Toads. In a sample of 30, the mean SUL was 56.3 mm. In a ­later sample of 76, the mean was 54.6 mm SUL (range 44–70 mm) (Gordon et al., 2020). Additional morphological mea­sure­ments are in Gordon et al. (2020). Larvae. The larvae have not been described but are presumably similar in morphology to related toad species. Eggs. Unknown, but presumably similar to related toad species.

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None known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Gordon et al. (2017) described this species based on an extensive morphological and ge­ne­tic (­whole genomic DNA using the mitochondrial control region as the ge­ne­tic marker) analy­sis of this species in comparison with A. boreas, A. exsul, and A. nelsoni. Anaxyrus williamsi represents a unique and ­sister lineage to the Humboldt-­Lahontan lineage of A. boreas and the northern lineage of A. canorus. Ge­ne­tic evidence suggests a recent divergence from A. boreas, which corresponds to an age of aquatic isolation in ­Dixie Valley 650,000 yrs ago. Using mitochondrial DNA and microsatellites, Forrest et al. (2017) confirmed the uniqueness of this species and noted that it warranted conservation as a distinct management unit.

Anaxyrus williamsi tadpole. Photo: Alexa Killion and Kelsey Ruehling (USGS)

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ADULT HABITAT

This species is restricted to spring-­fed wetlands at the western edge of the ­Dixie Valley Playa. The known range is ca. 370 ha (Forrest et al., 2017).

parotoids and tibial glands contain bufotoxin that may deter predators. POPULATION BIOLOGY

Nothing is known. AQUATIC AND TERRESTRIAL ECOLOGY

Anaxyrus williamsi is nocturnal, inhabiting moist vegetation in shallow ­water with ­little vegetative canopy (Gordon et al., 2017). Activity occurs throughout the warmer portions of the season, and toads are very closely associated with ­water. This is especially true for males in spring and females in autumn (Halstead et al., 2021). They prefer warmer ­waters and substrates. When they are away from ­water, Halstead et al. (2021) found them at a median distance of 4.2 m (range 3.3–5.3 m) from ­water, and 95% of the time they ­were within 14 m of ­water. ­Little is known concerning life history during the nonbreeding portions of the year, but the work by Halstead et al. (2021) suggests they remain in or very close to ­water year-­round. In autumn at least, toads avoid bare ground and grasses. The mean home range in autumn was 974 m2 (range 46–5,363 m2), with a corresponding mean core activity area of 173 m2 (range 8–823 m2) (Halstead et al., 2021). Males moved an average of 9.2 m/day, and females 4.6 m/day. As winter approaches, they select dormancy sites in, over (i.e., within dense vegetation), or near ­water in springs where temperatures are stable. Toads overwinter in mud, under­ground, in dense bulrushes, or directly in ­water. ­Dixie Valley Toads enter dormancy from September to early November (Halstead et al., 2021) and exhibit much individual variation in the onset of dormancy.

DISEASES, PARASITES, AND MALFORMATIONS

None known at pre­sent, although amphibian chytrid fungus has been detected in American Bullfrogs at the southern edge of the toad’s range (Forrest et al., 2013). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

The isolation and extremely small range of this species suggest that it should be protected. Although the population

CALLING ACTIVITY AND MATE SE­L ECTION

Breeding occurs from March to June (Forrest et al., 2013) when males congregate in shallow ­waters among wetland vegetation. ­There is no advertisement call, but males have a release call when in contact with other males.

Juvenile Anaxyrus williamsi. Photo: Tim Burkhardt

BREEDING SITES

Breeding occurs in shallow ­water in the spring-­fed wetlands. REPRODUCTION AND LARVAL ECOLOGY

Eggs are deposited in shallow ­water where tadpoles develop in thermally suitable microhabitats. Metamorphosis occurs ­after 10 weeks. DIET

Nothing is known, but presumably insectivorous. PREDATION AND DEFENSE

Although the toads are distinctively olive colored, they are cryptic when hidden among wetland vegetation. Both the

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Adult Anaxyrus williamsi. Photo: Tim Burkhardt

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158  Bufonidae

Anaxyrus williamsi breeding habitat. Photo: Alexa Killion and Kelsey Ruehling (USGS)

Anaxyrus woodhousii (Girard, 1854) Wood­house’s Toad Crapaud de Wood­house ETYMOLOGY

woodhousii: a patronym honoring the first collector of this toad, S.W. Wood­house (1821–1904). Wood­house was the naturalist and Assistant Army Surgeon who accompanied Captain L. Sitgreaves on his expedition to the Zuni and Colorado rivers in 1851 during which the toad was collected (Wood­house, 1854). Wood­house had previously participated in western exploring expeditions with Sitgreaves in 1849– 1850 (Tomer and Brodhead, 1992). NOMENCLATURE

Conant and Collins (1998) and Stebbins (2003): Bufo woodhousii Fouquette and Dubois (2014): Bufo (Anaxyrus) woodhousii Synonyms: Bufo aduncus, Bufo antecessor, Bufo compactilis woodhousii, Bufo dorsalis, Bufo frontosus, Bufo lentiginosus frontosus, Bufo lentiginosus wood­housei, Bufo planiorum, Bufo velatus, Bufo wood­housei, Bufo wood­ housei australis, Bufo wood­housei bexarensis, Bufo woodhousii velatus, Incilius wood­housei This species was described as Bufo dorsalis, but this name had been used previously and was thus unavailable for the toad. The name woodhousii is attributed to Girard (1854).

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size is unknown, it is likely very small. Pos­si­ble threats include development for geothermal energy, groundwater depletion, habitat loss, and the introduction of amphibian chytrid fungus via nonnative American Bullfrogs (Forrest et al., 2013). ­Because of the species’ near total de­pen­dency on spring ­water, any action that affects groundwater or hydroperiod could threaten this toad with extinction (Halstead et al., 2021). This species is considered a Sensitive Species in Nevada and was petitioned in 2017 for protection ­under the federal Endangered Species Act of 1973 (ESA). In April 2022, the species was listed as Endangered ­under emergency provisions of the ESA (USFWS, 2022). The listing allows for federal protection for 240 days ­until a formal proposal can be prepared, and was prompted by an imminent threat from a geothermal development proj­ect.

IDENTIFICATION

Adults. This is a medium to large, dry-­skinned, brown to tan to olive toad with small, dark brown to black irregular spots containing 1 wart per spot, although some animals may be greenish to gray. Light brown warts are observed within the darker blotches, and ­these blotches tend to increase in number with the size of the toad (Keeton and Carpenter, 1955). Parotoids are oblong, elongated, narrow, and elevated, and touch the cranial crests. A light mid-­dorsal stripe is pre­sent. Venters are mostly white to yellowish and unmarked, except that some animals may have a spot or spots in the pectoral region or on the anterior third of the venter (Blair, 1943a). Males have con­spic­u­ous dark throats during the breeding season, but ­these lighten considerably during the remainder of the year except in the Southwest; ­here, throats are dark year-­round in both sexes. Males also develop nuptial pads. A hypomelanic/leucistic adult was reported from Arizona (Stack­house and Foster, 2020). Adult males are usually slightly smaller than females. In Arizona, adults ­were 49–91 mm SUL (mean 74.5 mm) (Goldberg et al., 1996b) in 1 study, with males in another population having a mean of 83–88.1 mm SUL and females a mean of 93.2–104.2 mm SUL, depending on year (­Sullivan, 1987). Blair (1941a) provided mean SULs for adults from 9 populations; toads ranged from 62 to 81 mm. Other specific size mea­sure­ments include: females with a mean of 83.3 mm SUL from Texas (Meacham, 1962); a female mean of 88.8 mm SUL in Oklahoma (Bragg and Sanders, 1951); female means of 97.3 mm SUL (range 64–113 mm) and male means of 73.5 mm SUL (58–69 mm) in Nebraska (Ballinger et al., 2010); and male means of 55.2–64.4 mm SUL in southwestern Utah and adjacent Nevada (A.P. Blair, 1955).

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Anaxyrus woodhousii 159

Head pattern of Anaxyrus woodhousii. Illustration by Camila Pizano

Conant and Collins (1998) and McGinnis and Stebbins (2018) reported a maximum size of 127 mm SUL. Larvae. Tadpoles are small and very dark brown to gray or slate in coloration, often with light mottling or gold flecking. The dorsal musculature of the tail is lighter than the body, and the ventral musculature is immaculate. The tail fins have a few scattered dark flecks that are more numerous on the dorsal tail fin than on the ventral tail fin. The tail tip is rounded. Venters are lighter than the dorsum. At hatching, the larvae are tiny (2.5–3 mm) but reach a maximum size of 23 mm TL (Youngstrom and Smith, 1936). Stuart (1991) reported amelanistic tadpoles from New Mexico. Eggs. Eggs are deposited in 2 long strings within a continuous gelatinous casing with no partitions between the eggs. Within the strings, eggs are normally in single rows, although they occasionally may be “crowded” together (Smith, 1934). The vitellus mea­sures 1–1.5 mm in dia­meter and is black dorsally and tan to yellow ventrally. ­There is a single gelatinous capsule surrounding the vitellus that mea­sures 2.6–4.6 mm in dia­meter (mean 3.5 mm) (Livezey and Wright, 1947). DISTRIBUTION

Wood­house’s Toad is a species of the ­Great Plains and semi-­arid west and is absent from the Rocky Mountains, but it also appears to be expanding its range in the deserts of California and Nevada. It occurs from North Dakota and Montana southwards to the Gulf Coast in eastern Texas, and its range covers much of northern and central Texas, Utah and western Colorado, northern Arizona, and eastern and central Wyoming, Colorado, and New Mexico. In Missouri,

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Distribution of Anaxyrus woodhousii. The hatched area indicates an extensive area of hybridization with A. fowleri. It may not be pos­si­ble to assign specific status to toads in this area.

Wood­house’s Toad is found in the western part of the state, but the range extends along the Missouri River east nearly to the Mississippi River. Populations extend along the ­middle Colorado River into Nevada (once including the Vegas Valley; Stejneger, 1893) and extreme northwestern Arizona. Its range includes the Gila and lower Bill Williams river drainages in western Arizona, and isolated populations occur in Ash Meadows, Nevada, and along the Amargosa River drainage basin in California and Nevada (Greene and Branston, 2013; Bleich, 2020, 2021), in the Coachella Valley of California (Goodward and Wilcox, 2019), in Washington and Idaho (Snake and Columbia rivers; Bear River in southern Idaho), along the lower Colorado River in California, Arizona, and México, and along the lower Rio Grande and southern Texas. The species overlaps broadly with A. fowleri in east Texas and Oklahoma, southwestern Arkansas, and Louisiana; hybridization may be extensive and toads from this region may actually include genes from A. fowleri and A. americanus as well as A. woodhousii (Pauly et al., 2004). It overlaps with A. microscaphus in central Arizona and west central New Mexico, where hybridization also may occur. Impor­tant distributional references include: Arizona (Eaton, 1935; Vitt and Ohmart, 1978; Brennan and Holycross, 2006; Bezy and Cole, 2014; Murphy, 2019; Holycross et al., 2021), California (Vitt and Ohmart, 1978; Greene and Branston, 2013; Goodward and Wilcox, 2019; Bleich, 2020,

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2021; Flaxington, 2021), Colorado (Hammerson, 1999), Idaho (Slater, 1941b; Nussbaum et al., 1983; Mulcahy et al., 2002), Kansas (Smith, 1934; Collins, 1993; Collins et al., 2010), Missouri (Johnson, 2000; Daniel and Edmond, 2006), México (López et al., 2009), Montana (Black, 1970, 1971), Nebraska (Lynch, 1985; Ballinger et al., 2010; Fogell, 2010), Nevada (Stejneger, 1893; Linsdale, 1940; Bradford et al., 2005a; Bleich, 2021), New Mexico (Van Denburgh, 1924; Degenhardt et al., 1996), North Dakota (Wheeler and Wheeler, 1966; Hoberg and Gause, 1992), Oklahoma (Sievert and Sievert,2006), Oregon (Nussbaum et al., 1983; Leonard et al., 1993; Jones et al., 2005), South Dakota (Peterson, 1974; Fischer, 1998; Ballinger et al., 2000; Kiesow, 2006), Texas (as a mixture of species and subspecies: Dixon, 2000, 2013; Tipton et al., 2012), Utah (Tanner, 1931; Mulcahy et al., 2002), Washington (Slater, 1939b, 1955; Metter, 1960; Nussbaum et al., 1983; Leonard et al., 1993; Jones et al., 2005; McAllister, 1995; Corkran and Thoms, 2020), and Wyoming (Baxter and Stone, 1985). FOSSIL REC­O RD

Fossil A. woodhousii have been reported from Miocene deposits in Arizona, Pliocene deposits in Arizona, Kansas, and Texas, late Pliocene–­early Miocene deposits in Kansas, and Pleistocene deposits in Arizona, Colorado, Kansas, Nevada, New Mexico, South Dakota, and Texas (Tihen, 1962b; Holman, 2003). The species may be referred to as Bufo cf. B. woodhousii in the fossil lit­er­a­ture. A very large (100–160 mm SUL) fossil subspecies, Bufo woodhousii bexarensis, was described from the late Pleistocene of Texas (Mecham, 1958). SYSTEMATICS AND GEOGRAPHIC VARIATION

Anaxyrus woodhousii is a member of the Americanus clade of North American bufonids (Blair, 1963b), a group that includes A. americanus, A. baxteri, A. fowleri, A. hemiophrys, A. houstonensis, and A. terrestris. It is closely related to A. americanus and somewhat more distantly to A. fowleri and A. terrestris (Masta et al., 2002). Allopatric divergence among ­these lineages prob­ably occurred during the Pleistocene. Anaxyrus woodhousii evolved in the ­Great Plains and expanded into the southwest during the mid to late Pleistocene. Masta et al. (2003) reviewed the evolutionary history of A. woodhousii and its expansion to its pre­sent geographic distribution. ­There are 2 distinct clades with A. woodhousii, 1 in the southwestern United States originally described as Bufo woodhousii australis (Shannon and Lowe, 1955) and the other comprising the remainder of the range (Masta et al., 2003). Anaxyrus w. australis has distinct narrow interorbital ridges separated by a frontal trough, whereas in A. w.

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woodhousii the interorbital ridges are shallow and in contact, and ­there is no frontal trough. The Southwestern Wood­house’s Toad occurs from south central Colorado through New Mexico and Arizona, and along the Rio Grande of southwestern Texas. The distribution of A.w. australis is poorly defined over much of its northern and eastern distribution. Many authors (e.g., Brennan and Holycross, 2006) treat it as a subspecies of A. woodhousii, although A. w. australis may be considered a full species if supporting data become available. The subspecies A. w. velatus was described by Bragg and Sanders (1951) and elevated to specific status by Sanders (1986). The taxon was said to be smaller, less spotted, and darker than A. fowleri, to possess a dif­fer­ent call, and to have a pectoral velum (a darkened area ventrally between the front limbs). This phenotype was confirmed by Masta et al. (2002) as representing a hybrid population between A. woodhousii and A. fowleri. It is not currently recognized, despite the phenotypic-­based arguments in Dixon (2000). Although Sanders (1987) described 2 additional species (as Bufo antecessor and B. planiorum) based on morphological data, they are clearly referable to A. woodhousii, and their recognition is not supported by molecular data. Natu­ral hybridization with other Anaxyrus species occurs throughout the range of A. woodhousii and has been reported with A. americanus (Blair, 1956a; Ideker, 1968; Brown, 1970), A. cognatus (Gergus et al., 1999), A. fowleri (Meacham, 1962), A. houstonensis (Brown, 1971b; Hillis et al., 1984), A. microscaphus (A.P. Blair, 1955; ­Sullivan, 1986a, 1995; ­Sullivan and Lamb, 1988; Goldberg et al., 1996b; Lamb et al., 2000; Schwaner and ­Sullivan, 2009; ­Sullivan et al., 2015; Wooten et al., 2019), A. punctatus (McCoy et al., 1967; Malmos et al., 1995; Hammerson, 1999), Incilius alvarius (Gergus et al., 1999), and I. nebulifer (Thornton, 1955; Brown, 1971a; Sanders, 1986). Although hybridization occurs “naturally,” many hybrid populations are found in areas disturbed by ­human activity, indicating a breakdown in ecological isolating mechanisms that normally would keep the species apart (­Sullivan, 1986a). Hybridization also may be facilitated by the scramble competition mating strategy used by some toads. The advertisement calls of hybrids in nature tend to be intermediate in characteristics between the parental species (Blair, 1956a, 1956b; Ideker, 1968), although in some cases the calls may not be dif­fer­ent enough to prevent interspecific mating (Malmos et al., 2001). Some suspected hybrids, however, have aty­pi­cal calls that are ­little more than clicks, whereas ­others have very extended trills (Ideker, 1968). In contrast to intermediacy, the calls of hybrid A. woodhousii x A. microscaphus are highly repeatable in call duration and pulse rate (65–80 pulses/sec), much more so than their

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parental species (Malmos et al., 2001). This suggests that hybrids do not call less than the parentals nor are their calls more variable. In a Texas fish hatchery, 6% of the calling males consisted of A. woodhousii x I. nebulifer hybrids (Brown, 1971a), and in Arizona, 25% of a chorus was hybrid A. woodhousii x A. microscaphus with the remainder all A. microscaphus (Malmos et al., 2001). The extent of hybridization may change through time and contact zones may be dynamic areas of gene exchange. For example, A. woodhousii hybridizes with A. microscaphus along the Beaver Dam Wash where it intersects the Virgin River in southwestern Utah and adjacent Arizona. Based on collections from 1949 to 1953, hybrid individuals possessed a mostly A. microscaphus-­like phenotype. By 2001, however, hybrids within the contact zone exhibited mostly A. woodhousii-­like phenotypes (Schwaner and ­Sullivan, 2009). ­Sullivan and Lamb (1988) indicated that A. woodhousii appeared to be replacing A. microscaphus in this area. Molecular data confirmed a “hybrid swarm” at the contact zone, and demonstrated considerable A. woodhousii introgression with populations of A. microscaphus that extended a considerable distance up Beaver Dam Wash. The toad population at the confluence of the rivers consists of the parental species, F1 hybrids, and individuals resulting from reciprocal backcrossing. Note that ­these results are in contrast to ­earlier suggestions of temporal stability in this region (­Sullivan, 1995), although this study was based solely on morphological and acoustic characters. A dif­fer­ent outcome has occurred on the Agua Fria River in Arizona, where A. woodhousii has not replaced A. microscaphus (see A. microscaphus account; ­Sullivan et al., 2015; Wooten et al., 2019). In New Mexico, contact between A. woodhousii and A. microscaphus has not led to hybridization in an area of ­limited syntopy (Ryan et al., 2017). Hybrids between this species have been produced in numerous laboratory experiments with the following species (reviewed by Blair, 1972): A. americanus (A.P. Blair, 1941a, 1946; W.F. Blair, 1963a); A. boreas (Moore, 1955; Blair, 1959); A. californicus (Moore, 1955); A. cognatus (Blair, 1959); A. fowleri (Blair, 1941a; Meacham, 1962); A. hemiophrys; A. houstonensis; A. punctatus (Moore, 1955); A. microscaphus; A. speciosus (Moore, 1955); A. terrestris (Blair, 1961a, 1963a); and Incilius nebulifer (Thornton, 1955; Blair, 1956b, 1959). However, some crosses work only with 1 parent (e.g., ♂ I. nebulifer and ♀ A. woodhousii), and even then, resulting males and the offspring of the reciprocal cross are sterile (Thornton, 1955). In other cases (e.g., with A. fowleri), hybridization produces fully fertile offspring that are capable of backcrossing with other hybrids or the parental species and still produce offspring (Blair, 1941a; Meacham, 1962; Blair, 1963a). Some crosses, however, are

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not successful, such as ­those with other members of the Incilius nebulifer group of toads (Blair, 1966). ADULT HABITAT

Wood­house’s Toad occupies a wide variety of mostly mesic habitats from the central and southwestern ­Great Plains to arid habitats in the American Southwest. Habitat types include plains grasslands, mountain grasslands, pinyon-­ juniper woodland, riparian woodland, ponderosa pine forest, spruce-­fir forest, open fields, and suburban environments (Bragg, 1950b; Aitchison and Tomko, 1974; Ballinger at al., 2010). The toad is frequently observed in agricultural regions, where deep-­water wetlands imbedded in croplands are vital to the survival of this species (Mushet et al., 2012). This species is often the most abundant toad in sandy habitats, although not restricted to them, and is partial to sandy floodplains and bottomlands with deep friable soils (Bragg and Smith, 1942, 1943; Timken and Dunlap, 1965; Hammerson, 1999). It is found in the depths of the ­Grand Canyon (Miller et al., 1982; personal observation) and occurs at elevations to 2,133 m in Montana (Black, 1970), 2,440 m in Colorado (Hammerson, 1999), and 900–2,400 m in New Mexico (Degenhardt et al., 1996). The highest elevation recorded east of the Continental Divide is 2,325 m in Colorado (Livo et al., 2012). TERRESTRIAL ECOLOGY

Activity occurs from March or April through September or October, depending upon latitude (e.g., Collins, 1993; Engeman and Engeman, 1996; Geluso and Harner, 2013). This species is largely nocturnal in its foraging habits throughout much of the warm season. Diurnal activity by adults may occur on cloudy days and in wet weather, but small toads tend to be more diurnal than large adults. Wood­house’s Toads frequently congregate ­under streetlights or other bright lights to which insects are drawn. Dispersal of recent metamorphs occurs en masse, with diurnal activity occurring even during very hot weather; some dispersal may occur at night. The young toadlets seek moist locations as they move away from a breeding pond and avoid the driest locations. As the toads grow, they become less diurnal. This species is capable of considerable movement. They have been found up to 1.9 km straight-­line distance from their original point of capture in a mark-­release study in Oklahoma (King, 1960), and Thornton (1960) noted a movement of 643 m from the site where a toad was originally marked. Activity occurs over a range of temperatures, with lit­er­a­ture reports of 15–33.7°C (Scott and Carpenter, 1956; Fitch, 1956a; Stebbins, 1962; Brattstrom, 1963; Hammerson, 1999). However, Hammerson (1999) noted activity as low as 8°C. Although this species is often found in very cold

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environments, it is not tolerant of freezing temperatures and possesses no cryoprotectants (Swanson et al., 1996). They are sometimes found in mammal burrows, such as ­those of prairie dogs (Cynomys ludovicianus) (Taylor, 1929; Kretzer and Cully, 2001; Lomolino and Smith, 2004), or they dig shallow burrows where they can escape midday heat. Wood­house’s Toads are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1971, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). As such, Wood­house’s Toads likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Anaxyrus woodhousii is a spring to summer breeder, with most breeding occurring from March to May in Texas and Oklahoma (Force, 1930; Bragg, 1940a; Thornton, 1960; Meacham, 1962; Car­ter et al., 2018) and April–­August in Montana (Black, 1970). Even in Texas, however, calls may be heard as late as July, and Bragg (1940a) reported chorusing in Oklahoma in early August. The advertisement call is a loud burst, not a melodic trill as in related members of the Americanus group of toads. Calls are short, ranging from 0.9 to 2.6 sec, and cover a wide range of frequencies (dominant frequency 1500–1874 cps) (Blair, 1956b). Bragg (1940a) also described a call he termed a “whew,” that is, a high-­pitched, long-­duration call with a rising inflection. Calling may be initiated by rainfall, especially ­after long dry periods, but rainfall is not necessary for calling or breeding to occur (Thornton, 1960). Calling occurs at temperatures >14.5°C (Meacham, 1962), and Thornton (1960) and ­Sullivan (1982a) noted optimal temperatures for calling of 17–30°C. Most chorusing occurs in the early eve­ning. Males call from shallow w ­ ater near the shoreline, although they may call from farther from shore if the ­water is not deep. Calling rarely occurs on land. In some areas, chorusing males tend to be clumped in groups around a breeding site (­Sullivan, 1982a), and males in ­these large choruses call more often than males in small choruses (Woodward, 1984). ­Sullivan (1985a) characterized this as a lek mating system, a system which in this case is not ­shaped by female preference for large choruses (­Sullivan, 1986b). In other areas, males call from dispersed locations and do not tend to clump near one another (Bragg, 1940c). Calling sites are defended, and males act aggressively ­toward intruders (­Sullivan, 1982a). At an intruder’s approach, ­there may be a series of back-­and-­ forth calls between the rivals, followed by amplexus of the intruder by the resident male and extended wrestling. Calling

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males invariably win such matches. While they are calling, males are very alert and ­will dive into the ­water if approached. It is not unusual to hear only a few males calling at once, but choruses in the hundreds have been reported. Males arrive at breeding ponds ­earlier in the eve­ning than females. They are non-­specific in their choice of mates, and ­will amplexus any female that moves within their vicinity. Not surprisingly, interspecific amplexus is not uncommon. For example, Brown (1971a) noted that as many as 8.8% of amplexed pairs at a Texas fish hatchery consisted of mixed A. woodhousii x I. nebulifer individuals (also see Thornton, 1960), and Bragg (1939) observed amplexus between A. cognatus and A. woodhousii. Males have a characteristic release call that is uttered in conjunction with body vibrations when the toad is amplexed or handled (84–104 vibrations/sec; Blair, 1947b). The pulse rate during the release call is positively correlated with temperature, but the duration of the release vibrations and the frequency of the release call are inversely proportional to temperature (Brown and Littlejohn, 1972). A male can continue the release calls and vibrations for a considerable amount of time. Small females may utter a short “peep” when surprised, but the call is not repeated (Bragg, 1940a). Despite the extended breeding season—­which covers several weeks—­individual males and females may spend only a few days at the breeding pond. For example, males spent from 1.9 to 2.4 nights at a breeding site in New Mexico, whereas females spent from 1.0 to 1.4 nights at the pond (Woodward, 1984); chorus tenure was positively associated with male mating success. Similar residency was observed in Arizona (­Sullivan, 1982a), but the length of stay at a chorus was not an impor­tant determinant of mating success (­Sullivan, 1985a). Chorusing does not occur ­every night at all sites; ­Sullivan (1982a) noted that choruses formed 6–36 nights (mean 23) at 5 breeding choruses in Arizona. In 2 other populations, choruses lasted 23–45 days (­Sullivan, 1986b). ­Sullivan ­later (1989) noted a mean chorus duration of 19.8 days, with an average chorus size of only 5–6 males at another desert location. Some males and even females move among nearby choruses rather than remain at a single site throughout the breeding season. Males may revisit a chorus several times during the breeding season (Woodward, 1982b). Breeding population size varies annually. For example, Thornton (1960) marked 9–87 toads (6–64 males, 3–23 females) over a 3 yr period at a site in Texas. Although larger males may have greater reproductive success than small males (Woodward, 1982a, but see ­Sullivan, 1983), the sequence of arrival is not dependent on the size of the males (i.e., large males do not arrive ­earlier in the season than small males), and the operational sex ratio (1 male per 0.26–0.34 females) remains about the same at a

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breeding site throughout the season (Woodward, 1984). In contrast, ­Sullivan (1987) found that larger males tended to arrive ­earlier in the season than smaller males. The size of the male is not associated with chorus tenure; large males do not spend more time in a chorus than small males (Woodward, 1982b). Male call rates are positively correlated with mating success (­Sullivan, 1982a, 1983, 1987) and call rate is not ­limited by aerobic capacity (­Sullivan and Walsberg, 1985). This suggests that the male’s call rate is not an accurate indicator of his phenotypic vigor. Males lower their call rates and avoid acoustic overlap with all stimuli centered at 1.4 kHz (­Sullivan, 1985b; ­Sullivan and Meek, 1986). This reduces the extent of overlap with nearby calling males, and could lead to new chorus formation as the density of calling males increases. In this regard, male call rates are inversely proportional with chorus density (­Sullivan, 1985b), but the intensity of sexual se­lection increases with chorus size (­Sullivan, 1986b). Calling male size may or may not be correlated with mating success. In some years and at least some locations, the correlation is positive, whereas in other years it is not (­Sullivan, 1983, 1987). In addition, ­there is no evidence of size-­assortative mating (i.e., large females mating with large males) (Woodward, 1982a; ­Sullivan, 1983). In acoustic choice tests, females do not prefer the calls of large males over ­those of small males. Most males do not successfully breed within a season, whereas nearly all females do. Woodward (1984) calculated a male’s chance of finding a receptive female each night at 14%, whereas the female nightly success rate was 71–95% and 100% through the season. In Woodward’s (1984) study, the number of calling males varied from 1 to 120 per night. Males amplex 0–3 females during a breeding season (­Sullivan, 1982a; Woodward, 1982b), and mating success can vary from one location to another (­Sullivan, 1986b). Relatively few males (5–10%) and females (5–15%) captured one year at a pond return to the same pond to breed in subsequent years (­Sullivan, 1987). ­Sullivan (1987) attributed the low recapture rate to high population turnover rather than toads breeding at other sites.

et al., 2008). Both temporary and permanent sites are used (Woodward, 1987a). Wood­house’s Toad readily breeds in man-­made habitats, such as constructed ponds or lakes, stock tanks, irrigation ditches, borrow pits, garden ponds, and gravel pits, and ­will also breed in flooded fields. This species appears to prefer muddy or silty ­water but does not breed in buffalo wallows. REPRODUCTION

Breeding occurs over an extended period during the warm season, even if most breeding occurs in late spring to early summer. Based on histology, Goldberg (2017b) noted that males are capable of breeding from March to July, and females from March to October, in the southwestern United States. The smallest mature male was 68 mm SUL, whereas the smallest mature female was 71 mm SUL. Most breeding occurs in May and June in Nebraska (in Ballinger et al., 2010) and from April to June in Colorado (Hammerson, 1999). In Oklahoma, breeding extends from March to September (Bragg, 1950a). Despite the extended breeding season, successful breeding actually occurs on only a few nights (Woodward, 1982b). Rainfall is not necessary to stimulate breeding activity, although reproduction frequently occurs ­after periods of rain. The female chooses the oviposition site away from the main chorus of calling males. Eggs are deposited in 2 long strings, 1 from each ovary, along the substrate and among vegetation near the shoreline. Although breeding may occur throughout a pond, eggs are often oviposited by dif­fer­ent females in close proximity to one another. Clutch sizes range as high as 25,644 (Smith, 1934), with Woodward (1987a) giving a mean clutch size of 10,469 in New Mexico, Collins (1993) providing a mean clutch size of 8,500 for 2 Kansas

BREEDING SITES

Breeding sites include a wide variety of habitats, including swamps, small ponds, river backwaters and sloughs, flowing desert streams, and in shallow ­waters in the littoral zone of small lakes (Bragg, 1941b; Bragg and Smith, 1942). They may use pools in even major river channels, such as the Platte River in Nebraska, when ­water recedes during low-­flow periods (Geluso and Harner, 2013). They may also use backwaters left ­after spring floods or pools maintained by elevated groundwater levels adjacent to rivers (Bateman

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Egg strings of Anaxyrus woodhousii. Photo: Dana Drake

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clutches, and Krupa (1995) recording a clutch of 28,500 in Oklahoma. The body weight of the female is positively correlated with clutch weight, egg weight, and the number of eggs per clutch (Woodward, 1987a). Hatching occurs ca. 3 days following oviposition (Youngstrom and Smith, 1936). The ovarian cycle is described by Clark and Bragg (1950). LARVAL ECOLOGY

The larvae become ­free swimming when they attain a length of 6.5–7.0 mm ­after 5–6 days (Youngstrom and Smith, 1936). Developmental rate varies with temperature, food resources, and conspecific density. The length of the larval period is a minimum of 34 days and can take as many as 70 days, although Hammerson (1999) gives the larval period as 4–7 weeks. The front limbs appear ­after the maximum size has been attained. Metamorphosis begins when the tadpole is about 30 mm TL, and recent metamorphs average 10–15 mm SUL (Bragg, 1940a; Degenhardt et al., 1996; Hammerson, 1999). In South Dakota, Malaret (1978) mea­sured 3 recent metamorphs at 15, 15.9, and 17.9 mm SUL, but ­these likely had already grown several mm.

Larval Anaxyrus woodhousii. Photo: John Cossel DIET

Larvae eat algae and organic detritus and may consume animal ­matter. Postmetamorphic A. woodhousii eat a wide variety of prey, nearly all insects (Tanner, 1931; Smith, 1934; Smith and Bragg, 1949; Stebbins, 1962; Flowers, 1994; Flowers and Graves, 1995; Hammerson, 1999). Prey composition may change from one year to the next depending on availability. Specific items include ants, many types of beetles, lepidopterans, cicadas, crickets, grasshoppers, weevils, cut-­worms, spiders, phalangids, sowbugs, true bugs, millipedes, and centipedes. Juveniles eat many types of insects, particularly beetles and mites (Flowers, 1994). Gehlbach and Collette (1959) suggested that Wood­house’s

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Toad might feed underwater or at the ­water’s surface since prey included dytiscid, hydrophilid, and stratiomyid larvae. A large female was reported to eat an unidentified hatchling turtle in captivity (King, 1960), and another a neonate Crotalus viridis (Prairie Rattlesnake) (Ray et al., 2017). Males may have fewer items in the stomach per individual than females, and juveniles eat much the same prey as adults, except that small ants, collembolans, and mites make up a much greater proportion of the diet (Flowers and Graves, 1995). Of course, small invertebrates are preferred. Regional differences may occur in the diets of this species. For example, certain ground beetles (Agonum sp.) make up a high percentage of the diet of A. woodhousii from eastern Oklahoma, but only a tiny percentage from western populations. Just the reverse is seen in the prevalence of small ants in the diet (Smith and Bragg, 1949). Such differences likely reflect differences in prey availability. PREDATION AND DEFENSE

Wood­house’s Toads are cryptic and possess noxious skin secretions, especially on the parotoids and dorsal warts. ­These secretions are effective at deterring predators, as evident by the unsuccessful predation attempt on a Wood­ house’s Toad by an American Bullfrog (Brown, 1974). When approached, adults may crouch down against the substrate and remain motionless for some time before attempting escape (Bragg, 1945a). Predators of postmetamorphs include frogs (Lithobates catesbeianus), snakes (Heterodon nasicus, Nerodia sipedon, Pantherophis sp., Pituophis catenifer, Thamnophis cyrtopsis, T. elegans, T. marcianus, T. sirtalis), birds (hawks), and mammals (skunks, raccoons) (Bragg, 1940a; Gehlbach and Collette, 1959; Woodward and Mitchell, 1990; in Hammerson, 1999; in Johnson, 2000; Drost, 2020; Ford, 2020). Woodward and Mitchell (1990) noted that males ­were more likely to be eaten than females, and that mammalian predators tended to disembowel toads and leave the skin with the noxious granular glands intact. Larvae are eaten by a wide variety of aquatic invertebrates and by the larvae of spadefoots (Spea, Scaphiopus). Still, they are unpalatable to some species, although the effectiveness of larval unpalatability is variable (Adams et al., 2011). Schiwitz et al. (2020) found no effect on larval activity in the presence of predator chemical cues or conspecific alarm cues. POPULATION BIOLOGY

Individual growth is rapid in A. woodhousii, with Oklahoma toads reaching 43–65 mm SUL (mean 53.4 mm) by October prior to their first winter dormancy (Bragg, 1940a). The estimated growth rate in Bragg’s (1940a) study was 0.3 mm/ day. In general, toadlets double their SUL within the first

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Adult Anaxyrus woodhousii. Photo: C.K. Dodd, Jr.

month ­after metamorphosis. First reproduction likely occurs the first (males) or second (females) year ­after metamorphosis (Thornton, 1960). As with many frogs, the sex ratio at a breeding pond is usually male biased. Bragg (1940a), however, suggests that the overall sex ratio actually may be female biased based on captures at foraging locations. The fact that some females retain eggs late in the breeding season and that female sex ratios at breeding ponds are very dif­fer­ent from sex ratios in foraging areas suggests that some females may skip a breeding season even when environmental conditions are favorable (Bragg, 1940a; Bragg and Smith, 1943). Longevity in the wild is unknown, but Engeman and Engeman (1996) observed a toad trapped in a win­dow well in Denver living outdoors for 19 yrs. COMMUNITY ECOLOGY

This species is allopatric or comes into contact with a number of other closely related toads within the A. americanus complex. As noted above, hybridization may occur along contact zones between species, but introgression may not occur as extensively as hybridization experiments might suggest. For one ­thing, females are able to discriminate advertisement calls. ­These calls are often dif­fer­ent among species and serve as a pre-­mating isolating mechanism. Advertisement calls also help females discriminate conspecifics from hybrids. For example, females respond positively to the calls of their own males but not to the calls of hybrids (Awbrey, 1965). Closely related toads also often have dif­fer­ent breeding phenology or habitat preferences that may minimize hybridization potential (Blair, 1942). Differences in habitat preference can separate the species despite very close proximity, and mixed breeding assemblages generally do not occur (Bragg,

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1940a; Bragg and Smith, 1942; but see Collins, 1993). In Texas, for example, A. woodhousii contacts A. fowleri along the western margin of the eastern deciduous forest, and hybridization occurs along the 16–30 km wide contact zone (Meacham, 1962). Fowler’s Toad prefers sandy soils and mesic forest habitats, whereas Wood­house’s Toad prefers blackland prairie soils in more open habitats, although it is more versatile in its choice of habitats. Breeding site differences also are evident, with A. fowleri preferring creeks and A. woodhousii preferring prairie ponds (Blair, 1941a). U ­ nder pre-­ European colonization conditions, habitats presented a dendritic pattern along the contact zone, or they changed abruptly at the prairie-­forest interface. The demarcation between species was also abrupt. As ­humans have modified the habitats in this region, the species are more likely to come into contact, forming broad rather than narrow contact zones. Since A. fowleri tends to outcompete A. woodhousii, the nature of the interaction between species along the contact zone changes with the nature of the habitat, and the extent of introgression varies from one location to the next. ­There also may be differences in diet between certain toads in areas where they come into contact. For example, A. cognatus eats fewer small ants, harvester ants, sow bugs, and spiders than A. woodhousii, but more lepidopteran larvae, weevils, and centipedes (Smith and Bragg, 1949). ­Whether such dietary differences reflect habitat differences, preferences, or availability is unknown. In general, A. cognatus tends to replace A. woodhousii in the mixed-­grass prairies (Bragg, 1940a, 1940b), so a combination of ­factors may influence perceived dietary differences. In some anurans, the presence of a large number of heterospecific larvae may inhibit their growth. In A. woodhousii, however, the density of heterospecifics does not appear to inhibit growth, and A. woodhousii larvae do not contain the growth-­inhibiting cell observed in many other frog species (Licht, 1967). In contrast, the presence of large numbers of A. woodhousii larvae may inhibit the growth of the larvae of some species (A. houstonensis, G. olivacea, Lithobates sp.) but not ­others (A. debilis, A. speciosus, D. chrysoscelis, S. couchii). Density is inversely proportional to growth among conspecific A. woodhousii. DISEASES, PARASITES, AND MALFORMATIONS

The amphibian pathogen Batrachochytrium dendrobatidis (Bd) was not found at sites sampled in Colorado, New Mexico, and Oregon (Green and Muths, 20005; Adams et al., 2010; Suriyamongkol et al., 2019), but has been reported from Kansas (McTaggart et al., 2014), Nebraska (Harner et al., 2013), Oklahoma (Watters et al., 2016, 2018, 2019; Marhanka et al., 2017), South Dakota (Brown and Kerby, 2013), Texas (Todd et al., 2019), and Wyoming (Erdmann

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et al., 2018). Ranavirus has been detected in individuals from Nebraska (Davis and Kerby, 2016) and Oklahoma (Smith et al., 2019). Endoparasites include the trematodes Glypthelmins quieta, Gorgoderina amplicava, G. attenuata, Haematoloechus sp., Haematoloechus coloradensis, H. complexus, H. medioplexus, H. parviplexus, and Megalodiscus temperatus, the cestodes Cylindrotaenia americana and Distoichometra bufonis, the cestode Distoichometra bufonis and an unidentified metacestode, and the nematodes Aplectana incerta, A. itzocanensis, Cosmocercoides variabilis, Gyrinicola batrachiensis, Ozwaldocruzia pipiens, Physaloptera sp., Rhabdias americanus, and Rhabdias sp. (Trowbridge and Hefley, 1934; Parry and Grundman, 1965; Brooks, 1976; McAllister et al., 1989; Goldberg et al., 1996b; Green and Muths, 2005; Rhoden and Bolek, 2011, 2015). Waitz (1961) found no endoparasites in 2 animals. Green and Muths (2005) mentioned additional unidentified trematodes, nematodes, and cestodes. Rhoden and Bolek (2015) also mention the presence of Plagiorchid metacercariae and Echinostomatid metacercariae in this species. The myxozoan parasite Myxidium serotinum has been reported from A. woodhousii, as have the protozoans Opalina sp., Zelleriella sp., Nyctotherus cordiformis, Karatomorpha swazyi, Trichomonas sp., and Hexamita intestinalis (Parry and Grundman, 1965; McAllister et al., 1989). Ectoparasites include the leech Desserobdella picta in larvae (Bolek and Janovy, 2005) and the mite Hannemania dunni in adults (Loomis, 1956). A toad with an extra leg was reported from Oklahoma (King, 1960), and toads with missing eyes ­were observed in Colorado (in Hammerson, 1999).

and they actually grow larger, even at high densities. In essence, the pesticide reduces or eliminates competition for food resources from zooplankton, allowing greater tadpole growth and survival. The insecticide itself appeared to have no immediate adverse effects on the larvae. Boone and Semlitsch (2002) urged caution when interpreting the results, noting that effects on a community from an unnatural change in community structure could be subtle and occur long ­after the larval period. The insecticide malathion ­causes morbidity and mortality in adult A. woodhousii, even at sublethal doses (0.011, 0.0011 mg/g toad mass), when administered in conjunction with amphibian red-­leg (Aeromonas hydrophila) disease; no effects ­were seen in toads challenged by the bacterium alone (Taylor, 1998; Taylor et al., 1999a). The insecticide appears to cause toads to be more susceptible to naturally occurring disease pathogens than they would be when the pesticide was not pre­sent. Malathion at doses >0.110 mg/g toad mass is directly lethal when applied to the ventral skin. Malathion ­causes a decrease in brain cholinesterase activity levels. STATUS AND CONSERVATION

SUSCEPTIBILITY TO POTENTIAL STRESSORS

Threats to Wood­house’s Toad include habitat destruction, alteration, and fragmentation, mortality from roads and other transportation corridors, and pesticides. Declines ­were noted in urban areas as early as 1944 (Bragg, 1952). Bragg (1940a) noted that recent metamorphs ­were used as fish bait in Oklahoma and that thousands of A. woodhousii ­were killed on roads ­every year. Roadkill of im­mense proportions has occurred at least for 100 years, especially considering the extensive range of this species. The species generally avoids pine plantations, preferring instead the mixed-­hardwood riparian areas that bisect them (Rudolph and Dickson,

Chemicals. U ­ nder normal conditions (no insecticide), tadpole body mass is inversely proportional to density. The addition of the carbamate insecticide carbaryl (3.5 or 7.0 mg/L) actually increased the survivorship of A. woodhousii larvae in 1 set of experimental treatments, proportionally more so at high tadpole densities than at low densities (Boone and Semlitsch, 2002; Boone et al., 2004b). Body mass also was greater at metamorphosis in carbaryl-­ treated ponds than in ­those not treated with the insecticide. In contrast, an ­earlier study had suggested that carbaryl decreased survivorship of tadpoles to metamorphosis, an effect that was more pronounced at high tadpole densities than at low densities (Boone and Semlitsch, 2001). Even at low densities, survivorship greatly decreased at 7.0 mg/L carbaryl, although survivorship at 3.5 mg/L actually increased, as in the ­later study. Increased density also reduced mass at metamorphosis. As zooplankton are killed, ­there is a greater amount of algae available to the tadpoles

Breeding habitat of Anaxyrus woodhousii. Photo: David Pilliod

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1990). When pre­sent in clearcuts or intensely managed silvicultural areas, numbers are low (Fox et al., 2004). Nicoletto (2013) noted no adverse effects from a hurricane passing over his inland study site in Texas, despite considerable damage to the canopy cover. Wood­house’s Toads did not appear to be seriously impacted in Missouri River floodplain wetlands that had flooded catastrophically in 2011 (Grant et al., 2015, 2018). Adults occupied bare ground before the catastrophic flood and colonized areas with shallow slopes ­after the flood (Grant et al., 2015). Metamorphs, however, did not survive as well in such areas with decreased emergent vegetation, where such habitats offered ­little anchorage for egg strings (Grant et al., 2018). In contrast, floods along the Rio Grande River resulted in large numbers of toads breeding in an area with ­little previous activity over a long period of time (Bateman et al., 2008). Thus, ­there can be short-­term differences in response to disturbance events that vary by life

stage, differences that could not be detected by call surveys or tadpole sampling alone. ­There does not appear to be any indication of declines throughout substantial portions of this species’ range (Christiansen, 1981; Busby and Parmelee, 1996; Hammerson, 1999; Hossack et al., 2005). However, surveys in Big Bend National Park from 1998 to 2004 failed to detect A. woodhousii despite the availability of historical rec­ords (Dayton et al., 2007). The species has been monitored in a variety of ways. Corn et al. (2000) noted that automated recording devices and observer-­based call surveys did a poor job at estimating the abundance of this species, perhaps ­because the devices ­were located at some distance from the chorus being monitored. A mark-­recapture study yielded a population estimate of 213 toads, considerably larger than what had been estimated based on the call surveys alone. This species ­will occasionally colonize man-­made ponds during habitat restoration efforts (Briggler, 1998).

Incilius alvarius (Girard, 1859) Sonoran Desert Toad

whereas swimming males had a mean of 134.1 mm SUL. Using museum specimens, Goldberg (2018a) reported males 108–143 mm SUL (mean 122.9 mm) and females 88– 160 mm SUL (mean 132.6 mm). Brennan and Holycross (2006) noted a maximum size of 191 mm SUL. Larvae. The body of the tadpole is somewhat flattened, and is lightly pigmented and brassy in life. ­There is no pigmentation on the center of the belly. The tail musculature is evenly pigmented with large spots or blotches, but the tail fins are largely clear, although ­there may be a few small spots on the dorsal fin. Tail tips are rounded. Jaws are coarsely serrate. TL is  25 mm) of the summer monsoon, although the breeding season may be extended. For example, the presence of tadpoles in October suggests that breeding can occur through September (Degenhardt et al., 1996). Chorusing may be delayed till the second or third night ­after excessive rainfall events. Eggs are oviposited in long (to >1 m in length) jelly-­coated strings in temporary pools or shallow streams in ­water 30–45 cm in depth. Clutch size is 7,500– 8,000 eggs (Livezey and Wright, 1947). LARVAL ECOL­O GY

The larval period is short (17°C, with calls heard at eve­ning temperatures as high as 35°C (Thornton, 1960; Blair, 1961b). Wiest (1982) noted calling at 18.5–28.3°C over a 44 day period in Texas; in northeast Texas, calling occurs from March to July (Car­ter et al., 2018). Males may return to breed at a pond several times during a season when conditions permit. In addition, some males ­will call early in the season whereas ­others call ­later in the season. Thus, not all males are pre­sent at a breeding pool even ­under favorable conditions. The call of I. nebulifer is a sharp trill of moderate length. Trill rates vary from 30 to 43/sec at a dominant frequency of 1,250–1,700 cps; the trill lasts 1.9–5 sec (Blair, 1956b) and is repeated at 1–4 sec intervals. The sound has been likened to a wooden rattle. It is similar to the call of A. americanus but is more guttural and less musical. BREEDING SITES

Gulf Coast Toads breed in intermittent streams, temporary ponds, shallow pools, and virtually any available ­water body.

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Tadpole of Incilius nebulifer. Photo: Ronn Altig

Breeding also occurs in artificial pools, gravel pits, irrigation ditches, and stock tanks (Blair, 1960a; Hubbs and Martin, 1967). Breeding has been recorded in brackish ­water (Burger et al., 1949; in Neill, 1958). REPRODUCTION

The peak breeding season in Texas is April–­September, with toads first appearing at breeding ponds from February to April and the last reproduction occurring in September; variation in the timing of breeding is dependent on temperature, with toads in the south active ­earlier and ­later in the season than toads in the north (Blair, 1960a; Thornton, 1960; Salinas, 2009). Based on histology, Goldberg (2017a) noted that males are capable of breeding from January through autumn, and females from February to August, in Texas. The smallest mature male was 50 mm SUL, whereas the smallest mature female was 60 mm SUL. In Louisiana, calls are heard from April to September, although ­there are reports of breeding in March (Dundee and Rossman, 1989). In the spring, breeding is initiated by rainfall that fills the breeding ponds, and reproduction is sporadic and dependent on precipitation throughout the season. For example, Blair (1960a, 1961b) recorded only 1 rain event over a 2 day period that resulted in breeding in 1956, but in the wet year of 1957, the season extended 125 days. Although breeding may occur as late as August, the onset of dry weather may preclude successful metamorphosis. Females are capable of ovipositing more than 1 clutch per breeding season (Blair, 1960a). Spent females also return to breeding sites ­after periods of rainfall. Eggs are deposited in shallow ­water in long gelatinous strings about a meter in length. ­These strings can ­either rest on the substrate or be wound around vegetation or other submerged debris. Greuter (2004) reported exact counts of 1,887–5,614 eggs. Hatching occurs in 1.5–3 days. Embryos are capable of developing at 18–35°C, with an optimal temperature of 20–30°C (Volpe, 1957c; Hubbs et al., 1963; Ballinger and McKinney, 1966). Abnormalities increase at temperatures >31.5°C. Rates of development are similar among populations from vari­ous parts of its range in Texas and Louisiana.

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LARVAL ECOL­O GY

The larval period is 20–30 days u ­ nder laboratory conditions, with tadpoles reaching 25 mm prior to metamorphosis (Volpe, 1957c). Salinas (2009) mentions a larval period of 2–10 weeks. No other information is available on larval ecol­ogy. DIET

­ here are no published studies on the diet of this species. T Presumably, the toad eats a wide variety of invertebrates. PREDATION AND DEFENSE

Gulf Coast Toads have noxious and poisonous skin secretions that are effective in deterring predators. For example, Brown (1974) reported an unsuccessful predation attempt by an American Bullfrog on an adult I. nebulifer, and Salinas (2009) noted an opossum carry­ing a toad. Larvae are unpalatable to some species, but unpalatability is variable in effectiveness (Adams et al., 2011). The eggs are toxic to some species when injected (Licht, 1968), but not when eaten by mosquitofish (Gambusia) (Grubb, 1972). Garter Snakes (Thamnophis) may be immune to the toxins in eggs. Western Ribbonsnakes (T. proximus) have been observed eating young toads (Wright and Wright, 1949). Larvae are able to detect chemical cues from potential predators, but the response varies by predator and aggregation status of the tadpoles. In addition, tadpoles can detect cues from conspecifics that have been attacked by predators. Solo and small groups of tadpoles reduce their activity when exposed to predated conspecific cues, whereas only solo tadpoles reduce activity in response to a nearby predator’s cues. In effect, ­whether a tadpole is solo or a member of a group affects resource acquisition and predation response, and thus indirectly affects the life history of this species (Preston and Forstner, 2015a). In contrast to ­these results, Schiwitz et al. (2020) found no effect on larval activity in the presence of predator chemical cues or conspecific alarm cues.

females) over a 3 yr period at a site in Texas. Likewise, sex ratios vary, although they are male biased at breeding sites. For example, Blair (1960a) recorded sex ratios of 1.94 males per female over 3 yrs, but 4:1 one yr and 13.3:1 another yr over a 5 yr study. In south Texas, Salinas (2009) recorded a male skewed sex ratio of 1.96:1 over a 1 yr survey. Skewed ratios result from dif­fer­ent amounts of time spent at breeding ponds—­with females staying only ­until egg deposition—­and dif­fer­ent climatic conditions during the breeding seasons. Thus, females may be missed during sampling while males are more likely to be encountered by researchers. In addition, a drought-­shortened breeding season may result in only a portion of the adults visiting the breeding site, and some toads may skip a breeding season. In Blair’s (1960a) study, the entire breeding population was estimated at 97–208 adults but was still suspected of being male biased, whereas Salinas (2009) estimated her population at 2,935 individuals. Survivorship is likely greater in females based on mark-­recapture data. Gulf Coast Toads are relatively long lived, with males surviving as long as 8 yrs and females 5 yrs (Blair, 1960a).

POPULATION BIOLOGY

COMMUNITY ECOL­O GY

Growth occurs rapidly in young toads (0.55 mm/day), slows during the winter (0.11 mm/day), then accelerates in spring (0.2 mm/day) ­until the toads reach maturity. ­After maturity, growth slows to 0.04 mm/day. Males attain breeding size the summer following metamorphosis at about 10 months of age at 61–78 mm SUL (Blair, 1953; Thornton, 1960). Age of first reproduction in females is not known. Greuter (2004) reported a survivorship of 10 ppt are lethal to larvae, but mass at 1–5 ppt does not differ from controls at 0.44 ppt (Hua and Pierce, 2013). Tadpoles exposed to sublethal levels of salinity do not have increased tolerance upon ­later exposure to higher salinity concentrations. Instead, ­these tadpoles died at a faster rate. Increased levels of salinity in habitats occupied by this species pose a potential threat. Plant extracts. In mesocosm experiments, Chinese tallow (Triadica sebifera) leaf litter had no effect on survival or development of larval I. nebulifer (Cotten et al., 2012).

DISEASES, PARASITES, AND MALFORMATIONS

The myxozoan parasite Myxidium serotinum has been reported from I. nebulifer as have the protozoan Opalina sp., the apicomplexan Eimeria, and an Adelina-­like coccidian, the cestode Mesocestoides sp., the trematode Mesocoelium cf. monodi, and the nematodes Aplectana incerta and Cosmocercoides variabilis (McAllister et al., 1989; McAllister et al., 2017). The virulent fungal pathogen Batrachochytrium dendrobatidis also has been found in this species at an 83% rate of infection in south central Texas in 2006 (Gaertner et al., 2010). An adult male with an extra forelimb was reported from Texas (Garcia and Schalk, 2020). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Salinity. Embryos had 90–95% hatching success at salinities of ≤4 ppt, 74% at 6 ppt, and no hatching >8–10 ppt. (Alexander et al., 2012). Salinity also had mea­sur­able effects on larval time to metamorphosis (longer), mass (lower), and metamorph hind limb length (shorter) at salinities of ≤2 ppt than at 4–6 ppt. Thus, even relatively low levels of salinity

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STATUS AND CONSERVATION

Although this species likely has lost populations due to habitat destruction and alteration, it appears to be common throughout its range. Still, ­there are no long-­term data showing status and trends. In urban situations, Gulf Coast Toads may be unable to breed in managed ponds, especially when ­water levels are drawn down or chlorinated during the breeding season thus killing eggs and tadpoles (Salinas, 2009). In addition, toads may be killed on roads and walkways, and by lawn maintenance equipment. A single study indicated no appreciable effects from a wildfire (Brown et al., 2014). Likewise, Nicoletto (2013) noted no adverse effects from a hurricane passing over his study site in Texas. Recent evidence has suggested that ­human disruption of breeding sites has facilitated hybridization, and even may be contributing to the decline of other toad species as I. nebulifer extends its range (Vogel, 2007). Acute artificial lighting alters calling be­hav­ior by decreasing the number of males calling in a chorus and decreasing the intensity of calling (Hall, 2016).

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176  Bufonidae

Rhinella marina (Linnaeus, 1758) Cane Toad Poloka (Hawaiian) ETYMOLOGY

marina: from the Latin marinus meaning ‘pertaining to the sea.’ Linnaeus described the Cane Toad from a drawing in Seba (1734), who mistakenly believed the species lived on land and in the sea.

it was in the 1980s. Krakauer (1968) reported a mean adult size of 108 mm SUL. In Taylor and Wright’s (1932) series from south Texas, toads ­were 66–168 mm SUL. Larvae. Tadpoles are small, round, dark brown to black, and usually 10–25 mm TL. Throats are pigmented. Viewed laterally, the body appears flattened. ­There is a distinctive pale cream stripe on the lower edge of the tail musculature, but ­there is no evidence of ­saddles on the dorsal part of the tail musculature. Tail fins are a uniform translucent gray. Eyes are dorsal in position. Eggs. The eggs are small (1.7–2 mm in dia­meter) and black. They are oviposited in very long gelatinous strings.

NOMENCLATURE

Pramuk et al. (2007): Chaunus moved to Rhinella Frost et al. (2006a): Chaunus marinus Fouquette and Dubois (2014): Bufo (Rhinella) marinus Synonyms: Bufo gigas, Bufo horridus, Bufo horribilis. Complete synonymy in Amphibian Species of the World 6.0, an Online Reference. Information in this account pertains primarily to Florida, Hawai’i, and Texas. ­There is an extensive lit­er­a­ture on the biology of this species, particularly on its impacts to native fauna in Australia. Readers should consult Zug and Zug (1979), Lever (2001), and Shine (2018) for more extensive reviews of Cane Toad biology.

DISTRIBUTION

Rhinella marina occurs naturally from south Texas throughout Central Amer­i­ca into the Amazon Basin of northern South Amer­i­ca. Cane Toads are native in the United States only in a few Texas counties bordering the lower Rio Grande, although Brown (1950) noted deliberate releases of R. marina from Monterey, México, into Hogg County, Texas. Introductions throughout tropical areas of the world ­were

IDENTIFICATION

Adults. This is a very large, heavy-­bodied toad, and full-­ grown adults cannot be confused with any other species. The ground color usually is a uniform light brown in males, although some individuals may appear yellowish or dark brown. Females and juveniles have cream-­colored and dark patches dorsally in a somewhat interlaced reticulated pattern. Ovate to triangular parotoids are large and distinct, and large warts are pre­sent dorsally and on the tops of the legs. Male warts tend to be spiny whereas female warts are smooth. The top of the head is smooth with ridges pre­sent from the nose to the back of the head. The maximum body width is nearly three-­fourths of the body length. Venters are mottled in adults, but the venters of newly metamorphosed R. marina are black in the Tampa Bay region. Males are smaller than females. Adults are usually 100–180 mm SUL, but the largest Cane Toad on rec­ord is 230 mm SUL from Suriname. Rossi (1981) indicated the largest R. marina from Florida was 168 mm SUL. Meshaka et al. (2004) reported males to 150 mm SUL and females to 175 mm SUL from Highlands County, Florida. In 5 south Florida locations, males averaged 99.3–113.2 mm SUL (range 75–135 mm) and females 107.9–140.2 mm SUL (range 89–165 mm) (Meshaka et al., 2004). According to ­these authors, the mean adult body size is now smaller than

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Distribution of Rhinella marina. Populations in Florida are introduced.

Distribution of Rhinella marina in Hawaii.

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conducted ostensibly to control sugarcane beetles and cut-­worms. Although Cane Toads ate the beetles, they ­were in­effec­tive at controlling this pest ­because they are generalists that do not specialize on sugarcane beetles or cut-­worms. Despite repeated failure as pest control agents, introductions have continued, resulting in the pest control agent becoming the pest. Cane Toads ­were introduced from Puerto Rico and other areas unsuccessfully several times in Florida beginning in 1936, with intentional releases from the pet trade supplementing agriculture-­related releases. The pre­sent population was possibly the result of an accidental release from an animal dealer in 1955, but the population was supplemented by intentional releases in 1963 and 1964 (Riemer, 1958; King and Krakauer, 1966; Krakauer, 1970a; Lever, 2001; Krysko et al., 2019). ­These latter releases included toads from Suriname and Columbia. Riemer (1958) also mentions unsuccessful releases of the ­giant toads Rhinella arenarum and R. paracnemis. Cane Toads are currently reported from many scattered locations throughout the Florida peninsula from Clay and Columbia counties southward (Krysko et al., 2019). Calling was reported from Orlando in Orange County in 1979 (Rossi, 1981), but the species has expanded northward since then. Not all sightings represent breeding populations, however. Johnson and McGarrity (2010) reported an isolated population from Bay County along Florida’s northern Gulf Coast, but this population likely was extirpated by cold weather. Range expansion may continue along coastal areas, but northward expansion and establishment likely ­will be prevented by periodic cold weather. Introductions into Louisiana ­were not successful (Lever, 2001). Rhinella marina adults have been transported in horticultural shipments from Florida to Mississippi (Holbrook, 2015). Cane Toads ­were introduced into Hawai’i from Puerto Rico in March and April 1932; the site of introduction was the Manoa Valley and Waipio on the island of Oahu. The introduction was immediately successful and young toads ­were observed by August. The population exploded by the thousands. Subsequent introductions took place in 1933 to Hawai’i, Kauai, Maui, and Molokai. The history and success of introductions is reviewed by Pemberton (1933, 1934), Oliver and Shaw (1953), and Lever (2001). The species does not occur on the other Hawaiian Islands. In addition to islands in Hawai’i, Cane Toads are found on islands in the Florida Keys (Stock Island and Key West; Krakauer, 1970a; Lazell, 1989). In addition to its native range, the Cane Toad occurs in Australia, Bermuda, many Ca­rib­bean islands (including Puerto Rico), the Chagos Archipelago, Guam, Japan, the

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Philippines, Papua New Guinea, throughout the Pacific islands, and Taiwan (Zug and Zug, 1979; Easteal, 1981; Lever, 2001; Kraus, 2009). Impor­tant distributional references include: Florida (Krakauer, 1970a; Krysko et al., 2019), Hawai’i (McKeown, 1996), and Texas (Taylor and Wright, 1932; Dixon, 2000, 2013). FOSSIL REC­O RD

Miocene fossils attributed to R. marina are reported from Kansas (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

The Cane Toad is a member of the South American toad radiation, and is most closely related to R. jimi, R. poeppigii, R. cerradensis, R. veredas, and R. schneideri (Pauly et al., 2004; Pramuk et al., 2007; Maciel et al., 2010; Vallinoto et al., 2010). This group diverged from the rest of the R. marina clade during the late Miocene, about 10.5 mya. Fossil evidence suggests the species may have had a more extensive range in North Amer­i­ca during the Miocene. Laboratory crosses between R. marina and A. woodhousii or A. fowleri ­were not successful (Blair, 1959). ADULT HABITAT

In south Texas, Cane Toads occur along the Rio Grande Valley. Introduced populations of Cane Toads in Florida are found mostly in moist, open, urban, and agricultural environments as opposed to natu­ral habitats (Krakauer, 1968; Meshaka et al., 2004; Surdick, 2005). They do not occur in wet prairies, dry sandy areas, or in natu­ral habitats within the Everglades (Meshaka et al., 2000). Rossi (1981) recorded them in disturbed riparian habitats in the Tampa area, suburban yards, agricultural areas, areas dominated by exotic plants, brushlands dominated by castor bean (Ricinus communis), willows, and strangler fig (Ficus aurea) and pine flatwoods. In Hawai’i, Cane Toads are found in the lowlands throughout the islands on which they have been introduced. They occur from sea level to 300–600 m on Maui and Oahu, depending on location (in Lever, 2001). TERRESTRIAL ECOL­O GY

Rhinella marina usually does not stray far from ­water, and activity is stimulated by heavy rainfall. Most adult and subadult activity occurs at night from March to November in Florida (Krakauer, 1968; Rossi, 1981; Meshaka et al., 2004), although activity may be extended in the most southern portions of the peninsula and Keys. For example, Krakauer (1968) observed them with full stomachs being active in winter and sitting along canals. Metamorphs tend to be diurnal. In Hawai’i, activity is both diurnal and nocturnal, but most are active ­after sunset. Toads hide ­under

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178  Bufonidae

surface debris such as rocks, logs, and pine ­needles, and in canal banks. Dispersal occurs along roads and highways in Australia (Shine, 2018); toads avoid heavi­ly vegetated habitats (Brown et al., 2006). It is not known how dispersal occurs in Florida or Hawai’i, but it seems reasonable that Cane Toads use transportation corridors to facilitate movements. Rossi (1981) also mentions deliberate and inadvertent transport by ­people. Throughout much of its range, the ambient temperatures experienced by R. marina are rather stable. The species can tolerate a wide range of temperatures. For example, the CTmax for Florida adults is 40–40.8°C and the CTmin is 5–10°C, depending upon acclimation temperature (Krakauer, 1968, 1970b). ­Under experimental conditions, Cane Toads select temperatures from 21 to 28°C, with neither light nor season influencing mean temperature se­lection (Sievert, 1991). Still, the species may experience cold spells at the northern limit of its range, and this may prohibit northward expansion. Krakauer (1970b) recorded that 6 of 10 adults died at 4.2°C over a 96 hr period. Interestingly, cold weather does not seem to result in immunosuppression in this species as it does in other ectotherms (Carey et al., 1996). Cane Toads are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (probable “blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). Cane Toads likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males have what has been described as a deep, slow, booming melodic trill. Calling has been reported from December to September in Florida, depending on location (Krakauer, 1968; Rossi, 1981; Meshaka et al., 2004). Calling has been heard at ambient temperatures as low as 15°C (Meshaka et al., 2004). It is likely that some reproduction occurs year-­round in the extreme south inasmuch as gravid females are found in all months of the year. In Hawai’i, calling occurs year-­round. ­There appears to be no information on the Texas population. Amplexed male R. marina have a slow series (mean 6.9/ sec) of warning vibrations to alert a conspecific as to the sex of the grasped toad (Blair, 1947b). Male A. terrestris have been observed in amplexus with female R. marina in Florida (in Lever, 2001).

recorded in ponds, puddles, rock pits, and man-­made canals. Krakauer (1970a) and La Rivers (1948) mentioned breeding in slightly brackish ­water and lily ponds, and successful development in salinities of 10–15% dilute sea ­water was noted by Ely (1944). Chris Lechowicz (personal communication) has observed successful breeding in rainwater-­filled runnels on the beach on Sanibel Island, 12 m from the Gulf of Mexico, where salinities ­were 3–5 ppt. REPRODUCTION

Most oviposition occurs from March to September in Florida and year-­round in Hawaii. No information is available for Texas. Krakauer (1968) found Florida females with mature eggs year-­round, but suggested that cold weather might limit oviposition. Breeding may be stimulated by hurricanes that result in high amounts of rainfall over a very short period (Meshaka, 1993), but rainfall is not necessary for reproduction. Eggs are deposited in very long strings containing 5,000–32,000 eggs (Krakauer, 1968, 1970a). ­These strings are loosely attached to vegetation and aquatic debris. Hatching occurs in 1.5–4 days. LARVAL ECOL­O GY

Larvae are herbivores consuming algae and other plant detritus. Larval Cane Toads form large aggregations (Mares, 1972; photo in McKeown, 1996). They swim actively both day and night, and schools can cover a large distance. Tadpoles can tolerate rather high ­water temperatures, with the CTmax from Florida larvae being 41.6–42.5°C (Krakauer, 1970b). Larvae acclimated at 7°C can survive normally ­after 72 hrs exposure, with half being killed ­after 96 hrs. ­These results suggest that Cane Toads from northern populations can tolerate low and even freezing temperatures better than their tropical conspecifics. The duration of the

BREEDING SITES

Breeding occurs in a wide variety of habitats, from small shallow pools to open wetlands. The species has been

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Recent metamorph of Rhinella marina. Highlands County, Florida. Photo: Kevin Enge

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Rhinella marina 179

larval period is about 45–60 days in south Florida (Krakauer, 1970a; Meshaka et al., 2004) and 30 days in Hawaii (Oliver and Shaw, 1953). New metamorphs are 9.75–13 mm SUL (Meshaka, 1993; Meshaka et al., 2004). DIET

Cane Toads eat just about any invertebrate or vertebrate they can catch and stuff into their mouth; they have a prodigious appetite. They even readily consume stationary items such as pet food left out in a dish, garbage, fruits, and vegetables (Alexander, 1965). Apparently they can use olfactory as well as visual cues to orient ­toward and identify food (Rossi, 1981, 1983), and toads are reported to see prey up to a meter away (Pemberton, 1934). The forelimbs are used to stuff large prey into the mouth. Specific items in the diet of Florida toads include snails, isopods, land crabs, ants, cockroaches, lepidopterans, dragonflies, true bugs, beetles, earwigs, spiders, flies, juvenile R. marina, other frogs (Anaxyrus quercicus, A. terrestris, Dryophytes squirellus), small snakes (Diadophis punctatus, Thamnophis sauritus, Indotyphlops braminus), and a hatchling Apalone ferox (Krakauer, 1968; Rossi, 1981; Meshaka et al., 2004; Meshaka and Powell, 2010; Schuman and Bartoszek, 2019a). In Hawai’i, the diet includes grasshoppers, armyworms (Spodoptera mauritia), cut-­worms, borers, weevils, beetles (including large quantities of the ­rose beetles Adoretus sinicus and Pantomorus godmani), centipedes, millipedes, spiders, true bugs, earthworms, scorpions, wasps, moths, bees, ants, caterpillars, slugs, snails, cockroaches (including the exotic pest Pycnoscelus surinamensis), sow bugs, and geckoes (Pemberton, 1934; Illingworth, 1941; Oliver and Shaw, 1953; Lever, 2001). Rossi (1981) also found mammal hair, bird bones, and another unidentified bufonid, and observed Cane Toads scavenging dead fish and eating dog feces. Plant material is frequently ingested. In a laboratory setting, they consumed ­every vertebrate offered, including vari­ous frogs and small snakes. Although Cane Toads consume large quantities of insect pests, their effectiveness at controlling pests is questionable.

Hawk), and snakes (Drymarchon couperi, Heterodon platirhinos, Nerodia fasciata, Thamnophis sauritus, T. sirtalis) (Rossi, 1981; Meshaka, 1994; Schuman et al., 2020). Birds and some mammals learn to roll the toads onto their backs and eviscerate them, thus avoiding the poisonous secretions. In Hawai’i, mongooses (Herpestes) may occasionally eat Cane Toads (La Rivers, 1948; Oliver and Shaw, 1953), but they are not a major portion of the diet and mongooses are not an effective control agent. Tadpoles are noxious or poisonous to some predators but are eaten by Bullhead Catfish (Ameiurus natalis). ­Under laboratory conditions, fertilized eggs of R. marina are toxic to gastropods (Lymnaea sp.), goldfish (Carassius auratus), and larval frogs (Anaxyrus terrestris, Dryophytes cinereus, Osteopilus septentrionalis, Lithobates sphenocephalus, Scaphiopus holbrookii) (Punzo and Lindstrom, 2001). However, many aquatic invertebrates, including crayfish, predaceous ­water bugs, and larval Gastrophryne carolinensis eat R. marina eggs with no ill effects. POPULATION BIOLOGY

Growth is rapid, with toads reaching 60–80 mm SUL in Hawai’i within 3 months of hatching. Individuals reach sexual maturity in 1 yr in Hawai’i (Pemberton, 1934). Longevity in wild populations is unknown, but Pemberton (1949) kept a captive for 16 yrs in Hawai’i. COMMUNITY ECOL­O GY

The presence of R. marina larvae did not affect the growth, development, or survivorship of larval Anaxyrus terrestris or Dryophytes cinereus ­under experimental conditions (Smith, 2005a). Larval Osteopilus septentrionalis consume larval R. marina, and eating the toxic toad larvae had no effect on

PREDATION AND DEFENSE

­These ­giant toads likely have few major predators as adults, particularly ­because of the toxic and noxious skin secretions that are produced in copious amounts from the parotoids and other granular glands. When threatened, they inflate their body and orient it in such a manner as to pre­sent the attacker with the toxin-­containing parotoid glands, and they may even jump ­toward the attacker. However, a number of predators on R. marina have been reported from Florida, including the ­Virginia Opossum (Didelphis virginiana), birds (American Crow, Bluejay, Mockingbird, Red-­shouldered

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Adult Rhinella marina, Hillsborough County, Florida. Photo: C.K. Dodd, Jr.

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180  Bufonidae

the subsequent survival of the Cuban Treefrog larvae, although tadpoles had symptoms of intoxication for 24 hrs following ingestion (Smith, 2005b). The diets of adult A. terrestris and R. marina overlap considerably (0.86) suggesting the potential for competition between ­these species should resources become limiting (Meshaka and Powell, 2010). DISEASES, PARASITES, AND MALFORMATIONS

The fungus Basidiobolus sp. was found on Cane Toads from Tampa, Florida (Okafor et al., 1984). Bd, but not ranavirus, has been found on Cane Toads on the island of Hawai’i (Goodman et al., 2019). The exotic tick Amblyomma rotundatum may have been introduced with R. marina into south Florida and is now established ­there (Oliver et al., 1993). Other reported tick parasites include A. americana and Dermacentor sp. (Rossi, 1981). The rat lungworm, Angiostrongylus cantonensis, was found in Cane Toads examined on the island of Hawai’i (Niebuhr et al., 2020). Eggs of the hookworm parasite Aclyostoma caninum, the threadworm Strongiloides sp., the coccidian Isopora sp., and large numbers of unidentified protozoans have been reported from toad feces; ­these likely resulted from the ingestion of dog and cat feces (Rossi, 1981). In other parts of its range, R. marina is parasitized by protozoans, helminths, and ticks, and a variety of pathogens affect this species, including viruses, bacteria, and fungi (Zug and Zug, 1979; Lever, 2001). Neoplasias (cancers) also have been reported. In south Florida, increased gonadal abnormalities and the frequency of intersex Cane Toads are correlated with agricultural activities. Gonadal abnormalities in turn are associated with impaired gonadal function, especially regarding the effects of testosterone. Secondary sex charac­ ere ­either feminized (increased skin mottling) or teristics w demasculinized (reduced forearm width and nuptial pad number) in intersex toads. Male toads had hormone concentrations and secondary sex traits intermediate between intersexes and toads from non-­agricultural areas (McCoy et al., 2008). Taken together, ­these results suggest that toads from agricultural areas have reduced fitness and reproductive success.

Breeding habitat of Rhinella marina in uplands habitat. Highlands County, Florida. Photo: Kevin Enge

SUSCEPTIBILITY TO POTENTIAL STRESSORS

Chemicals. Cane Toads in Hawai’i ­were poisoned by strychnine ­after ingesting flowers of the tree Strychnos nuxvomica (Arnold, 1968). STATUS AND EFFECTS ON NATIVE FAUNA

The status of the native Texas population is not reported in the lit­er­a­ture. Throughout the world, the non-­aggressive R. marina has had detrimental effects on native faunas wherever they have been introduced (e.g., Burnett, 1997; Lever,

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Breeding habitat of Rhinella marina. Runnel beach pool on Captiva Island, Florida. Photo: Chris Lechowicz

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2001; Griffiths and McKay, 2007; Letnic et al., 2008; Shine, 2018), despite glowing support for their introduction to control pests particularly among organic farmers (Pfeiffer, 1949). ­Little is known of their effects on the fauna of Florida and Hawai’i. In Australia, where most research on invasive toads has occurred, Cane Toads have caused high rates of mortality of native fauna and have drastically altered community composition and structure. Adverse effects are largely due to the toxic nature of the toad’s skin secretion coupled with the tendency of naïve predators to attack novel prey. This may not be as much of a prob­lem in Florida where other bufonids occur and to which predators have been exposed. Effects on the endemic Hawaiian bird and insect fauna are not known. Cane Toads may have negative impacts on native frogs and toads. ­Because of their large size and appetite, they consume almost any small invertebrate or vertebrate within range. In laboratory feeding ­trials, they aggressively outcompete domestic toads by striking them into submission with their tongue (Boice and Boice, 1970). Cane Toads have lateral vision and can observe an approaching competitor in order to slap it with their tongue and discourage further approach ­toward a prey item (Robins et al., 1998). ­Whether feeding competition occurs in nature or was an artifact of captivity is not known. Like adults, larval Cane Toads are toxic to many naïve predators, including other tadpoles (e.g., Crossland et al., 2008). Indeed, predators exposed to Cane Toad larvae even learn to avoid native

Dodd_Canada_int_5pgs_B1&B2.indd 181

palatable prey a­ fter an encounter with the R. marina larvae (Nelson et al., 2010). Domestic animals, especially cats and dogs that refuse to let go of the toad, are routinely poisoned by the cardiac glycosides in the toad’s secretions (Otani et al., 1969). This can sometimes result in the death of the pet. As long as ­humans wash their hands ­after ­handling a Cane Toad, ­there is minimal danger of poisoning. ­Children, however, should be advised to leave Cane Toads alone. ­After ­handling R. marina, a person should avoid contact with their eyes or mucous membranes. Domestic animals should be treated by a veterinarian. It seems highly unlikely that Cane Toads in Florida and Hawai’i ­will be eradicated. As early as 1956, a Dade County official proposed a bounty on the toad ­because of its threat to pets, but the toad is still pre­sent and even expanding its range. For use at a local level in south Florida, Muller et al. (2020) used an acoustic-­based trap (the Toadinator®, Animal Control Technologies, Australia) that draws R. marina to it using certain aspects of the toad’s call, particularly male choruses, and a treatment that involved a single call at 80 dB with a dominant frequency of 735 Hz and a pulse rate of 18 pulses/sec. Other call treatments using dif­fer­ent dominant frequencies and pulse rates ­were not as effective. The trap worked best on warm, moist nights. As might be expected, captures ­were heavi­ly male biased with 110 males, but only 7 females, captured over 30 nights at 6 trapping sites.

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F ­ amily Craugastoridae

Craugastor augusti (Dugès in Brocchi, 1879) Barking Frog ETYMOLOGY

augusti: a patronym honoring Auguste Henri Andre Dumeril (1812–1870), Professor of Herpetology and Ichthyology at the Museum National d’Histoire Naturelle, Paris; cactorum: from the Greek word kaktos meaning ‘prickly plant’ and the Latin suffix -­orum meaning ‘of the.’ Literally, a frog of the cactus; latrans: from the Latin word meaning ‘barking’ and referring to the frog’s call. NOMENCLATURE

Stebbins (2003): Eleutherodatylus augusti Fouquette and Dubois (2014): Craugastor (Hylactophryne) augusti Synonyms: Eleutherodactylus augusti fuscofemora, Eleutherodactylus augusti, Eleutherodactylus augusti cactorum, Eleutherodactylus augusti latrans, Eleutherodactylus bolivari, Eleutherodactylus cactorum, Eleutherodactylus fuscofemora, Eleutherodactylus latrans, Hylactophryne augusti, Hylodes augusti, Hylodes latrans, Lithodytes latrans IDENTIFICATION

Adults. The ground coloration is brownish gray, tan, reddish brown, or olive gray. ­There is regional variation in ground color, with C. a. cactorum from Arizona primarily brown and C. a. latrans from Texas and New Mexico from sulphur yellow to beige (Goldberg et al., 2004b). A dorsolateral fold is pre­sent, and the granular skin lacks warts or tubercles. A distinguishing fold of skin is pre­sent across the back of the head. Heads are broad. Large brown or black spots may be pre­sent with pale greenish or light centers. The outer and inner surfaces of the limbs are bright yellowish green; this color may extend along the sides of the body. Indistinct bars are pre­sent on the upper surfaces of the limbs (illustrated by Zweifel, 1956b). A black line is pre­sent on the underside of

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the lip. Venters are pale gray or pinkish and dotted in white. Prominent tubercles are pre­sent on the undersides of the feet below the joints. Toes are not webbed. During the calling season, male C. a. cactorum have dark throats with dark tympanums, whereas female throats are white with pink tympanums; male throats ­later change to mottled gray. Sexually dimorphic throat coloration is not pre­sent in C. a. latrans (Goldberg et al., 2004b). Adults are 64–75 mm SUL. Although based on a small sample size, Goldberg (2002) noted males ­were 71.4–74.4 mm SUL and females ­were 77.2–82.8 mm SUL in an Arizona population. Schwalbe and Alberti (1998) mentioned an 85 mm SUL female. The largest Texas male was 77.2 mm SUL, and the largest female was 94 mm SUL (Zweifel, 1956b). Juveniles. Juveniles are patterned differently from adults. They have a dark head and shoulder region, a light band across the ­middle of the back, and a dark posterior third of the body (Zweifel, 1956b; Brennan and Holycross, 2006). Dark spots may be vis­i­ble through the dark dorsal pattern. The band dis­appears in the adults, although it may be very faintly vis­i­ble in some individuals. Throats are white and bellies are translucent. Juveniles in New Mexico have been reported to be strikingly green and ivory or white (Koster, 1946; in Wright and Wright, 1949), or only with the dorsum of the legs yellow tinged in green (in Degenhardt et al., 1996). Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg, and froglets hatch as miniature adults. Eggs. Eggs are deposited terrestrially. They are large, with a dia­meter of 6–7.5 mm. DISTRIBUTION

The Barking Frog occurs from southern Arizona (Santa Rita, Patagonia, Quinlan, Pajarito, Baboquivari, Huachuca mountains, possibly Sierra Ancha), New Mexico (Pecos Valley, Otero County), and west central Texas south to Oaxaca, México. Impor­tant distributional references include: Koster (1946), Bezy et al. (1966), Degenhardt et al. (1996), Dixon (2000, 2013), Murray and Painter (2003), Brennan

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ADULT HABITAT

The Barking Frog is associated almost exclusively with limestone caves, cliffs, crevices, and rock outcroppings in extreme southern Arizona, New Mexico, and Texas (Strecker, 1910b). The frogs rarely move between even adjacent rock outcroppings, and population size on any par­tic­u­lar outcrop is small (2–10 in Goldberg’s, 2002 study). Craugastor augusti is also found associated with rodent burrows in creosote bush flats near limestone and gypsum outcrops in New Mexico (Koster, 1946; Degenhardt et al., 1996) and along bluffs bordering streams in the prairies of west Texas (Strecker, 1910b). In Arizona, their habitat is Oak Woodland–­Grassland ecotones characterized by junipers, manzanita, oaks, sumac, yucca, agave, sotol, and prickly pear cactus (photo­graphs in Bezy et al., 1966). In Arizona, they are found from at least 1,397 to 1,890 m in elevation (Rorabaugh, 2004). Distribution of Craugastor augusti. Craugastor a. latrans occurs in Arizona, with C.a. augusti in Texas and New Mexico. ­These may represent dif­fer­ent species.

and Holycross (2006), Tipton et al. (2012), Bezy and Cole (2014), Murphy (2019), and Holycross et al. (2021). FOSSIL REC­O RD

Fossil Barking Frogs have been reported from Pleistocene deposits in New Mexico, Texas, and Sonora in México (Holman, 1969, 2003). Holman (2003) provided a description of the bones useful in identification of fossils. SYSTEMATICS AND GEOGRAPHIC VARIATION

Craugastor augusti is a member of the Craugastor (Hylactophryne) augusti species series that mostly inhabits México (Hedges et al., 2008). The species has been associated phyloge­ne­tically with the ­Middle American clade of eleutherodactylines (genus Craugastor) (Heinicke et al., 2007); Hedges et al. (2008) elevated this clade to ­family status and placed C. augusti within the subgenus Hylactophryne. The subspecies C. a. latrans has been referred to populations in Texas and New Mexico, and C. a. cactorum has been referred to Arizona populations (Zweifel, 1956b). Craugastor a. latrans is distinguished from C. a. cactorum by differences in relative tympanum size, body size, coloration, mtDNA, and skin toxicity (Zweifel, 1956b; Goldberg, 2002; Goldberg et al., 2004b). Goldberg et al. (2004b) suggested that ­these differences could represent sufficient justification to recognize latrans and cactorum as separate species but did not formally designate them as such. In the United States, 3 dif­fer­ent clades ­were revealed by mtDNA analy­sis within C. augusti (Goldberg et al., 2004b). Additional molecular data are needed from throughout the species’ range.

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TERRESTRIAL ECOL­O GY

Activity is nocturnal, with frogs leaving protected under­ ground retreats only during the summer rainy season. Daylight hours (05:00–17:00) are spent in rock crevices (usually of limestone but also of granite), caves, and even mine tailings. Goldberg (2002) found most frogs ­either on or within 30 m of limestone outcrops, with home ranges of 112.5–2,327 m2 depending upon estimator used. Nocturnal movements ­were usually short, averaging only 4.6 m (range 2.6–7.7 m). Occupied crevices are not oriented in any par­tic­u­lar direction, but west-­facing crevices may provide suboptimal conditions. In Arizona, they return to their subterranean refugia from August to October depending on weather conditions (Goldberg, 2002). Body temperatures range from 15 to 28°C. Barking Frogs are photonegative in their phototactic response, suggesting they are highly nocturnal (Jaeger and Hailman, 1973). CALLING ACTIVITY AND MATE SE­L ECTION

The call is similar to the barking of a dog (“a resounding bark,” Wright and Wright, 1938) or the caw of a raven at a distance, but more of a “whurr” at closer range (Schwalbe and Alberti, 1998). Advertisement calls from Arizona Craugastor a. cactorum are longer in duration (mean 573 ms vs. 343–376 ms), higher in frequency (mean 1,137 Hz vs. 919–995 Hz), and have longer pulses (mean 9.26 ms vs. 4.28–5.6 ms) than ­those of Craugastor a. latrans from Texas (fundamental frequency of 200–300 cps, a duration of 0.24–0.28 sec, and a call rate of 40 calls/min) and New Mexico (Fouquette, 1960; Goldberg et al., 2004b). Calls can be heard by a ­human observer 1.6 km distant in Texas (C. a. latrans) and 600 m in Arizona (C. a. cactorum).

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184  Craugastoridae

Calling occurs from only a 2 to 4 week period in Arizona, with most males calling on the first few nights following the initiation of summer rains (Goldberg, 2002). Calling also is associated with high humidity, no wind, lower temperatures, and high hourly rainfall. In a 2 yr study, Goldberg (2002) heard frogs on only 8 and 16 nights. In contrast, Jameson (1954) reported hearing calls from February to August in Texas. Calls are very loud, but frogs ­will stop calling if an observer approaches within 14–22 m. Calling occurs from within crevices, from ­under rocks, or from small chambers. They also have been observed perched on boulders and calling from late after­noon (17:00) into the eve­ning (Rorabaugh, 2004). Males move up to 50 m from their retreats to call. Calls are often made in tandem whereby 1 frog initiates calling and is followed by other nearby males in sequence. This sequential chorusing extends to all chorusing males, with 1 particularly loud frog taking the role of the leader. If the lead male does not call, a second loud frog ­will assume the leadership role (Jameson, 1954). Males sometime vie back and forth in an apparent calling contest to determine which ­will take the lead; weaker-­voiced males within the chorus then follow. Females in some populations also have a startle vocalization (central Texas) when grasped, whereas ­others do not (New Mexico, west Texas, Arizona) (Jameson, 1954; Goldberg et al., 2004b). OVIPOSITION SITES

Eggs are deposited terrestrially deep in moist cracks, crevices, and fissures, or ­under rocks in the moist earth. The question of adult attendance needs further examination. Males are said to attend to the clutches to protect them from predators and desiccation, even urinating on the clutch to keep it moist (Jameson, 1950). In contrast, the female constructed and attended the nest in captivity, whereas the male remained outside (Streicher and Fujita, 2014). The female constructed the nest by kicking soil into a PVC pipe and then compressed the soil using her body and limbs. Strecker (1910b) noted a pair in amplexus near a stream and incorrectly inferred that eggs ­were deposited in ­water; he suggested that eggs ­were deposited in wet leaves along the stream margins. Other early investigators also have incorrectly assumed aquatic breeding (in Jameson, 1950). REPRODUCTION

Egg clutches are unpigmented and not deposited in a frothy mass. Clutch size is 6–80 eggs (Livezey and Wright, 1947; Schwalbe and Alberti, 1998). Jameson (1950) reported a single clutch of 67 eggs in Texas, and a single clutch of 60 eggs was reported from a New Mexico female

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Juvenile Craugastor augusti have a very dif­fer­ent pattern from adults. Photo: Jay Withgott

(in ­Degenhardt et al., 1996). In captivity, Streicher and Fujita (2014) reported a single clutch of 122, with 119 eggs hatching. Hatching from the eggs as miniature adults is estimated to require 25–35 days (Jameson, 1950). Schwalbe and Alberti (1998) reported a single hatchling of 21 mm SUL, and a 16 mm SUL individual was observed in New Mexico (in Degenhardt et al., 1996). Strecker (1910b) reported eggs being deposited in February in aquatic habitats. He further provided information on tadpoles, suggesting he was confusing this species with some other species. Information from this paper should be used with caution. Livezey and Wright (1947) gave the breeding season as February–­May, but males have been heard calling ­later in the season (Jameson, 1954). DIET

Prey in the scat of Barking Frogs includes centipedes, field crickets, scorpions, leafhoppers, mites, grasshoppers, spiders, ant lions, and katydids (Goldberg, 2002). Other prey includes kissing bugs, cave crickets, and land snails (Bulimulus, Succinea) (Olson, 1959; Schwalbe and Alberti, 1998). Strecker (1910b) reported beetles, ants, and spiders in the diet of Texas frogs. PREDATION AND DEFENSE

Barking Frogs have been described as “prodigious” jumpers when disturbed (Schwalbe and Alberti, 1998). ­These authors noted leaps of 70 cm between boulders, even when carry­ing a radio transmitter. Arizona C. a. cactorum does not possess toxic skin secretions, but C. a. latrans from Texas and New Mexico does (Goldberg et al., 2004b). They also puff up their body to enhance contact with the skin secretions and to make the it difficult for a predator to ­handle them. Jameson (1954) noted that females ­will deliver a “blaring screech” when handled, which presumably serves to startle a wouldbe predator.

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POPULATION BIOLOGY

Using a variety of techniques, Goldberg (2002) and Goldberg and Schwalbe (2004) estimated a survival probability of 66% between years and a capture probability of 13% for a population of C. augusti in the Huachuca Mountains of Arizona. Survival rate was estimated at 93%. ­These latter authors further estimated density at 2 populations as 1 frog

Rocky habitat favored by Craugastor augusti. Photo: Cecil Schwalbe

per 736 m2 at 1 location and 1 per 3,507 m2 at another location, noting that it took 7.6 hrs to find a frog at the first and 5.1 hrs at the second. Longevity is unknown but likely longer than most frogs; frogs have lived for 11 yrs in captivity (Murphy, 2019). Adult Craugastor augusti. Photo: James Stuart

DISEASES, PARASITES, AND MALFORMATIONS

No information is available. SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Adult male (left) and female (right) Craugastor augusti. Note the difference in tympanum coloration. Photo: Cecil Schwalbe

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Sampling and monitoring this species in difficult terrain is not an easy task considering its ­limited calling and period of activity. Distance sampling, visual encounter surveys, and call surveys are not useful, and mark-­recapture is ­labor intensive and does not give sufficient statistical power for monitoring (Goldberg, 2002; Goldberg and Schwalbe, 2004). Using an occupancy approach focusing on rock outcrops might be the best method to follow status and trends. Fortunately, the species is abundant throughout much of its extensive range in México.

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F ­ amily Eleutherodactylidae

Eleutherodactylus cystignathoides (Cope, 1878 “1877”) Rio Grande Chirping Frog ETYMOLOGY

cystignathoides: According to Lynch (1970), the name is derived from Cystignathus, an old generic name for several leptodactylid frogs. campi: a patronym honoring R.D. Camp who presented the type specimen to the US National Museum. NOMENCLATURE

Frost et al. (2006a): Syrrhophus cystignathoides Fouquette and Dubois (2014): Eleutherodactylus (Syrrhophus) cystignathoides Synonyms: Eleutherodactylus campi, Eleutherodactylus cystignathoides campi, Phyllobates cystignathoides, Syrrhophus campi

Eggs. The eggs are small and unpigmented, mea­sur­ing 3–5 mm in dia­meter (Livezey and Wright, 1947; Hayes-­ Odum, 1990). DISTRIBUTION

The Rio Grande Chirping Frog is found naturally from the Rio Grande embayment south to Nuevo Leon, Tamaulipas, San Luis Potosi, and Veracruz in México. Scattered populations are reported from at least 49 counties in east and central Texas (Quinn, 1979; Lott, 2012, 2014; Swanson and Swanson, 2017) and 6 parishes in Louisiana (Hardy, 2004; Lott, 2012, 2014; Boundy and Carr, 2017). Other isolated populations occur in Mobile County, Alabama (McConnell et al., 2015) and Maricopa County, Arizona (Holycross et al., 2021). This species has been widely introduced throughout Texas and Louisiana via the nursery plant trade, and individuals are likely to be encountered throughout the northwest Gulf region. Impor­tant distributional references

IDENTIFICATION

Adults. This is a small, rather nondescript frog with a brownish gray to brownish green ground color. The body is elongate and flattened, and the snout is pointed. A dark bar is pre­sent from the eye to the end of the nose, but ­there are no interorbital bars. Dark spots are pre­sent dorsally on a finely granular skin. The rear part of the body has irregular flecking. Vertical bars are pre­sent on the legs. Toes on the digits are only slightly expanded. Venters are smooth and translucent, and the ventral vein appears as a dark line down the ­middle of the belly. Adult males are 16.3–23.5 mm SUL, whereas females are 16–25.8 mm SUL (Lynch, 1970). Eleven individuals captured in Houston ranged from 7 to 23.7 mm SUL (Quinn, 1979). Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg, and froglets hatch as miniature adults. Hayes-­Odum (1990) described development within the egg.

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Distribution of Eleutherodactylus cystignathoides. This species is rapidly expanding its range via horticultural introductions. An isolated introduced population also occurs in Maricopa County, Arizona (not pictured).

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Eleutherodactylus cystignathoides 187

are Dixon (2000, 2013), Lott (2012, 2014), Tipton et al. (2012), Boundy and Carr (2017). FOSSIL REC­O RD

Fossil Rio Grande Chirping Frogs are reported from Pleistocene deposits in Tamaulipas, México (Holman, 2003). Holman (2003), however, noted that osteological characters useful in identification of fossils of this species are not available. SYSTEMATICS AND GEOGRAPHIC VARIATION

Rio Grande Chirping Frogs ­were described as Syrrhophus campi by Stejneger (1915), and ­were ­later assigned as a subspecies of S. cystignathoides (Martin, 1958). This relationship needs to be reexamined using molecular analy­sis. In his 1970 review of the genus Syrrhophus, Lynch placed this species in the Leprus species group. Hedges (1989) synonymized the genus Syrrhophus with Eleutherodactylus, and Heinicke et al. (2007) noted that for the Ca­rib­bean clade of eleutherodactyline frogs, including frogs formerly considered Syrrhophus, the proper generic name was Eleutherodactylus. Hedges et al. (2008) placed E. cystignathoides within the Eleutherodactylus (Syrrhophus) leprus species series. Some authors, however, continue to recognize Syrrhophus as a genus rather than as a subgenus (e.g., Tipton et al., 2012). ADULT HABITAT

The Rio Grande Chirping Frog is found along sandy river alluvium rather than in rock outcrops as are the other Texas chirping frogs. Wright and Wright (1938) described the habitat as “moist earth ­under boards, brick, or stone piles, ­under walks, or any cover of yard, field, grass, or brush.”

rock walls, and buildings also are used. Calling occurs both in rainy and dry weather, but calling is more intensive on rainy nights. Females move ­toward calling males and may initiate amplexus by lightly touching the males. An amplexed pair may move around ­until an appropriate oviposition site is found. OVIPOSITION SITES

Oviposition occurs at or just below the substrate surface in moist soil or ­under surface debris. Hayes-­Odum (1990) found eggs in a shallow flowerbed, and in captivity, eggs ­were deposited at or just below the soil surface. REPRODUCTION

The clutch size is 5–13 with a breeding season from April to May (Wright and Wright, 1938; Livezey and Wright, 1947; Tipton et al., 2012). It may be that this species deposits multiple clutches over a longer breeding season than current information suggests. Eggs are deposited in a clump, which the female then spreads around and covers using her rear legs (Hayes-­Odum, 1990). Development takes 14–16 days at 27–33ºC. Hatchlings are dark brown and ca. 6 mm SUL (Hayes-­Odum, 1990). DIET

The diet includes small beetles and spiders (Hardy, 2004), but a comprehensive assessment of diet has not been undertaken. Tipton et al. (2012) reported a frog regurgitating cockroach eggs. PREDATION AND DEFENSE

This frog is very ­adept at ­running or scurrying ­under vegetation and into rock crevices to avoid capture.

TERRESTRIAL ECOL­O GY

POPULATION BIOLOGY

Nothing has been reported on the terrestrial ecol­ogy of this species.

No information is available.

CALLING ACTIVITY AND MATE SE­L ECTION

In Louisiana, Hardy (2004) reported calling in April and from August to November, but Tipton et al. (2012) noted calling at any time during the warm months. Calling occurred mostly at night, but calling also occurred in the morning from 07:30 to 09:15 hrs. The minimum air temperature at which calling takes place is ca. 20ºC (Hayes-­Odum, 1990). Early in the eve­ning, calling occurs from perches as high as 22 cm above the substrate (Hayes-­Odum, 1990). As the night progresses, most calling occurs at lower perch heights and from beneath vegetation, where males call from crevices in leaf litter and mulch. The crevices appear to penetrate into the soil, and frogs retreat deeper into the burrows and tunnels should they be disturbed or ­after calling ceases at dawn. Crevices in sidewalks,

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Adult Eleutherodactylus cystignathoides. Photo: Kenny Wray

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188  Eleutherodactylidae

DISEASES, PARASITES, AND MALFORMATIONS

Unidentified nematodes ­were reported by Hardy (2004). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Population status and trends have not been investigated. However, the species is widely distributed and expanding its range due to the nursery trade.

Habitat of Eleutherodactylus cystignathoides. Brazos County, Texas. Photo: Gary Nafis

Eleutherodactylus guttilatus (Cope, 1879) Spotted Chirping Frog ETYMOLOGY

Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg and froglets hatch as miniature adults. Eggs. The eggs are small and unpigmented, mea­sur­ing ca. 4 mm in dia­meter (Gaige, 1931). However, Gaige may have been referring to E. marnockii since she considered populations in central Texas to be conspecific with ­those in the Chisos Mountains.

guttilatus: from the Latin guttula meaning ‘spotted’ or ‘flecking.’

DISTRIBUTION

NOMENCLATURE

Frost et al. (2006a): Syrrhophus guttlilatus Fouquette and Dubois (2014): Eleutherodactylus (Syrrhophus) guttilatus Synonyms: Eleutherodactylus petrophilus, Eleutherodactylus smithi, Malachylodes guttilatus, Syrrhophus gaigeae, Syrrhophus petrophilus, Syrrhophus smithi Information on E. guttilatus is frequently included with E. marnockii, as the species ­were once considered conspecific. Collecting locations should be verified when using the lit­er­a­ture.

Spotted Chirping Frogs are known only from the Big Bend region of Texas and several scattered localities in Sierra Madre Oriental in México. Dixon (2000) also mentioned calls heard from the east side of the Davis Mountains.

IDENTIFICATION

Adults. This is a small frog with a wide head and light brown to black flecking and vermiculation on the dorsal pattern. The ground color is cream, gray, or brown. The eyes are prominent, and an interorbital bar is pre­sent. The dorsal skin is smooth. The thighs are usually not banded, and the digital pads are slightly expanded. Adult males are 20.6–29 mm SUL, whereas females are 25.7–31 mm SUL (Lynch, 1970). Gaige (1931) reported individuals 24–32 mm SUL, and Milstead et al. (1950) gave 20.9–26.5 mm SUL for 4 adults.

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Distribution of Eleutherodactylus guttilatus

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Eleutherodactylus guttilatus 189

Impor­tant distributional references include Dixon (2000, 2013), Dayton et al. (2007), and Tipton et al. (2012). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

In his 1970 review of the genus Syrrhophus, Lynch placed this species in the Marnockii species group. Hedges (1989) synonymized the genus Syrrhophus with Eleutherodactylus, and Heinicke et al. (2007) noted that for the Ca­rib­bean clade of eleutherodactyline frogs, including frogs formerly considered Syrrhophus, the proper generic name was Eleutherodactylus. Hedges et al. (2008) placed E. guttilatus within the Eleutherodactylus (Syrrhophus) marnockii species group. Some authors, however, continue to recognize Syrrhophus as a genus rather than as a subgenus. Schmidt and Smith (1944) described Syrrhophus gaigae from the Chisos Mountains, but the name S. guttilatus (Cope, 1879) has nomenclatural priority.

Adult Eleutherodactylus guttilatus. Photo: Robert Hansen

ADULT HABITAT

This species is found ­under rocks, on rock bluffs, among rockslides, and along stream beds in canyons (Gaige, 1931; Jameson and Flury, 1949). TERRESTRIAL ECOL­O GY

Activity appears to be stimulated by rainfall. CALLING ACTIVITY AND MATE SE­L ECTION

The call is described as a metallic note repeated at irregular intervals (Gaige, 1931), but Dayton et al. (2007) noted ­there are actually 2 distinct calls: a short, sharp chirp and a 1–2 sec trill. Calling occurs in the eve­ning ­after rain at temperatures of 15.6–26.7°C (Dayton et al., 2007). OVIPOSITION SITES

Oviposition sites are unknown but are presumed to be in moist rock crevices or in cavities beneath the soil ­under rocks. REPRODUCTION

Breeding occurs from April to July (Dayton et al., 2007). Individual clutches contain 6–20 eggs. A female with 5 eggs was observed on 20 July (Gaige, 1931). DIET

Few data are available. Gaige (1931) observed ants, beetles, termites, and an isopod in the gut of a single female. PREDATION AND DEFENSE

Habitat of Eleutherodactylus guttilatus, Big Bend National Park. Photo: C.K. Dodd, Jr.

Like other chirping frogs, E. guttilatus is agile and rapidly leaps to safety at the approach of an observer or predator.

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190  Eleutherodactylidae

POPULATION BIOLOGY

SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available.

No information is available.

DISEASES, PARASITES, AND MALFORMATIONS

STATUS AND CONSERVATION

The mite Hannemania hylae is an ectoparasite of this species in Texas (Gaige, 1931; Lynch, 1970; Jung et al., 2001).

No information is available. Dayton et al. (2007) suggested that the species might be more widely distributed than is currently recognized.

Eleutherodactylus marnockii (Cope, 1878) Cliff Chirping Frog

2013), Tipton et al. (2012), Owen et al. (2014), and Davis and LaDuc (2018).

ETYMOLOGY

marnockii: a patronym in honor of G.W. Marnock of Helotes, Texas, who collected the original specimen.

FOSSIL REC­O RD

Cliff Chirping Frogs are reported from Pleistocene deposits in a variety of locations in Texas (Holman, 1969, 2003). Holman (2003) provided a description of the ilium useful in identifying fossils of this species. SYSTEMATICS AND GEOGRAPHIC VARIATION

NOMENCLATURE

Frost et al. (2006a): Syrrhophus marnockii Fouquette and Dubois (2014): Eleutherodactylus (Syrrhophus) marnockii Synonyms: Eleutherodactylus marnockii, Hylodes marnockii, Syrrhophus marnochii IDENTIFICATION

Adults. Adults are small, stout bodied and flattened, mea­sur­ ing 19–38 mm SUL. The head is noticeably large. The ground color is greenish brown with dark flecks throughout the dorsum. Mohr (1948) noted that individuals appeared white in caves but turned a normal color in light. An interorbital bar is pre­sent. The dorsal skin is smooth to weakly pustular, but the venter is smooth and white. The thighs are banded, and the digital pads are expanded. Adult males are 18.4–28.9 mm SUL, whereas females are 20.4– 35.4 mm SUL (Lynch, 1970). Milstead et al. (1950) reported adults from the Edwards Plateau at 18–32.7 mm SUL and the Stockton Plateau at 28.8–37.7 mm SUL. Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg, and froglets hatch as miniature adults. Eggs. The eggs are small and unpigmented, mea­sur­ing ca. 4 mm in dia­meter (Livezey and Wright, 1947).

In his 1970 review of the genus Syrrhophus, Lynch placed this species in the Marnockii species group. Hedges (1989) synonymized the genus Syrrhophus with Eleutherodactylus, and Heinicke et al. (2007) noted that for the Ca­rib­bean clade of eleutherodactyline frogs, including frogs formerly considered Syrrhophus, the proper generic name was Eleutherodactylus. Hedges et al. (2008) placed Eleutherodactylus marnockii within the Eleutherodactylus (Syrrhophus) marnockii species group. Some authors, however, continue to recognize Syrrhophus as a genus rather than as a subgenus (e.g., Tipton et al., 2012).

DISTRIBUTION

This species is endemic to Texas and is known only from the Edwards Plateau and the edge of the Stockton Plateau as far west as the Sierra Vieja. Calls also have been reported from Upton County. Impor­tant distributional references include Milstead et al. (1950), Dixon (2000,

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Distribution of Eleutherodactylus marnockii

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Eleutherodactylus marnockii 191

ADULT HABITAT

CALLING ACTIVITY AND MATE SE­L ECTION

This is a species of limestone caves and crevices throughout the Edwards and parts of the Stockton plateaus. ­These areas are heavi­ly dissected by canyons and stream channels surrounded by blackland prairie. The topography consists of escarpments, terraced hills, bluffs, talus slopes, rolling grasslands, and rock outcrops. Jameson (1955) discussed topography and vegetation in connection with E. marnockii occupancy in the dif­fer­ent geologic formations, and Milstead et al. (1950) noted its presence in the cedar-­oak association on the Stockton Plateau. Rock crevices in the vicinity of creeks appear to be preferred habitat. The species is common in caves (Nicholson, 1932; Mohr, 1948; Jameson, 1955). Cliff Chirping Frogs are terrestrial, inhabiting rock outcrops and the substrate adjacent to rocky cliffs. They also have been found to 2.4 m off the ground in trees and Smilax vines (Jameson, 1955), and along stream banks. Cliff ­faces may be isolated from one another by grasslands, cedar breaks, and deep soils.

Calling occurs from protected sites, such as cracks, fissures, caves, and from ­under ­houses. Most calling occurs at night but calls occasionally are heard during the day. Even when in amplexus, a male may continue calling. The usual call is likened to a cricket chirping and can be heard by ­human observers up to 30 m distant. Calls are often made in tandem among individuals (trios or quartets), whereby 1 frog initiates calling followed by other nearby males in sequence. If the lead male does not call, the ­others also do not call (Jameson, 1954, 1955). This sequential calling pattern is heard throughout the year and not just during the breeding season. It is pos­si­ble that the sequence represents a territorial system or a means of nearest-­neighbor recognition. Jameson (1955) noted that the advertisement call is much more rapid, clear, and sharp, and is only made during the breeding season. This latter call is pronounced when males are already in amplexus or when ­there is a female close by. A male approaches a female and scratches her back, neck, and legs with his hind legs. If the female does not move away, he ­will clasp her in axillary amplexus. The male continues to stimulate the female with his hind legs. She ­will dig a trench with her forelegs and deposit the eggs as the clasped pair move along the trench. Both the male and female cover the eggs with soil ­after they are deposited.

TERRESTRIAL ECOL­O GY

Activity is primarily nocturnal, beginning shortly ­after dusk, with peak activity periods from April to early June and from September to October corresponding to moderate temperatures. Cliff Chirping Frogs are more active when rainfall is above normal than during droughts, and spring drought may delay spring activity into early summer. Some activity, however, occurs year-­round, with frogs becoming active in winter as temperatures rise following a period of cold weather. Frog activity also is associated with high humidity, low wind, low light intensity, and high substrate moisture (Jameson, 1955). Frogs may be observed moving sluggishly in crevices at temperatures of 4.4–10°C. At higher temperatures, they move around much more often. Jameson (1955) observed frogs from 1.1 to 33.3°C and at a humidity of only 35%. Other than juvenile dispersal, movements prob­ably are ­limited, with most frogs remaining in the immediate vicinity of 1 location their entire life. Indeed, breeding season home ranges averaged only 0.034–0.07 ha at 12 Texas sites, with no differences between the sexes (Jameson, 1955). The largest home ranges occurred on open slopes. In Jameson’s study, marked juveniles dispersed 112.5–300 m (mean 211 m), with most frogs moving parallel to the bluff. A few frogs established immediate residency. Dispersal likely depends on topography, available habitat, weather conditions, and time of year. Jameson (1955) further showed that dispersal readily occurs into areas of low population density and away from areas of high population density.

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OVIPOSITION SITES

Despite intensive searches, Jameson (1955) was unable to locate egg clutches in nature. Presumably they are deposited deep in moist soil in protected crevices and fissures under­ground. REPRODUCTION

Gravid females have been reported from February to December, suggesting an almost year-­round breeding season. Most breeding occurs during the spring and fall rains, based on the presence of tiny juveniles; juveniles have been observed from February to November (Jameson, 1955). Eggs are deposited shortly ­after mating. Jameson (1955) found no evidence of nest guarding. Individual clutches contain 6–20 eggs. Multiple clutches may be oviposited, suggesting a total reproductive output of 60 or more eggs per season. DIET

Cliff Chirping Frogs eat a variety of invertebrates, including ants, beetles, crickets, termites, and spiders (Jameson, 1955). They likely consume any small animal they can. PREDATION AND DEFENSE

As with other chirping frogs, this species is ­adept at ­running into cracks and crevices in limestone in order to avoid

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192  Eleutherodactylidae

capture. Natu­ral predators include snakes (Agkistrodon contortrix, Crotalus atrox, Thamnophis eques) and wolf spiders (Jameson, 1955). POPULATION BIOLOGY

Females reach sexual maturity at 19–22 mm SUL, whereas males reach maturity at 18–22 mm SUL (Jameson, 1955). Sex ratios are difficult to determine ­under natu­ral circumstances. Of 245 frogs marked by Jameson (1955), 48 ­were juveniles, 66 ­were females (with eggs), and 141 ­were classified as “probable males.” For another study, Jameson had collectors search for frogs and found 57 males and 53 females. Density varies among populations, with Jameson’s 12 study sites containing densities of 1.2–8.9 frogs per 0.45 ha. Density estimates also vary annually and within a season. Populations appear greater in the spring than in the fall. Jameson (1955) estimated ­there ­were from 3 to 31 frogs per “average home range” at 12 study locations. Population

Habitat of Eleutherodactylus marnockii. Photo: Dirk Stevenson

turnover was considerable between seasons and years, but sample sizes (years and locations observed) ­were very small. Jameson (1955) suggested a complete population turnover ­every 2.5–3 yrs. DISEASES, PARASITES, AND MALFORMATIONS

The mite Hannemania hylae is an ectoparasite of this species (Lynch, 1970). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Adult Eleutherodactylus marnockii. Photo: Dirk Stevenson

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This species appears to be reasonably common throughout its range, but ­there are no studies on its status and population trends. Acute artificial lighting alters calling be­hav­ior by decreasing the number of males calling in a chorus and decreasing the intensity of calling (Hall, 2016).

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F ­ amily Hylidae

Acris blanchardi Harper, 1947 Blanchard’s Cricket Frog Rainette grillon de Blanchard ETYMOLOGY

blanchardi: a patronym honoring Frank N. Blanchard (1888–1937), a herpetologist at the University of Michigan. NOMENCLATURE

Synonyms: Acris crepitans blanchardi, Acris crepitans paludicola, Acris gryllus blanchardi, Acris gryllus paludicola, Hyla ocularis blanchardi ­There is considerable confusion in the primary lit­er­a­ture about ­whether the species A. crepitans or A. blanchardi is being discussed. ­Until 2008 (­Gamble et al., 2008), ­these taxa ­were considered subspecies of a single wide-­ranging species, and many authors did not make a distinction between them. Even as late as 2020, some authors continue to lump ­these species together (e.g., Sonn et al., 2019, 2020). Geo­graph­i­cal overlap may not be extensive, however, although some distributional details remain to be determined. IDENTIFICATION

Adults. This is a slender, small frog (maximum 35 mm SUL) with a blunt snout and at least 1 black line on the posterior portion of the thigh. The upper black line on the back of the thigh is usually irregular; the lower area of the thigh consists of black stippling suggestive of a second line. Background colors are brown to gray. The dorsal stripes of Blanchard’s Cricket Frogs are variable in coloration, with colors of gray to brownish red to green vertebral stripes. Down the back, the stripe is often bordered on ­either side by black spots. ­These colors may be vivid to pale, and the band itself can be continuous or interrupted. A few rare individuals have both red and green dorsal coloration. The coloration is not related to sex and can be variable within the population. Even frogs with a nongreen vertebral stripe may have occasional green

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spots, and frogs rarely have a red-­green stripe (Pyburn, 1961a). The dorsal surface of the skin has a profusion of tiny warts of irregular size. Small, black spots may be pre­sent, which are rimmed in white. A dark triangle is pre­sent between the eyes, although in some individuals the triangle may appear pale. Upper jaws are black to brown, and from the ­angle of the eye to the jaw, a white line is usually pre­sent. Bellies are white, and some individuals have a very pale yellow throat. The undersides of limbs are white. The tips of the fin­gers have slightly enlarged disks and the fin­gers lack webbing. The feet are well webbed. Males have a yellow to dark subgular vocal pouch. ­There are no morphological characters that definitively separate A. crepitans from A. blanchardi. Females are slightly larger than males. In Wisconsin, adults range from 20 to 30 mm SUL (mode 25–26 mm) (Dernehl, 1902). In South Dakota and Nebraska, males have a mean SUL of 22.6 mm (range 15.5–26.7 mm), whereas the female mean is 24.5 mm SUL (range 16.3–31.5 mm SUL) (McCallum and Trauth, 2004). In Indiana, males are 20.3–25.5 mm SUL (mean 22.6 mm), whereas females are 20–30 mm SUL (mean 24.5 mm) (Minton, 2001). Nevo (1973b) recorded the following mean male and female SUL lengths in mm: west Texas (25.5, 28.7), central Texas (23.6, 26.4), west Gulf Coast Plain (22.5, 24.0), northern interior lowlands (22.5, 24.3) and northern ­Great Plains (25.8, 27.2). Larvae. Larvae are olive to brown, and are speckled with small black markings. The mottled upper and lower tail fins are narrow, and the tail musculature may have a dark bordering line at the junction between the upper tail fin and the tail musculature. The tail usually has a con­spic­u­ous black tip, as in other cricket frog tadpoles. It is also much longer than the body and has a pointed tail tip. The throat is gray in the center, and the body is mottled ­toward the side and belly; the belly itself is white to pale yellow. The total length may reach 40–50 mm. Cricket frogs have the distinction of having the largest tadpole in relation to metamorph size of all North American anurans. A description of the tadpole, contrasting it with larval A. gryllus, is in Orton (1947).

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194  Hylidae

Eggs. The eggs are small and dark brown to black dorsally and somewhat tan ventrally. The mean dia­meter of the vitellus is 0.85–1.0 mm in Ohio (Dickson, 2002) and 1.13 mm in Texas (range 1.06–1.17 mm). ­There are 2 gelatinous envelopes surrounding the egg, although the inner envelope may not be vis­i­ble without the aid of a microscope. The inner envelope is 2.38–2.95 mm in Ohio (Dickson, 2002) and 2.34–2.74 mm (mean 2.6 mm) in Texas; the outer envelope is 2.98–3.7 mm (mean 3.34 mm). The total number of eggs ranges between ca. 150 to >400, but the sample sizes of ­actual counts are small. The eggs ­were described by Livezey (1950). DISTRIBUTION

Blanchard’s Cricket Frog occurs from central Ohio southward into western West ­Virginia and northern Kentucky. Morse (1904) reported A. blanchardi from several locations in Ohio where Walker (1946) could not relocate them. The species has not been observed in West ­Virginia since 1948 and is apparently extirpated within the state (Dickson, 2002). The range once included a small part of southwestern Ontario (now extirpated; Choquette and Jolin, 2018). Populations occur in southern Michigan and Wisconsin and extend up the Mississippi River to Washington and adjacent counties in eastern Minnesota (Moriarty et al., 1998; Casper et al., 2017). Populations in southwestern Minnesota ­were

Distribution of Acris blanchardi. An isolated population once existed in extreme southeastern Arizona but is no longer extant. The hatched area in Mississippi indicates uncertainty in the identity of Acris in this region.

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thought to be extirpated, but ­there are recent rec­ords from Rock County (Smith, 2018). The range includes southeastern South Dakota and most of Iowa, south through much of central Nebraska, most of Kansas and Oklahoma, and into the eastern three-­quarters of Texas and the adjacent Rio Bravo drainage in México. ­There ­were populations along the Republican and Platte rivers in northern Colorado, but ­these dis­appeared in the 1970s, and the last cricket frog was observed in Colorado in 1979 (Hammerson, 1999; Hammerson and Livo, 1999). The range extends into New Mexico along the Pecos River, and A. blanchardi may have once reached as far west as Cochise County, Arizona (Frost, 1983; Murphy, 2019); this population at the springs and cienegas of San Bernardino no longer exists. To the east, A. blanchardi is found throughout Louisiana (except the Florida Parishes) and may cross into Mississippi in the counties bordering the Mississippi River in the Delta Region. ­There is no evidence, however, of it crossing the Mississippi and Ohio rivers between Memphis and the vicinity of Louisville, Kentucky. Many A. blanchardi populations have dis­appeared since the 1970s, and the current range no longer includes much of the area previously occupied, especially in the West and upper Midwest. Blanchard’s Cricket Frogs have been recorded on North Bass, ­Middle Bass, Kelley’s, and Pelee (not seen since 1987) islands in Lake Erie (Walker, 1946; Logier and Toner, 1961; Langlois, 1964; Weller and Green, 1997; Davis and Menze, 2000; Hecnar et al., 2002). Heavy pesticide use on Ontario’s Pelee Island might have exterminated A. blanchardi ­there (Russell et al., 2002). Impor­tant distributional references include: Arizona (Frost, 1983), Arkansas (Burt, 1935; Trauth et al., 2004), Colorado (Hammerson, 1999; Hammerson and Livo, 1999), the ­Great Lakes region (Harding, 1997; Harding and Mifsud, 2017), Illinois (Schmidt and Necker, 1935; Smith, 1961; Phillips et al., 1999), Indiana (Minton, 2001), Iowa (Hemesath, 1998), Kansas (Smith, 1934; Collins, 1993), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Michigan (Ruthven, 1912), Minnesota (Oldfield and Moriarty, 1994; Moriarty and Hall, 2014; Casper et al., 2017), Missouri (Johnson, 2000; Daniel and Edmond, 2006), Nebraska (Hudson, 1942; Lynch, 1985; McCallum and Trauth, 2004; Ballinger et al., 2010; Fogell, 2010), New Mexico (Van Denburgh, 1924; Degenhardt et al., 1996), Ohio (Walker, 1946; Pfingsten, 1998; Davis and Menze, 2000), Oklahoma (Burt, 1935; Sievert and Sievert, 2006), South Dakota (Ballinger et al., 2000; McCallum and Trauth, 2004; Naugle et al., 2005; Kiesow, 2006; Burdick and Swanson, 2010), Texas (Hardy, 1995; Dixon, 2000, 2013; Tipton et al., 2012, as ­either or both A. crepitans and A. blanchardi), West ­Virginia (Green and Pauley, 1987), and

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Acris blanchardi 195

Wisconsin (Suzuki, 1951; Vogt, 1981; Mossman et al., 1998). Dixon (2000) and Trauth et al. (2004) include a separate map for A. crepitans, but only A. blanchardi occurs in Texas and Arkansas, respectively (­Gamble et al., 2008). The maps in Dundee and Rossman (1989) and Boundy and Carr (2017) include both A. crepitans and A. blanchardi. FOSSIL REC­O RD

Pliocene fossils attributed to A. blanchardi (as A. crepitans) are known from Texas, as are Pleistocene Irvingtonian fossils. Pleistocene Rancholabrean fossils are more common, having been described from many localities in Kansas and Texas (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

Acris blanchardi has long been considered a subspecies of A. crepitans based on morphological data (Harper, 1947). However, McCallum (2003) and McCallum and Trauth (2006) reported on the results of an extensive morphological study (including analyses of toe webbing, spot patterns, stripe patterns, presence of warts, morphological mea­sure­ ments) and concluded that ­there ­were no meaningful differences between ­these taxa; they recommended that A. c. blanchardi not be recognized as a valid taxon. Using both mtDNA and nDNA, however, ­Gamble et al. (2008) concluded that ­there ­were enough differences in the molecular data to warrant recognition of A. blanchardi as a full species. ­There are also regional differences in physiological variables between the species (e.g., Salthe and Nevo, 1969). The Coastal Cricket Frog, A. gryllus paludicola, was described by Burger et al. (1949), based on individuals found from near Sabine Pass, Jefferson County, Texas. The subspecies was described based on its relatively smooth skin, a lack of or reduced presence of anal warts, ill-­defined postfemoral stripes, and a rose-­pink coloration of the male’s vocal sac. Based upon a reanalysis of 7 newly found individuals from the type locality, Rose et al. (2006) concluded that recognition of the subspecies was warranted based on mtDNA comparisons, but that it was allied with A. crepitans instead of A. gryllus. ­Gamble et al. (2008), however, did not recognize the subspecies, since it was nested within the A. blanchardi clade. Ward et al. (1987) examined 16 enzyme loci for populations in north central and south central Texas. Ten of the loci ­were monomorphic. They concluded that differentiation among remaining loci was low between northern and southern populations. However, ­there was a considerable amount of allelic heterozygosity, indicating high levels of differentiation at the population level. Ge­ne­tic similarity was not related to geographic proximity, and gene flow between even adjacent populations was minimal. Ward et al. (1987)

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hypothesized that inbreeding and/or stochastic events related to environmental ­factors ­were responsible for the maintenance of the ge­ne­tic variation. In Kansas, Gorman and Gaines (1987) examined 17 loci from 16 populations occurring across the state. Some populations ­were found along the margins of ponds, whereas ­others occurred along streams. Allelic frequencies varied considerably among populations, and differences ­were evident in the frequencies of alleles between adults and juveniles. ­These differences ­were maintained across years and suggested strong se­lection. In general, ge­ne­tic variation declined ­toward the western portion of the state. It appears that temperature and moisture exert significant influence upon allele frequencies. In addition, ­there ­were strong differences in allele frequencies between cricket frog populations living at ponds and ­those living along streams. ­There ­were no differences in allele frequencies among dif­fer­ent color-­morph stripes (Gorman, 1986). Gorman and Gaines (1987) hypothesized that se­lection operated on physiological characteristics of Blanchard’s Cricket Frogs rather than through predator-­mediated se­lection. Whereas Nevo and Capranica (1985) differentiated 3 dif­fer­ent call patterns in Acris, roughly corresponding to forests (A. crepitans), grasslands (A. blanchardi), and pinelands (A. gryllus), it is clear that ­these patterns do not hold in forested areas in the southern part of the range of A. blanchardi. In Louisiana and Texas, for example, A. blanchardi has a “forest” call similar to A. crepitans, where populations occur in forested habitats. Similarities in calls relate more to habitat characteristics (open versus closed habitats) than to phylogeny. Two basic patterns of dorsal stripe coloration in A. blanchardi are—­from a ge­ne­tic point of view—­green and nongreen (red or gray). The red stripe appears dominant to gray, but its ge­ne­tic relationship to green stripes is unclear. The green coloration is determined by ­either a heterozygous or homozygous condition at a single gene, and the green stripe results from ­simple dominance (Pyburn, 1961a). Thus, crossing a pair of heterozygous green-­stripe frogs should produce a roughly 3:1 ratio of green to nongreen F1 offspring, and this prediction is easily verified in laboratory crosses (Pyburn, 1961b). Dorsal coloration can include vari­ous degrees of color intensity of the grayish, red, and green morphs. Vogt (1981) even has a photo­graph of a very light colored yellowish-­ brown individual. In Wisconsin, for example, Dernehl (1902) recorded 8 frogs with very ­little red coloration, 3 with medium shades of red, and 12 with intense shades of red. Corresponding values for the green morph ­were 6, 1, and 5. Thirty-­five of the 100 frogs he examined had at least some shades of red or green. Nongreen morphs predominate. For

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example, in Texas, 83% of 521 recent metamorphs w ­ ere nongreen and 17% ­were green (Pyburn, 1961b), whereas in Indiana, Isaacs (1971) recorded 273 gray-­striped frogs, 134 red-­striped frogs, and only 110 green-­striped frogs. Pyburn (1961b) reported on an additional collection in Texas in which 28% ­were green and 72% ­were nongreen. In Kansas, western populations are composed almost entirely of gray-­striped frogs, with the proportions of green-­and red-­striped frogs increasing ­toward the east (Gorman, 1986). Pattern polymorphism is common in cricket frogs. Some authors have suggested that ­there are dif­fer­ent se­lection pressures on frogs in seasonally changing backgrounds or habitats. Burkett (1984) noted that the proportion of green-­striped frogs seemed highest when ­water levels ­were high and vegetation was abundant. Green frogs also ­were scarcer in open situations than in wooded habitats. As with Pseudacris maculata, the frequencies of ­these morphs may change seasonally and annually within a population (Nevo, 1973a), and may reflect dif­fer­ent se­lection pressures depending on seasonal changes in background coloration. However, color changes do not vary seasonally in all populations (Isaacs, 1971; Gray, 1983; Gorman, 1986), and ­there may be dif­fer­ent escape be­hav­iors associated with dif­fer­ent stripe colors. For example, Wendelken (1968) noted that red-­ striped A. blanchardi tended to jump more often ­toward ­water than land (but see Gray, 1978). Thus, pattern polymorphism in cricket frogs could be maintained via substrate matching, predator avoidance, and dif­fer­ent escape be­hav­ iors (Milstead et al., 1974). Not all authors agree. In rather ­simple experiments, Gray (1978) could find no differences among color morphs in predation susceptibility, substrate or ­water preferences, or in distribution along the shoreline. He found no differences in color morphs in desiccation rates, re­sis­tance to prolonged stress, thermal preference, or CTmax among cricket frogs in Illinois (Gray, 1977). Fi­nally, he found no differences in movement patterns, dispersal, growth, survivorship, or seasonal frequencies of stripe pattern at vari­ous study sites (Gray, 1983, 1984). However, ­there ­were differences geo­graph­i­cally and annually in color-­morph frequencies among locations. Isaacs (1971) also found no seasonal differences in the frequencies of color morphs over a 3 month study at a single site in Indiana. Gray (1983, 1984) suggested ­simple chance might account for variation in color-­morph frequencies in Blanchard’s Cricket Frog, as effective population size was small in his Illinois populations. A.P. Blair (in Moore, 1955) attempted to hybridize female “A. gryllus crepitans” (presumably A. blanchardi) with male Pseudacris streckeri and P. clarkii. The attempt was unsuccessful.

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ADULT HABITAT

Acris blanchardi is a frog of open habitats throughout much of its western range. In the Midwest and in east Texas and adjacent Louisiana, Blanchard’s Cricket Frogs are found around permanent ponds, lakes, marshes, and small streams surrounded by both deciduous and piney woods habitats with substantial amounts of edge habitat (Knutson et al., 2000; Lehtinen and Witter, 2014). They are found less often around impoundments, swales, and in riverine situations, at least in the South (Lichtenberg et al., 2006). They can also be found associated with agricultural crops (Brown, 1974; Anderson and Arruda, 2006), gravel pits (Schmidt and Necker, 1935), or formerly strip-­mined areas (Myers and Klimstra, 1963; Anderson and Arruda, 2006). In the arid High Plains, they are more confined to river bottoms than wetlands directly imbedded in the grasslands (see, for example, the distribution map in Lynch, 1985; Burdick and Swanson, 2010). Acris blanchardi is rarely found far from ­water, and they do not frequent temporary wetlands ­unless they are in proximity to permanent wetlands. ­These frogs prefer moist and muddy substrates without much bare ground in close proximity to shelter sites such as rocks (Smith et al., 2003; Burdick and Swanson, 2010). When along streams, they prefer pool to riffle habitats, and use both sunny and shaded microhabitats close to ­water; Burdick and Swanson (2010) found them usually within 13–44 cm from ­water. Blanchard’s Cricket Frogs have been reported at elevations as high as 1,275 m in New Mexico (Degenhardt et al., 1996). AQUATIC AND TERRESTRIAL ECOL­O GY

Activity normally occurs throughout the warmer months of the year, although activity extends year-­round in many areas if environmental conditions permit (e.g., Pyburn, 1958). This species is often active at cold temperatures when “normal” frogs are dormant, but they also are active at temperatures approaching 38°C (Clarke, 1958). When active, frogs prefer soils with high moisture content both day and night. When active at night, they show no preference for ground (i.e., coverboard) or canopy cover (Youngquist and Boone, 2014). Activity normally occurs well into the autumn. For example, A. blanchardi in Ohio ­were observed to be active from February to late March ­until early September (Wilcox, 1891) or mid-­November (Walker, 1946; Brenner, 1969). In Kansas and New Mexico, activity normally occurs into October or November (Smith, 1934; Heinrich and Kaufman, 1985; Collins, 1993; Degenhardt et al., 1996). In southern Illinois, cricket frogs have been observed in January (Gray, 1983). Other reports of winter activity are available for Kansas (Linsdale, 1927; Busby et al., 2005), Illinois (Ross-

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man, 1960; Smith, 1961; Tucker et al., 1995), Indiana (Blatchley, 1892; Evermann and Clark, 1916), and Ohio (Dickson, 2002). In Texas, activity tends to decline greatly in December and January, but the frogs never ­really dis­appear as long as temperatures permit. Overwintering occurs terrestrially in crayfish burrows, other small burrows, and cracks in the mud banks along ponds and rivers (Gray, 1971; Irwin et al., 1999; Swanson and Burdick, 2010; Badje et al., 2016), ­under surface debris (Garman, 1892; Walker, 1946, 1963; Pope, 1964; Bayless, 1966), among tree roots (Walker, 1946), ­under rocks near springs (Blair, 1951), or in gravel slopes along and above streams (McCallum and Trauth, 2003b). Overwintering may occur singularly or communally (Badje et al., 2016). Badje et al. (2016) reported a minimum mean hibernacula depth of 4.7 cm, with exposed south-­facing riverbanks as critical overwintering habitat. Movement to overwintering sites occurs by early October, and by mid-­November, all frogs have dis­appeared above ground as freezing temperatures set in. ­These frogs are capable of surviving for short periods in oxygenated ­water, but not in hypoxic conditions, and they even may survive short periods (90 m away (Pyburn, 1958; Burkett, 1984). Movements of ­these distances can occur over a period of a few days to months. Individuals seem to remain within an area for an extended period, with long-­distance movements back and forth between distant locations occurring periodically. Movements occur during periods of wet weather, and individuals move between adjacent ponds, pools, or other moist habitats. They even have been observed 60 m from the nearest ­water during rainfall. The greatest movement distance Pyburn (1958) recorded was 167 m,

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whereas Burkett (1984) recorded a single frog moving 213 m and Fitch (1958) recorded movements up to 183 m. ­These frogs ­will readily return to near their point of capture when displaced over distances >30 m. Despite most observers noting the adherence of this species to moist areas near ­water, ­there are discrepancies in the lit­er­a­ture. For example, Collins and Collins (2006) stated that “Northern Cricket Frogs evidently wander ­great distances from ­water during both dry and wet weather, and many apparently die during ­these wanderings, keeping populations at an optimal level.” It seems doubtful, however, that se­lection would operate so forcefully for the group as opposed to the individual. Fitch (1958) noted that most movements occurred along pond margins and moist ravines, and that frogs could move considerable distances ­under wet conditions. Many frogs likely die during dispersal, but not to keep populations “optimal.” Body length in this species is positively correlated with body mass, and the larger body size of A. blanchardi compared with A. crepitans and A. gryllus may be associated with the tendency of large frogs to lose ­water more slowly than small frogs (Nevo, 1973b). This is impor­tant, since much of the range of A. blanchardi is in the semiarid grasslands of the Plains and upper Midwest. The CTmax for this species is 38.8–39.5°C, and the thermal preference is 27–28°C (Gray, 1977). Blanchard’s Cricket Frogs likely are photopositive in their phototactic response as is the phyloge­ne­tically related A. crepitans (Jaeger and Hailman, 1973), suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities. They prob­ably are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds. Blanchard’s Cricket Frogs also likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males produce an advertisement call that both attracts females and provides information about the male’s position and calling status to rival males. Males tend to call from the same location on consecutive nights, and if displaced ­will return to the original calling location (Perrill and Shepherd, 1989). In addition to and in conjunction with calling, male frogs may extend their rear legs in a visual display directed ­toward one another (see Fig.1 in Horne et al., 2014). Frogs near each other in a chorus ­will circle within a small area while extending their rear legs and irregularly vocalizing. Occasionally frogs would attack one another by hopping on an opponent, then hopping off. Bouts ended when individuals ­stopped displaying. Circling bouts involved leg extensions,

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Foot-­flagging by male Acris blanchardi ­toward conspecific male. Kansas. Photo: Greg Sievert

direction changes, forward movements, and vocalization. Display bouts averaged 2.9 min. Calling occurs both by day and by night. In Arkansas, calling occurred at temperatures from 13 to 24°C (Briggler, 1998), whereas in Texas calls ­were heard at temperatures from 6 to 34°C. Wiest (1982), also in Texas, noted calling at 8.7–28.3°C over an 85 day period. Diurnal calling occurs at the lowest temperatures (1,000 ppb and for (24 hr) endrin is 23 ppb. However, the mean LC50 (96 hr) for toxaphene is only 76 ppb and for endrin is 10 ppb (Hall and Swineford, 1981). ­These results ­were for continuous flow toxicity tests, and survivorship was determined ­after 8 days.

Metals. In Northern Cricket Frog tadpoles, levels of aluminum, iron, magnesium, and manganese may be quite concentrated (Sparling and Lowe, 1996). Body concentra-

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STATUS AND CONSERVATION

Acris crepitans often is an abundant frog, although Weir et al. (2014) and Villena et al. (2016) suggested populations in the Northeast and the South ­were declining based on probability of occurrence through time. Populations at the northern extent of its range are vulnerable, however, and some populations have been extirpated. Northern Cricket Frogs dis­appeared from Long and Staten islands, New York, by the 1930s and 1970s, respectively (Gibbs et al., 2007). On Long Island, Overton (1914) characterized them as “extremely abundant and noisy in several marshes in the vicinity of Jamaica.” Only a few populations remain in New York on private lands. Other populations are threatened by agriculture and urbanization. In the western Piedmont of North Carolina, Rice et al. (2001) ­were only able to document A. crepitans at 1 site, although up to 8 sites had been documented historically prior to 1970. In terms of mitigation, Northern Cricket Frogs may use constructed ponds, but their abundance is not ­great at restored ponds. Merovich and Howard (2000) recorded only 10 A. crepitans using ponds >4–5 yrs old over a 2 yr monitoring period in Mary­land. No use was recorded in constructed ponds 19.0 mm SUL and females are 20–24.5 mm SUL (Alexander, 1966). Nevo (1973b) recorded the following mean male and female lengths in mm SUL: western Gulf Coastal Plain (22.8, 24.3), eastern Gulf Coastal Plain (22.2, 23.4), and southern Florida (22.1, 22.5). Males are 15–29 mm SUL and females 16–33 mm SUL in Georgia (Jensen et al., 2008). Adults are 16–22 mm SUL in Alabama (mean 19.7 mm) (Brown, 1956) and a mean of 21.1 mm SUL from another North Carolina study (Micancin and Mette, 2009). Larvae. The tadpoles are medium sized, light to dark gray, and have long tails with low tail fins. The body is slightly depressed, the nostrils are large, and the tail fins do not have bold markings, although they are finely flecked. Throats are dark. Many larvae have black tail tips, but a few do not, even within the same pond. The ­free part of the spiracle’s tube is longer in A. gryllus than in A. crepitans. Cricket frogs have the distinction of having the largest tadpole in relation to metamorph size of all North American anurans. A ­description of the tadpole, contrasting it with larval A. blanchardi, is in Orton (1947). Note that the larval description for A. gryllus in Wright (1929) is a composite that includes A. crepitans and A. blanchardi. Eggs. The eggs are small and deep brown to buffy olive above and cream to white below. The vitellus is 0.9– 1.43 mm in dia­meter (mean 1.16 in Alabama; Brown, 1956). ­There is a single, firm, gelatinous coating around the egg mea­sur­ing 2.4–3.6 mm in dia­meter and without an inner envelope according to Wright and Wright (1949). However, Brown (1956) noted 3 gelatinous membranes surrounding eggs deposited in captivity. The second envelope was firm but not gelatinous. Brown (1956) recorded the mean dia­meter of the inner envelope as 1.71 mm (range 1.62– 1.81), the second envelope as 1.96 mm (range 1.9–2.0), and the third envelope as 2.68 mm (range 2.57–2.95). DISTRIBUTION

Southern Cricket Frogs occur on the Coastal Plain from southeastern ­Virginia through the Florida Parishes of Louisiana. The range extends northward into the Coosa Valley of Alabama and in western Georgia northward through the Piedmont to the south end of the Ridge and Valley Province. Populations may extend elsewhere into the Piedmont along river floodplains. Isolated populations occur in southeastern Tennessee and in east central Georgia. In the vicinity of the Fall Line, the range of this species overlaps somewhat with A. crepitans, but perhaps not as much as statements in the older lit­er­a­ture indicate. The Mississippi

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Distribution of Acris gryllus. It is likely that A. gryllus crosses the border into east central Alabama.

River largely forms the western boundary of this species, but its presence in the Delta Region of Mississippi has been questioned (Boyd, 1964). The range extends northward to southwestern Tennessee, then eastward again south of the Fall Line from Alabama back though southeastern ­Virginia. Southern Cricket Frogs may be found on larger islands, such as Roanoke and Bodie islands, North Carolina (Braswell, 1988; Gaul and Mitchell, 2007), and Isle of Hope, St. Simons and Cumberland islands, Georgia (Williamson and Moulis, 1994; Shoop and Ruckdeschel, 2006; O’Hare and Madden, 2018). Impor­tant distributional references include: Alabama (Brown, 1956; Mount, 1975), Florida (Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Louisiana (Viosca, 1944; Dundee and Rossman, 1989; Boundy and Carr, 2017), Mississippi (Boyd, 1964), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007; Micancin and Mette, 2009), South Carolina (Dodd and Barichivich, 2017; Fields, 2019), Tennessee (Niemiller and Reynolds, 2011), and ­Virginia (Tobey, 1985; Mitchell and Reay, 1999; Micancin et al., 2012). Note that the maps in Williamson and Moulis (1994) pre­sent a confused distribution pattern between A. crepitans and A. gryllus in Georgia. Many of the dot locations likely are based on misidentified individuals (see Jensen et al., 2008). Micancin et al. (2012) also noted that many Acris from southeastern ­Virginia in museum collections ­were misidentified.

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FOSSIL REC­O RD

Pleistocene Rancholabrean fossils of A. gryllus have been reported from north central Florida (Holman, 2003). A Miocene Acris was also described from northern Florida as A. barbouri, based on differences in the shape of the ilium (Holman, 1967). SYSTEMATICS AND GEOGRAPHIC VARIATION

At one time, all cricket frogs in North Amer­i­ca ­were thought to be part of a single wide-­ranging species. Comparisons of blood and liver proteins (Dessauer and Nevo, 1969), molecular data (­Gamble et al., 2008), call structure (Nevo and Capranica, 1985), chromosomes (Bushnell et al., 1939), distributional patterns (Neill, 1954), ecol­ogy, and morphology have conclusively demonstrated that A. gryllus is a distinct species. ­There are 2 major clades within A. gryllus, an eastern clade occurring from the Florida Panhandle to southeastern ­Virginia, and a western clade found in Mississippi and adjacent Tennessee (­Gamble et al., 2008). The boundary between ­these clades is undetermined but is likely in the Mobile Basin and Tombigbee River system. The distinction between ­these clades does not follow previous subspecific nomenclature (A. g. gryllus throughout most of the range and A. g. dorsalis in Florida). Despite some morphological differences (Neill, 1950b), the recognition of A. g. dorsalis is not supported by molecular data. Laboratory experiments indicate a high degree of ge­ne­tic compatibility between A. crepitans and A. gryllus, regardless of which species is male or female (Mecham, 1964). However, few larvae could be reared through metamorphosis, and none released into the environment ­were subsequently recovered by Mecham (1964). Backcrossed hybrids had low fertilization rates and poor development, and the hybrids appeared stunted. Hybridization does not appear to occur in nature, perhaps ­because of differences in mating calls and ­limited geographic overlap. However, ­there is at least 1 observation of interspecific aggression between males of ­these species at a breeding pond (Micancin and Mette, 2010). Pattern polymorphism is common in cricket frogs and may reflect dif­fer­ent se­lection pressures on dif­fer­ent seasonally changing backgrounds (see discussion in Acris crepitans account). In Southern Cricket Frogs, Nevo (1973a) found that 24–100% of frogs ­were gray, 22.6–62.5% had a red stripe, and 9–30% had a green stripe at the 9 populations sampled. ADULT HABITAT

Acris gryllus is a wide-­ranging species occupying many dif­fer­ent habitats throughout the southeastern United States. Primarily a lowland species, it prefers well-­drained habitats

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in the vicinity of permanent or temporary ponds, wet prairies, seepage bogs, sloughs, canals, sawgrass prairies, bayheads, lakes, swamps, and grassy meadows. Examples of surrounding habitats include longleaf pine–­turkey oak sandhills, mixed-­pine–­oak woodlands, hardwood forest, mesic and xeric hammocks, pine flatwoods, subtropical tree islands, agricultural fields, and silvicultural operations (Anderson et al., 1952; Harima, 1969; Dodd, 1992; Enge, 1998a, 2002; Enge and Wood, 1998; Meshaka et al., 2000; Baber et al., 2005; Greenberg and Tanner, 2005b; Means and Franz, 2005; Surdick, 2005; Dodd and Barichivich, 2007; Meshaka and Layne, 2015; Chandler et al., 2015a; Erwin et al., 2016). The species ventures into the uplands in many areas, marking the division between the lowland Coastal Plain and the more upland Piedmont provinces. Southern Cricket Frogs are abundant along the shorelines of ponds and wetlands, especially where ­there is an abundance of low-­growing grasses such as Eleocharis (Bancroft et al., 1983). They readily are found on mats of floating vegetation. They respond favorably to prescribed fire management in the southeastern longleaf pine ecosystem (Klaus and Noss, 2016). AQUATIC AND TERRESTRIAL ECOL­O GY

The first field notes on this ­little terrestrial frog ­were recorded by Bartram (1791:278) as he traveled in the southeastern United States, although he may have mixed in observations of other species as he characterized its habits. For example, A. gryllus does not climb on vegetation or trees, although Bartram certainly got the sound of the call correct (“a feeble note sounding like crickets”). Throughout the year, ­these frogs congregate in considerable numbers around the margins of ponds and lakes and in grassy wetlands in savannas. During the nonbreeding season, they are frequently found in nearby lowland cypress swamps where they occupy the margins of the swamp. They normally do not occur very far from ­water and are among the most aquatic of native anurans with the exception of the ranids (Lithobates, Rana). They frequently occur on emergent and floating vegetation and along wet trails in prairie, swampy habitats, and along streams in steephead ravines (Wright, 1932; Enge, 1998b). Indeed, Wright (1932) characterized them as being found “in almost ­every type of plant habitat which has any moisture at all.” When in wetlands, they do not always remain close to shore. For example, Beck (1948) frequently found them on floating vegetation 91 m from shore. Southern Cricket Frogs are active year-­round, weather permitting. They tolerate colder weather better than most other southern frogs. In laboratory experiments, the mean CTmin is ca. 3°C and the mean CTmax is ca. 38°C

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(John-­Alder et al., 1988). Southern Cricket Frogs are active at ­every temperature in between ­these values. ­There is no relationship between winter activity and canopy cover or bank slope, but A. gryllus avoids areas inhabited by Lithobates grylio, presumably ­because of predator avoidance (Bomske and Bickford, 2019). Winter occurrence is positively associated with plant species richness. When at the breeding site or sitting along a pond shoreline, Southern Cricket Frogs move very ­little from 1 part of the pond or wetland to another. In general, they are philopatric with a small home range, preferring to remain within 5 m of the site of previous occurrence. Bayless (1966) found the mean movement distance to be 1.3 m (range 0.5–3 m). Some frogs do make extensive movements over the course of an activity season, however. For example, 1 frog in Louisiana moved ca. 33 m in ca. 35 days, and the greatest distance Bayless (1966) recorded movement was ca. 65 m over a period of 180 days between captures. In Florida, Telford (1952) encountered them often at distances of 91 m or more from the nearest ­water, and Carr (1940a) recorded them 274 m from ­water. Also in Florida, Dodd (1996) recorded A. gryllus 255–492 m (mean 383 m) from the nearest ­water in sandhills habitat; movements tended to be uniform in direction to and away from a temporary pond in any 1 year, but over a 5 yr period, the immigration and emigration pattern was nonuniform as frogs moved through the habitat during an extended drought (Dodd and Anderson, 2018). In North Carolina, Micancin (2010) found a single female 562 m from the nearest breeding chorus. The extent to which A. gryllus uses terrestrial habitats needs further investigation. Like Northern Cricket Frogs, Southern Cricket Frogs are able to use Y-­axis celestial orientation to determine dispersal and migratory directions to and away from a perpendicular shoreline or within a pond. This ability requires a learned shore position, a view of a celestial cue (sun, moon, and stars), and an internal clock phased to local time. Orientation is most effective by day when the sun is vis­i­ble and by night when the moon is vis­i­ble (Ferguson et al., 1965; Ferguson, 1966a). If only the stars are vis­i­ble, frogs show a bipolar orientation. Displaced frogs are able to return to the shoreline where they ­were captured or to the point of release using the position of the sun for orientation. Ferguson (1963) reported that displaced frogs easily returned 91 m to the point of capture, and that many frogs returned as far as 223 m, although sometimes stopping at intervening ponds along the way. Celestial orientation can be mediated through the eyes, but extraoptic light receptors in the brain allow proper orientation and setting of biological clocks even in blind A. gryllus (Taylor and Ferguson, 1970).

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Southern Cricket Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). Southern Cricket Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Male Southern Cricket Frogs call both by day and by night, but the majority of calls are made between 19:30 and 02:00 (Mohr and Dorcas, 1999). Calling extends throughout the year in Southern Cricket Frogs, but nearly all successful reproduction takes place from late spring throughout the summer. For example, calling begins in April and extends ­until August in North Carolina (Alexander, 1966) and Mississippi (Smith, 1975), and from February to October in Georgia (Jensen et al., 2008). In Florida, calls may be heard year-­round depending on location and weather conditions, although Bancroft et al. (1983) and Meshaka and Layne (2015) only reported calls from February to October in central and southern Florida. In most locations, calling likely precedes ­actual reproduction by several weeks as it does in other cricket frogs. The reason for extensive calling late in the season is unknown. As with other cricket frogs, the advertisement call sounds like a series of clicks or “ticks,” which are composed of pulses repeated at intervals of 0.2 sec (Blair, 1958a). The dominant frequency of the call is a mean of 3.55 kHz. Some characteristics of the call include: a click rate of 1.2–6.21 (mean 2) per entire call, a mean click duration of 34–36 ms, a mean number of pulses of 10.2–10.9, a call duration of 12–28 sec (mean 16.5 sec), a total number of clicks of 13–40 (mean 27.8), and an average click rate of 1.4–3.8. Interpulse intervals average 1.7–2.1 sec. The number of clicks increases with the duration of the call. Nevo and Capranica (1985) and Micancin and Mette (2009) provided extensive information on regional variation in both gross and fine structure of the call of A. gryllus. Males call from along the shoreline of a pond or lake and, like A. blanchardi, tend to defend an acoustic calling space. In Georgia, calling males occupied a territory of only 0.56 m2 (range 0.028–1.362 m2) (Forester and Daniel, 1986). The size of the territory is not correlated with the number of nights a male is pre­sent or with mating success. Males remain within a par­tic­u­lar area from 1 night to the next, and movements are very small. Forester and Daniel (1986) recorded mean movements of only 52 cm (range 0–205 cm).

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Females move ­toward a calling male. She ­will turn her head from side to side in the direction of the caller as if this assists her in orientation, and she ­will perform a series of circling hops if she has difficulty in locating his position. The female does not initiate contact. In time, the male usually observes the female, and then moves ­toward and amplexes her. Forester and Daniel (1986) described such a courtship sequence that included 8 female circles over a 5 min period. A male may successfully amplex more than 1 female (Forester and Daniel, 1986). BREEDING SITES

Southern Cricket Frogs breed in a wide variety of temporary to permanent wetlands, including small grassy meadow wetlands, depression marshes, cypress savannas, cypress-­gum ponds, woodland ponds, and the quiet shallows of very large lakes (e.g., Liner et al., 2008). They tend to prefer shorelines with minimal or no vegetation when breeding along pond or lake margins. Fish may or may not be pre­sent. Acris gryllus also breeds in human-­ created wetlands, such as ­those that form ­under power-­line rights-­of-­way, in roadside ditches, and along trails through wet areas. As long as seasonal wetlands are available, they ­will breed on golf courses (Scott et al., 2008). Breeding sites often have extensive mats of floating vegetation or lily pads on which frogs sit and call. The vegetation also provides concealment from predators and from direct solar insolation. A generalist approach to breeding site choice may allow this species to adapt to extreme environmental changes across a landscape. For example, Babbitt and Tanner (2000) studied breeding at a site in south Florida that experienced a drought 1 year followed by flooding the next. The flooding allowed fish to invade formerly fishless wetlands. Acris gryllus was able to use temporary isolated ponds during the first year’s drought as well as the extensive wetlands resulting from the flooding during the second year. Reproductive output was reduced during the drought as some ponds dried, but it was successful both years despite the extremes in precipitation. REPRODUCTION

Southern Cricket Frogs normally breed throughout the spring to early summer, with the exact timing depending on environmental conditions. Calling occurs throughout the warmer period of the year and extends well into the autumn or even winter, but the extent of successful reproduction ­after the primary calling period is unknown. Carr (1940a) stated that they bred ­every month of the year in Florida. In Louisiana, however, most reproduction occurs in March and

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April, although calling may continue ­until September. Bayless (1966) found no clasping pairs ­after 8 April and considered this species possibly to be an explosive breeder whose breeding activity was triggered by heavy rainfall. In Georgia, calling occurs from February to October, but most breeding occurs from April to June (Jensen et al., 2008). As noted above, it is unlikely that midsummer to autumn calling results in amplexus and reproduction except perhaps well south in the Florida peninsula. Testis mass and male fat body mass are lowest in winter and highest in spring, and testis mass decreases in summer and increases in autumn. Ovarian mass is correlated with both body mass and snout-­ urostyle length. In contrast, male fat body mass was correlated with body mass but not snout-­urostyle length. Female fat body mass reaches its maximum in winter and is used to yolk developing eggs in the coming spring. ­These ­factors, together with the timing of spermatogenesis and ovarian cycles, do not lend themselves to successful reproduction ­after the spring to early summer breeding season that exists throughout much of the species’ range (Smith, 1975). In central and south Florida, however, some reproduction likely occurs year-­round, even if greatly decreased during the winter. Tadpoles of A. gryllus are pre­sent year-­round at Loxahatchee National Wildlife Refuge (Baber et al., 2005) and at the MacArthur Agro-­Ecology Research Center (Babbitt and Tanner, 2000). Tadpole densities ranged from 0.06 to 0.43 m2, but varied annually and monthly. The location at which breeding grades from occurring solely in spring and summer to occurring year-­round in Florida has not been determined. Eggs are deposited singly and sometimes in small groups of 3–4. The eggs may stick together in a loose mass of 7–10 eggs per mass. ­These masses cling to vegetation via the gelatinous envelopes, or eggs may rest freely upon the substrate in shallow ­water. ­Actual counts of egg clutches are few in the lit­er­a­ture. Females oviposited 99–156 eggs per clutch in North Carolina (Alexander, 1966), and Wright (1932) reported a single clutch count of 241 eggs. Many authors state that clutch size can reach 250, perhaps referring to Wright’s count. Mitchell and Pague (2014) recorded 109–340 eggs (mean 170) from ­Virginia. Hatching occurs quickly, within 4 days of deposition. LARVAL ECOL­O GY

The larval period lasts from 32 to 94 days (Wright, 1932; Bayless, 1966; Jensen et al., 2008). Wright (1932) recorded a mean of 67 days for the larval period, but stated that 50–90 days seemed the “probable” period. Metamorphs are 9–15 mm SUL (Wright, 1932; Alexander, 1966; Jensen et al., 2008).

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Larval Acris gryllus. Some larvae have the black tail spot, whereas o ­ thers do not. Photo: C.K. Dodd, Jr.

DIET

The diet of Southern Cricket Frogs includes small invertebrates, especially ants. In a Florida study, 70% of the stomachs contained ants, and they made up 68% of the prey ingested. Other prey included mosquitoes, leaf bugs, spiders, chironomid flies, and beetles (Franz, 1972). In the latter study, frogs captured between 22:00 and 07:00 had no food in their stomachs, suggesting solely diurnal feeding. In Louisiana, specific items include collembolans, ants, flies and fly larvae, spiders, mites, beetles, true bugs, aphids, termites, and miscellaneous larval and adult insects (Beck, 1948; Bayless, 1969b). Carr (1940a) recorded them feeding on emerging midges, mayflies, and chironomids. He further noted that they sometimes attacked prey too large to be eaten. In areas of sympatry, the food items of A. crepitans and A. gryllus overlap, showing no prey specialization. Feeding occurs year-­round when weather conditions permit. PREDATION AND DEFENSE

Larval A. gryllus appear to be palatable, and even in the presence of chemicals from potential fish predators do not spend more time in refugia than they do when fish are absent (Kats et al., 1988). They may, however, reduce their activity in the presence of dragonfly naiads, fish, or Eastern Red-­ spotted Newts (Richardson, 2001). As with A. blanchardi, many have black tail tips, which serve as disruptive coloration or help to distract a potential predator away from the head and body. Caldwell (1982) demonstrated that larvae with black-­tipped tails ­were found most often in ponds, whereas ­those without this coloration ­were found more frequently in lakes and creeks. In ponds, dragonfly larvae are the primary predator. The black tail tip draws their attention, and larvae in wetlands containing Anax have much higher frequencies of black-­tipped tails than in their absence. In creeks and lakes, the uniform body coloration may help in concealment from predaceous fishes (Caldwell, 1982). This polymorphism is pre­sent even in nearby populations and is thus predator mediated but habitat specific. The Southern Cricket Frog jumps readily at the approach of an intruder, and makes sudden, rapid leaps. The first leap

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is usually the longest (mean 72.7 cm: Blem et al., 1978). Jumping distances are correlated with temperature, with jumps of 20 cm at 5°C to nearly 1 m at 35°C (John-­Alder et al., 1988). A frog ­will jump ­toward the ­water and swim rapidly among vegetation, circling back ­toward shore. If disturbed again, it may dive and bury itself among vegetation or mud, but it often pops back up rather rapidly. During jumping, the frog is able to quickly change directions, making pursuit difficult. Dodd (1991) noted that they ­were capable of easily crossing a drift fence by simply jumping on it and walking the remaining way to the top, at least in laboratory experiments; ­there appeared to be less tendency to trespass the fence in the field. Predators undoubtedly include a variety of invertebrates, ranid frogs, snakes (Thamnophis sp.), birds (particularly wading birds, Florida grackles), and mammals (pigs). Few specific observations are available, most notably for garter and ribbon snakes (T. sirtalis, T. sauritus) (Wright, 1932). Goin (1943) reported a large Dolomedes spider eating an adult cricket frog. POPULATION BIOLOGY

Southern Cricket Frogs are among the most abundant frogs in their environment, where a short walk along a lake or pond shore results in dozens of ­these tiny frogs hopping in ­every direction. Wright (1932) called it the most abundant frog in the Okefenokee Swamp, and it is certainly one of the most abundant frogs around ponds and lakes in north central Florida. Distribution, however, is somewhat nonrandom in the vicinity of more contiguous shallow grassy pools and associated terrestrial habitat. Robust population estimates or even empirical counts are few in the lit­er­a­ture. In a Louisiana population, Turner (1960a) found that individuals ­were spaced at intervals ranging from 1.7 to 1.9 m from their nearest neighbor in and around a grassy wetland, and that densities changed with time. He estimated that ­there ­were 0.066 frogs per m2 for a total population of about 90 in the 1,350 m2 wetland in December. By April, the density increased to 0.085 frogs per m2 with a population estimate of about 140. In Florida, Greenberg and Tanner (2005b) captured 888 adults and 297 juveniles around 8 isolated temporary ponds surrounded by dry sandhills over a 7 yr period. Also in Florida, Dodd (1992) captured 255 A. gryllus over a 5+ yr study at a temporary pond surrounded by xeric hammock and sandhills. Gibbons and Bennett (1974) captured 340 A. gryllus around 2 ponds in 2 yrs; most frogs, however, ­were captured at 1 of the ponds in only 1 year. Also in South Carolina, Russell et al. (2002a) documented 1,298 A. gryllus using 5 small, isolated wetlands over a 2 yr period; tadpoles also ­were observed in the wetlands.

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Acris gryllus 219

Adult Acris gryllus, green phase. Photo: Alan Cressler

Both A. crepitans and A. blanchardi are essentially annual species with very few individuals surviving beyond the first breeding season. Some authors also state that A. gryllus reaches maturity quickly and is ready to breed in the spring following metamorphosis (e.g., Jensen et al., 2008). However, this may not be the case with A. gryllus, as longevity was >3 years in the North Carolina population studied by Alexander (1966). He found that 50–70% of females failed to reach sexual maturity 1 year ­after metamorphosis, and consequently first bred the second year. Thus, the normal life-­span may be greater in Southern Cricket Frogs than in the other 2 species of Acris. As with many frogs, the number of males is greater at breeding ponds than females, but overall population sex ratios remain poorly documented. In Georgia, Forester and Daniel (1986) reported an operational sex ratio of 5.6 males per 0.3 females, but acknowledged that this represented an underestimation of the number of females in the population. COMMUNITY ECOL­O GY

In many areas of the South, populations of A. gryllus and A. crepitans overlap geo­graph­i­cally (Viosca, 1944; Neill, 1954; Mecham, 1964; Bayless, 1969b; Micancin and Mette, 2009). However, it appears ­these species have dif­fer­ent habitat preferences where they occur together. For example, A. crepitans is found in bottomland swamps or ­water close to the shore of lakes, whereas A. gryllus occurs more in uplands or along drainage ditches, where the species overlap (Norton and Harvey, 1975; Dundee and Rossman, 1989). Acris gryllus tends to avoid shrub-­dominated pond margins whereas A. crepitans prefers such areas; although A. gryllus ­will breed in ponds, it prefers open shorelines. Acris gryllus also prefers to

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breed in shallow grassy meadow pools whereas A. crepitans does not. Neither species breeds in lowland swamps, but both species may be found si­mul­ta­neously year-­round in such habitats, especially during the nonbreeding season. According to Bayless (1969b), A. gryllus tends to occur along the swamp margins, with A. crepitans occurring in the muddy bottoms. ­These differences may help segregate the species, at least during the breeding season, although other authors (e.g., Mecham, 1964) could find no consistent differences in ecological separation between the species. Despite dif­fer­ent preferences for microhabitats, however, both species may occur at a pond. Bayless (1966) found that abundance was about equal between species in the winter, but that during the warm months of the year, A. crepitans far exceeded A. gryllus in abundance. Newly metamorphosed young tend to swell in abundance in midsummer, but ­these frogs disperse soon thereafter. In addition, A. crepitans tends to occupy the more shaded (or less exposed) portions of the pond, whereas A. gryllus occupies the more exposed sections (Bayless, 1966). This suggests the potential for observer bias when assessing abundance through visual encounter surveys. In addition to habitat differences, ­there are differences in the advertisement calls of the species. The calls sound like a series of clicks, and the dominant frequencies of the calls overlap. However, ­there are differences in the temporal structure and amplitude of the clicks (Micancin and Mette, 2009). Females are able to recognize the advertisement calls of conspecific males (Micancin, 2008). DISEASES, PARASITES, AND MALFORMATIONS

The fungal pathogen Basidiobolus sp. has been reported from Southern Cricket Frogs from Florida (Okafor et al., 1984). As of 2008, the amphibian chytrid fungus had not been detected in A. gryllus in the Southeast (Rothermel et al., 2008). However, it is now known from Alabama, Florida, Georgia, and North Carolina (Rizkalla, 2010; Hill and Levy, 2014; Hughey et al., 2014; Chiari et al., 2017; Lentz et al., 2021). Ranavirus has been reported from North Carolina (Lentz et al., 2021). SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. In Louisiana, Bayless (1966) routinely found A. gryllus in ­water with a pH of 4–5. In the related A. crepitans, the toxic effects of pH are evident at pH values 1 km away. Males call from vegetation within and surrounding acid seeps and bogs. Most perch heights are located 1–1.5 m off the ground, rarely above 2 m (Bullard, 1965; Means and Longden, 1976). Males leave their arboreal calling perches well ­after dark and move to the margins of slow-­moving streams and seepage bogs. ­There, they begin calling from the substrate near an oviposition site. They frequently call in duets, with the second male calling immediately ­after the first. The call has a ventriloquist quality that makes it difficult for a ­human observer to locate the male. Calling begins shortly ­after sunset with most calling occurring 2–3 hrs ­after sunset. Occasional males may be heard around daybreak. A female moves ­toward a calling male and makes contact by striking him 1 or more times. This usually occurs on the ground, although initial contact may be made on a calling perch since amplexed pairs have been observed in trees. Amplexus is axillary, with the male riding on her back with his toes resting on the top of her thighs. The female initiates oviposition by bowing her back (head and posterior up, abdomen down) and thrusting her legs laterally. This movement places her vent in close proximity to that of the male’s vent. The male moves his legs to her sides in a stroking manner, whereupon she straightens her back and 4–9 unattached single eggs are extruded suddenly and fertilized (Noble and Noble, 1923; Means and Longden, 1976). Noble and Noble (1923) illustrated a mating pair. Males appear to be evenly spaced about 5–10 m apart. Calling D. andersonii males are territorial and have an aggressive encounter call if an intruder approaches too closely. The encounter call is similar to the advertisement call but has an increased repetition rate made by shortening the interval between low frequency peaks, much as D. squirellus does (Fellers, 1979a).

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BREEDING SITES

Breeding occurs in acidic hillside seepage bogs and shrub bogs, although the plant composition varies among the disjunct populations. ­Water should be clear, shallow, and slow moving; stagnant ­waters are avoided. Breeding sites in North Carolina are mostly temporary to semipermanent ditches, streams, and pools. REPRODUCTION

Breeding occurs in shallow ­water, with eggs lying ­free on the substrate and unconnected to one another. Clutch size is 800–1,000 in New Jersey (Noble and Noble, 1923), and Means and Longden (1976) reported a female ovipositing 206 eggs in Florida. Hatching occurs within 3 days. LARVAL ECOL­O GY

Larvae prefer shallow w ­ ater where they largely remain immobile during daylight hours. I experimental ­trials, they reduced their activity patterns when exposed to predators (Notophthalmus viridescens, Enneacanthus obesus, Pantala) and assumed a more benthic position in the ­water column than they other­wise would have (Lawler, 1989). The larval period is estimated to be 49–75 days (Wright, 1932; Gosner and Black, 1957b). Newly metamorphosed froglets are 11–15 mm SUL.

Breeding habitat of D. andersonii. Photo: John Bunnell

Tadpole of D. andersonii. Photo: John Bunnell

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224  Hylidae

DIET

The adult diet consists of beetles, grasshoppers, ants, flies, and larval insects (Noble and Noble, 1923). PREDATION AND DEFENSE

When handled, D. andersonii emits an odor of “raw peas,” which could have an antipredator function (Noble and Noble, 1923). The green coloration and stripe of postmetamorphs suggest crypsis with surrounding vegetation. The stripe on the tail of the tadpole may function in counter-­ shading, which aids in crypsis. POPULATION BIOLOGY

Growth of postmetamorphs is rapid, and juveniles can reach breeding size by the first summer following metamorphosis, especially if hatched from eggs oviposited early in the season. It seems likely, however, that juveniles hatched from eggs oviposited late in the season might not actually breed ­until the spring of their second year. Larvae of dif­fer­ent sizes can be found throughout the summer, indicating a prolonged breeding season. Estimates of effective population sizes (Ne) have been derived for New Jersey (4,241), North Carolina (403,718), South Carolina (72,504) and Florida/Alabama (22,014) (Oswald et al., 2020). As expected, populations in North Carolina and South Carolina have the highest migration rates between populations, but even ­there the rates are quite low (3.8 is easily tolerated by D. andersonii, with the critical pH from 3.6 to 3.8 (Gosner and Black, 1957a). Successful hatching can occur at a pH of 3.7 (Freda and Dunson, 1986). The estimated lethal pH is just over 3.4. Still, larval D. andersonii grow significantly slower at a low pH (40% slower at 3.75 and 25% at 4.0) than they do at higher pH (Freda and Dunson, 1986). The pH of most natu­ral ponds is >4.0.

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STATUS AND CONSERVATION

The precarious status and need for conservation management of D. andersonii has long been recognized (Bury et al., 1980). Pine Barrens Treefrog habitat may be threatened by the encroachment of woody canopy vegetation, especially in areas where fires have been excluded. Habitat degradation, especially outside of protected areas, is always a threat. In managed habitats, this species may persist even in some areas of commercial forestry and along energy corridors where shrubby vegetation is maintained. The use of herbicides to control vegetation, however, is not advisable ­until the lethal and sublethal effects on eggs and larval development are understood. In surveys from 2013 to 2016, Moler et al. (2020) detected the species at 49 of 111 historical sites in the Florida Panhandle, many of which ­were ­either in Blackwater State Forest or on Eglin Air Force Base. They further identified 33 new sites on public land and 4 new sites on private land. Frogs ­were not detected at historical sites in Holmes County, however, although ­there ­were only 4 populations known from this county. Detection probability

Dryophytes arenicolor (Cope, 1866) Canyon Treefrog ETYMOLOGY

arenicolor: from the Latin arena meaning ‘sand’ and color meaning ‘color.’ The name refers to the frog’s ground color. NOMENCLATURE

McGinnis and Stebbins (2018): Hyla arenicolor Fouquette and Dubois (2014): Hyla (Dryophytes) arenicolor Synonyms: Hyla affinis, Hyla coper, Hyla copii, Hyliola digueti In the early lit­er­a­ture (e.g., Storer, 1925; Slevin, 1928; Wright and Wright, 1949; Stebbins, 1951), accounts of D. arenicolor may have information on Hyliola cadaverina intermingled with it. This has often resulted in incorrect range delineation and confused descriptions, such as statements that D. arenicolor is sometimes greenish. IDENTIFICATION

Adults. The ground color is light gray to dark brown, depending upon substrate and temperature, with a rugose dorsal surface. Individuals may exhibit a lichen-­like pattern that allows them to blend in well with their background

Dodd_Canada_int_5pgs_B1&B2.indd 225

was ca. 43%, with an estimated 63% site occupancy. ­These authors recommended at least 5 site visits to infer species absence via call surveys. They further considered the Florida population to be stable in terms of status and distribution. The amount of available wetland habitat declined between 2001 and 2016 in all 4 populations, to 2,695 km2 in Florida, 2,482 km2 in South Carolina, 7,384 km2 in North Carolina, and 3,238 km2 in New Jersey as of 2016 (Oswald et al., 2020). Total herbaceous wetlands, the habitat preferred by this species, comprise only 130 km2 in Florida, 73 km2 in South Carolina, 334 km2 in North Carolina, and 848 km2 in New Jersey as of 2016. Indeed, Oswald et al. (2020) noted that ­these ­were only coarse county-­wide habitat estimates that likely overestimate the extent of available or occupied habitat. Clearly, ­these discrete populations should be managed for their unique ge­ne­tic and phenotypic diversity, and efforts should be made to preserve habitats not already ­under management agreements. This species is considered Threatened in New Jersey and a Protected Nongame Species in Alabama. Take is prohibited in Florida via Florida Administrative Rule 68A-26.002.

and, like many other treefrogs, they readily change their color and pattern to match their substrate. A dark-­edged white spot is pre­sent ­under the eye, and ­there is a fold of skin above the tympanum. The skin is granular with many small tubercles. The toe tips are expanded, allowing them to move on slick canyon walls. The fin­gers are not webbed, but the toes are partially webbed. Venters are creamy white, with bright yellow or orange on the undersides of the hind legs; venters are closely granular. The maximum size is 57 mm SUL (McGinnis and Stebbins, 2018). Wright and Wright (1949) gave the male SUL as 29–53 mm and the female as 30–54 mm; Goldberg (2020e) reported males averaged 39.9 mm SUL (range 28–48 mm) and females 42.3 mm SUL (range 35–50 mm) based on museum specimens. Larvae. Newly hatched larvae are yellow brown dorsally and yellow ventrally. Mature tadpoles are dark brown to golden brown and somewhat globular in shape with abdomens that appear black; the area anterior to the abdomen is unpigmented. When viewed dorsally, the tail musculature does not have regularly spaced transverse light bars or ­saddles. Dorsal tail fins have slight reticulations, but ventral fins are unpigmented; the extent of tail fin pigmentation increases with development, especially as the rear legs appear. In addition, ­there appears to be regional variation in tadpole pigmentation, with Texas individuals having tail fins that are much more heavi­ly blotched than more western

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226  Hylidae

Hind foot of Dryophytes arenicolor. Illustration by Breck Bartholomew.

(1999), Dixon (2000, 2013), Brennan and Holycross (2006), Tipton et al. (2012), Davis and LaDuc (2018), Murphy (2019), and Holycross et al. (2021). Lateral head view of Dryophytes arenicolor. Illustration by Breck Bartholomew.

populations. ­There also appears to be regional variation in tadpole size, with tadpoles from the Chiricahuas smaller (to 38.3 mm TL) than ­those from along the Virgin River (to 47.3 mm TL) (Zweifel, 1961). Minton (1958) mentions tadpoles 35–40 mm TL and Wright (1929) noted tadpoles to 50 mm TL, both from Texas populations. Albino larvae and a recently metamorphosed juvenile w ­ ere reported from Texas (Van Devender, 1969). Zweifel (1961) provided figures of larvae at vari­ous stages of development and diagrams of the mouthparts. Eggs. The vitellus is 1.8–2.4 mm in dia­meter (mean 2.07 mm) with a single envelope 3.9–5 mm in dia­meter (mean 4.4 mm) (Livezey and Wright, 1947). DISTRIBUTION

Canyon Treefrogs are found from southern Utah southward into México as far as northern Oaxaca. Isolated populations occur in southeastern Colorado, eastern and southeastern New Mexico, and in west Texas (Chisos, Davis, Del Norte, and Sierra Vieja mountains). The species occurs in many deep canyons, including the ­Grand Canyon (Eaton, 1935; personal observation). Although it occurs on the Arizona side of the Colorado River, ­there are no rec­ords from California. Impor­tant distributional references include: Van Denburgh (1924), Degenhardt et al. (1996), Hammerson

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FOSSIL REC­O RD

Pleistocene fossils of D. arenicolor are known from New Mexico and Sonora, México (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

Dryophytes arenicolor is a member of the D. eximius species group, a primarily tropical assemblage from México. Barber (1999) identified 3 divergent lineages within D. arenicolor, based on analyses of mtDNA. ­These areas corresponded to populations in Texas, New Mexico, eastern and central Arizona, southern Utah, and the Chiricahua Mountains (clade 1), populations in sky islands of the Sonoran Desert and adjacent Mexican Highlands (clade 2), and populations in the ­Grand Canyon (clade 3). Clade 2 has 2 subgroups, 1 in the Pinalẽno, Ricon, and Catalina mountains and Arivaipa Creek, and the other in the Santa Rita, Huachuca, and Patagonia mountains. Clade 3 was highly divergent from clades 1 and 2. Indeed, the level of divergence among the 3 clades suggested they might represent dif­fer­ent species. In contrast, Klymus et al. (2010) reexamined the phylogeography of D. arenicolor using nuclear DNA and call analyses. They concluded that nuclear data and variation in call patterns ­were more congruent with a 2-­clade scenario. ­These latter authors, as well as Bryson et al. (2010, 2014), found evidence of past mtDNA introgression with D. wrightorum in Barber’s (1999) ­Grand Canyon populations, even though D. wrightorum no longer occurs in the area. In a further analy­sis including many

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Dryophytes arenicolor 227

canyons often have cottonwood trees and other plants frequently associated with desert riparian areas. Adjacent habitats include pinyon-­juniper woodland, pine-­oak associations, and semiarid mesquite grasslands (Aitchison and Tomko, 1974). Aitchison and Tomko (1974) observed them at elevations of 1,676–1,798 m near Flagstaff, and Degenhardt et al. (1996) reported them at 1,200–2,500 m in New Mexico. The species occurs from near sea level to 3,048 m (Stebbins, 2003). TERRESTRIAL ECOL­O GY

Distribution of Dryophytes arenicolor

more Mexican populations, Bryson et al. (2014) concluded ­there ­were actually 5 separate clades within D. arenicolor. Gene capture and mitochondrial introgression are common within the D. eximius species group, leading to a complex evolutionary history and phylogeography (Bryson et al., 2010, 2014). ­Under laboratory conditions, crosses of D. arenicolor with Smilisca baudinii or Hyliola cadaverina, H. regilla, Pseudacris clarkii, Dryophytes cinereus, or D. versicolor ­were generally unsuccessful, and no metamorphs ­were produced; crosses with D. femoralis, D. squirellus, or D. chrysoscelis produced some metamorphs, but backcrosses ­were unsuccessful (Pierce, 1975).

Dryophytes arenicolor usually are found perched on boulders and cliffs above pools of ­water. Activity occurs from February to November depending upon location and weather conditions (Gates, 1957; Swann, 2005; Lazaroff et al., 2006). Canyon Treefrogs are largely nocturnal but may become active a few hours before dusk as canyon shading deepens. They also bask in the direct sun at temperatures of up to 35°C (Wylie, 1981). For example, Jameson and Flury (1949) recorded them sitting in the direct sun on boulders and solid rock stream banks. Wylie (1981) suggested that sitting on exposed rock ­faces allows them to avoid certain predators, such as Garter Snakes (Thamnophis). During very dry weather, they may aggregate in moist situations. Swann (2005) mentions up to 100 frogs on rock surfaces over shrinking ­water pools, where they moved back and forth in order to soak in the remaining ­water. They are capable of ­water uptake through the ventral skin surface. During the summer rains, Canyon Treefrogs can forage away from streams for a considerable distance. Swann (2005) mentions finding them up to 100 m from ­water tucked into crevices, and Lazaroff et al. (2006) noted reports of them in a cave far from ­water.

ADULT HABITAT

As their name implies, Canyon Treefrogs occupy rocky canyons and arroyos with permanent to semipermanent ­water in the Desert Southwest. Despite the name “treefrog,” they ­really are a rock wall frog and rarely are found on trees in riparian habitats, although Lemos Espinal and Smith (2007a) noted them on trees and shrubs on rainy days. During the day, they seek refuge in rocky crevices in canyon walls and even in Black Phoebe (Sayornis nigricans) nests (Zylstra and Ward, 2013), but can be found in numbers in shallow ­water and ditches along creek beds. Minton (1958) mentioned them in “slide rock” at an unspecified “considerable” distance from a stream; Swann (2005) noted they ­were found in talus on Arizona’s “Sky Islands”; and Degenhardt et al. (1996) observed them in talus 1 km from the nearest ­water in New Mexico’s Animas Mountains. Inhabited

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Crevice habitat of Dryophytes arenicolor. Photo: Breck Bartholomew

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228  Hylidae

Canyon Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination (Hailman and Jaeger, 1974). Canyon Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling begins in March at temperatures as low as 10°C (Gates, 1957) and extends into August (Livezey and Wright, 1947; Lazaroff et al., 2006; Dayton et al., 2007). Gehlbach (1965), however, suggested 2 calling periods, which coincide with local precipitation patterns. Zweifel (1968b) found males calling usually at 21–25.2°C in the Chiricahua Mountains. Calling occurs in or immediately adjacent to a stream, such as ­under flat ledges located along a stream margin or from smooth rock walls at and above the ­water line (Degenhardt et al., 1996). Diurnally, they are heard calling from crevices or from spaces between rocks. Most calls are heard from just before dark into the night. The call of the Canyon Treefrog has been described as “that of a slightly hoarse lamb: Ba-­a-­a” (Eaton, 1935). Intercall intervals in 1 Arizona study ­were 0.20–0.28 sec, calls lasted 0.13–0.14 sec, the pulse repetition rate was 60–75 pulses/sec, and the dominant frequency was 0.50 kHz; in Utah, the intercall interval was ca. 0.59 sec, calls lasted 0.33–0.36 sec, and the pulse repetition rate was 60–240 pulses/sec (Pierce and Ralin, 1972). In another study, the number of pulses was 15–17.9, the duration ranged between 37 and 39.4 ms, the call period lasted 15.15–16.3 sec, and the high frequency peak was 2,093–2,487 Hz (all data are means; Klymus et al., 2010). Some of the variation in means represent dif­fer­ent phyloge­ne­tic histories and perhaps the

influence of past introgression with D. wrightorum (Klymus et al., 2010). Males also possess an encounter call made in response to other males calling nearby or when another male attempts amplexus. Pierce and Ralin (1972) liken this call to a series of high-­pitched “erps” or “yips.” This suggests that males establish a calling territory that they ­will defend against intruders. Pierce and Ralin (1972) described both calls in detail. Females approach calling males in response to the advertisement call. A male can detect an approaching female and move to amplex her as she approaches (Brown and Pierce, 1965). BREEDING SITES

Oviposition occurs in shallow ­water in potholes and pools in intermittent canyon streams. Preferred breeding sites lack fish and crayfish. Eggs are attached singly to weeds and brush or to rock surfaces on the bottom, although they may clump together (Campbell, 1934). Occasional eggs may break off from the substrate and float to the surface. REPRODUCTION

In Arizona, spermiogenesis occurs from at least March to September, with the smallest mature males at 28 mm SUL (Goldberg, 2020e). Females are ready to spawn from April to July, with the smallest mature females at 35 mm SUL (Goldberg, 2020e). Goldberg (2020e) suggested that females may spawn more than once in a reproductive season, based on a histological examination of ovaries from 2 females. Eggs are oviposited attached to leaves and other debris on or near the bottoms of shallow streams and potholes. Eggs ­will occasionally float on the surface or clump together, giving rise to reports of surface film egg clutches. Clutch size is several hundred eggs. Hatching occurs in ca. 3 days. LARVAL ECOL­O GY

Larval development occurs normally at temperatures >15.5°C, with the minimum developmental temperature ca. 13°C; Zweifel (1968b) suggested that 31°C was the maximum temperature for normal larval development. The duration of the larval period is 30–74 days, depending on ­water temperature (Zweifel, 1961; Wylie, 1981; Dayton et al., 2007), but some tadpoles may overwinter (in Swann, 2005). Newly transformed froglets are ca. 15 mm TL (Zweifel, 1961). DIET

Calling adult male Dryophytes arenicolor. Photo: Dennis Suhre

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The diet includes a variety of insects, including beetles, ants, true bugs, worms, and caddisflies (in Degenhardt et al., 1996).

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Tadpole of Dryophytes arenicolor. Photo: James Rorabaugh Eggs of Dryophytes arenicolor. Photo: Breck Bartholomew

PREDATION AND DEFENSE

Canyon Treefrogs are wary and readily jump t­ oward crevices to escape. Their ground color and lichen-­like patterns aid them in blending in with canyon habitats. In addition, their ability to rapidly change color allows them to blend into substrates ­under changing conditions. Their colors change quickly from light to dark, more so than from dark to light. Not surprisingly, Canyon Treefrogs choose darker substrates when given a choice, at least at lower temperatures (Swann, 2005). At higher temperatures, they ­will choose a light substrate. Canyon Treefrogs also have skin secretions that cause irritation to the eyes and mucous membranes. Tadpoles and adults are eaten by garter snakes (Thamnophis cyrtopsis, T. eques) and ­giant ­water bugs (Jameson and Flury, 1949; Wylie, 1981; Swann, 2005; Jones and Hensley, 2020). POPULATION BIOLOGY

In a study of ge­ne­tic population structure, Mims et al. (2015) found that ­there is greater ge­ne­tic structuring of populations with increasing ­water requirements in desert anurans, and this pattern is well illustrated in the population ge­ne­tic structure of Canyon Treefrogs. For D. arenicolor, the scale of population structure was rather consistent throughout the Madrean Sky Islands of southeastern Arizona, with high levels of differentiation and spatial structuring by mountain range. Ge­ne­tic distances ­were most correlated with uniform landscape re­sis­tance, with most support for the isolation by distance hypothesis. Inasmuch as aquatic habitats are threatened throughout the Desert Southwest, ­there is a threat of loss of ge­ne­tic diversity, especially since ge­ne­tic connectivity is highly corelated with terrestrial and aquatic connectivity.

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Breeding habitat of Dryophytes arenicolor. Photo: Breck Bartholomew COMMUNITY ECOL­O GY

Dryophytes wrightorum and D. arenicolor frequently are found at the same breeding sites in the mountains of central Arizona and northern México. Differences in calls, breeding season (although they do overlap), and response to rainfall

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230  Hylidae

normally are sufficient to prevent hybridization, although Klymus et al. (2010) found evidence of past introgression between them in at least 1 region.

sp., Nyctotherus cordiformis, Karatomorpha swazyi, and Hexamita intestinalis (Parry and Grundman, 1965). SUSCEPTIBILITY TO POTENTIAL STRESSORS

DISEASES, PARASITES, AND MALFORMATIONS

The fungal pathogen Batrachochytrium dendrobatidis was reported from 2 Arizona Canyon Treefrogs (Bradley et al., 2002). Canyon Treefrogs are parasitized by the cestodes Cylindrotaenia americana and Distoichometra kozloffi; the nematodes Cosmocercella haberi, Cosmocercoides dukae, and Physaloptera sp.; and larval forms of the trematode subfamily Plagiorchiinae (Parry and Grundman, 1965; Goldberg et al., 1996c). The mite Hannemania hylae is an ectoparasite of this species in Texas (Jung et al., 2001), and H. hegeneri is found on D. arenicolor from Utah (Parry and Grundman, 1965). The species also is infected by an undetermined species of trypanosome and the protozoans Zelleriella

Dryophytes avivoca (Viosca, 1928) Bird-­voiced Treefrog ETYMOLOGY

avivoca: from the Latin avis meaning ‘bird’ and voca meaning ‘call.’ The name refers to the male’s advertisement call. NOMENCLATURE

Powell et al. (2016): Hyla avivoca Fouquette and Dubois (2014): Hyla (Dryophytes) avivoca Synonyms: Hyla avivoca ogechiensis, Hyla phaeocrypta ogechiensis, Hyla versicolor phaeocrypta Cope (1889) originally assigned the name Hyla phaeocrypta to this frog, but it appears he was describing a small D. versicolor. Thus, Viosca’s (1928) name (now Dryophytes) avivoca is the proper name for the Bird-­voiced Treefrog. IDENTIFICATION

Adults. The Bird-­voiced Treefrog is very similar in appearance to members of the Gray Treefrog complex, but it is smaller and has an entirely unique voice. The dorsum is gray, greenish, or brown with a variable lichen-­like pattern, and ­there is a greenish-­white spot directly ­under the eye. If green, the coloration is entirely dorsal and may be bordered by an intermittent, thin dorsolateral stripe. The concealed portions of the thigh and sides are greenish, and the dark markings on the thigh sometimes form a vermiculate pattern. The body is

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No information is available. STATUS AND CONSERVATION

The Canyon Treefrog does not seem to have experienced population declines and is still rather abundant in habitats throughout the Desert Southwest. Individual populations may be adversely affected by introduced fish and crayfish, altered stream flow patterns, sand deposition, and ­human disturbance (Lazaroff et al., 2006). Drought may also impact local populations. Still, Griffis-­Kyle et al. (2018) considered this species one of the most vulnerable amphibians in the Desert Southwest ­because of its dependence on small stream habitats that are threatened throughout the region.

slender with a blunt snout, and the eyes are protuberant. The dorsal skin surface is smooth, but the ventral skin may be granular. Venters are dull white with throats peppered by small dark spots. Dark bars are pre­sent on both fore and hind limbs. Toe pads are con­spic­u­ous, and hind toes are webbed. The groin and posterior portion of the thighs have a light green–­spotted flash coloration. Mittleman (1945) described differences between D. avivoca and members of the Gray Treefrog complex. The maximum size is 52.5 mm SUL (Neill, 1948), and males are smaller than females. Mittleman (1945) gave a male size range of 29–40 mm SUL (mean 33.9 mm). In Oklahoma, Secor (1988) recorded calling males at 29.7– 40.5 mm SUL (mean 35.3 mm), and in Arkansas, males ­were 28.7–37.5 mm SUL (mean 35 mm) (Trauth and Robinette, 1990). In Tennessee, Parker (1951) recorded males 32– 42.8 mm SUL (mean 35.5 mm) and females to 52.5 mm SUL. Hellman (1953) recorded a single female 50 mm SUL in Florida. Alabama frogs ­were 29.9–37.7 mm SUL (mean 33.7 mm) (Brown, 1956). Larvae. The tadpole is distinctive. The body and tail are jet black with very light flecks of gold, and ­there is a yellow preorbital stripe extending from the eye to the tip of the snout. A pale, short postorbital stripe also is pre­sent. Larvae have 3–7 copper-­red ­saddles on the dorsal tail musculature. Bellies are black. The tail is long with a pointed tip, and the tail fins are grayish with black mottling. The iris has 4 gold spots. Tadpoles reach a maximum of 31–35 mm TL. Tadpoles are described by Siekmann (1949), Parker (1951), Hellman (1953), and Volpe et al. (1961). Mouthparts are figured in Volpe et al. (1961).

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Dryophytes avivoca 231

Eggs. ­There are 2 envelopes surrounding the egg, which is dark brown dorsally and cream to white ventrally. The vitellus is 1.1–1.4 mm in dia­meter; the inner envelope is 1.6–2.2 mm in dia­meter; and the outer envelope is 4–5.5 mm in dia­meter (Parker, 1951; Hellman, 1953; Volpe et al., 1961). In Tennessee, Parker (1951) recorded the vitellus as 0.8–0.9 mm in dia­meter, although the other mea­sure­ments ­were similar. This suggests the possibility of geographic variation in size. Volpe et al. (1961) suggested that the jelly film surrounding the outer envelope could be considered a third envelope; it is 0.22 mm in thickness. DISTRIBUTION

The Bird-­voiced Treefrog occurs from the South Carolina Coastal Plain (along the Savannah River just northwest of Savannah; Dodd and Barichivich, 2017) across Georgia (Altamaha River drainage to southwest of Townsend) to the Florida Panhandle, then west to the Florida Parishes of Louisiana. Populations occur west of the Mississippi River in Louisiana, southeastern Oklahoma (­Little River drainage), and southwestern and central Arkansas. The range includes the Black ­Belt of Alabama, most of Mississippi, and northward along the Mississippi River to southern Indiana (possibly extirpated) and Illinois. The species appears to be expanding its range eastward in Tennessee along the Cumberland River drainage and recolonizing some areas of southern Illinois where it had been considered extirpated

(Palis, 2021). Isolated populations also occur in north central Georgia and in the Coosa Valley of Alabama. Impor­tant distributional references include: Alabama (Mount, 1975), Arkansas (Davis and Hollenback, 1978; Trauth and Robinette, 1990; Trauth, 1992; Fulmer and Tumlison, 2004; Trauth et al., 2004), Florida (Krysko et al., 2019), Georgia (Harper, 1933; Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Illinois (Phillips et al., 1999; Redmer et al., 1999a; Palis, 2021), Indiana (Minton, 2001), Louisiana (Davis and Hollenback, 1978; Dundee and Rossman, 1989; Boundy and Carr, 2017), Oklahoma (Blair and Lindsay, 1961; Krupa, 1986b; Sievert and Sievert, 2006), and Tennessee (Redmond and Scott, 1996; Niemiller and Reynolds, 2011). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Dryophytes avivoca was described by Viosca (1928), who noted that the species previously had been confused with members of the Gray Treefrog complex. It is most closely related to D. andersonii and slightly more distantly to the Gray Treefrog complex (Hedges, 1986). Contrary to suggestions based on call structure, ­there is no D. versicolor group (containing D. avivoca) as proposed by Blair (1958c). ­There are 2 clades within D. avivoca, a western clade and an eastern clade, which was described as Hyla phaeocrypta ogechiensis (Neill, 1948; Hedges, 1986). The primary difference among subspecies involves postfemoral coloration (Neill, 1954). The validity of this taxon needs clarification. In the laboratory, D. avivoca hybridizes successfully with D. cinereus and D. chrysoscelis (Mecham, 1965; Fortman and Altig, 1973). Fortman and Altig (1973) described the hybrid tadpoles. Crosses with D. femoralis, D. gratiosus, and D. squirellus ­were generally unsuccessful, but success may depend on the sex of the parents (Mecham, 1965). Natu­ral hybridization with D. chrysoscelis was reported by Mecham (1960b) in Alabama. The diploid chromosome number is 24, and Bushnell et al. (1939) described the chromosomes. ADULT HABITAT

Distribution of Dryophytes avivoca. This species may occur in adjacent regions of Texas and northwestern Louisiana.

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This is a species of the cypress-­(Taxodium) tupelo-­gum headwater swamps, swampy floodplains, bottomland and slope forest, and swampy streams, lakes, rivers, and ponds of the Southeast. It frequents buttonbush (Cephalanthus), reed thickets, and other woody plants in riparian habitats. ­These habitats are often flooded in winter but hold only a ­little ­water in summer. The canopy is usually dense and the understory community diverse. In southern Illinois, D. avivoca has ventured away from its traditional habitat to

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232  Hylidae

Tadpole of Dryophytes avivoca. Photo: Stan Trauth

wiping is impor­tant in allowing frogs to remain exposed in arboreal perches. Bird-­voiced Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient t­ oward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Bird-­voiced Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Adult Dryophytes avivoca, gray phase. Photo: Aubrey Heupel

colonize newly created open-­water wetlands lacking trees, except at the margins from which they call (Palis, 2020b). Habitat descriptions are in Carr (1940a), Enge et al. (1996), and Redmer et al. (1999a). TERRESTRIAL ECOL­O GY

During the nonbreeding season, Bird-­voiced Treefrogs are found in lowland forests and swamps adjacent to summer habitats, where they are observed hiding in tree crevices, in stumps, and occasionally sitting on palmetto fronds. Juveniles are found in late summer to autumn perched in low shrubs (18.4°C. Males call from woody vegetation, trees, and downed logs at perch heights of 0.1–7 m, mostly above the ­water (from 2 m over land to 15 m into the ­water) (Secor, 1988; Redmer et al., 1999a); most males are 1–2 mm above the ­water. A few males call from the shoreline, whereas ­others are found to >3 m above ­water. Perch sites range from narrow vines to the trunks of large trees, but most males call from low branches and vines. Females approach a calling male and initiate amplexus by touching him. She ­will make short, frantic movements ­toward him ­after each call ­until she is close to him (Trauth and Robinette, 1990). As soon as he is touched, the male turns and amplexes the female in an axillary position. ­After initiating amplexus, the male continues to make a raspy call for a while. The amplexed pair then moves head first down from the perch site to the ­water. At the ­water’s surface, the female turns around so that she is one-­third submerged and the male is half submerged in the ­water. Oviposition begins immediately. The female arches her head and back and extends her rear legs in the ­water. Eggs are extruded in packets in 1–4 min intervals (Redmer, 1998b). Oviposition takes >1.5 hrs.

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Dryophytes avivoca 233

Males appear to be territorial and ­will engage another male that encroaches too closely to a calling perch. Secor (1988) recorded intercalling male mean distances of 3.9–7.8 m; however, occasional calling males may be much closer. Once a calling male detects an intruder (from as far away as 45 cm), he switches from the advertisement call to a short trilling chirp (Altig, 1972b). If the intruder does not depart, the calling male ­will approach and challenge him. A wrestling match ensues, with attempted amplectant be­hav­ior and constant chirping by the resident male. Resident males usually win wrestling matches, which end with the intruder’s departure. Encounters last up to 15 min. Male Bird-­voiced Treefrogs also produce a warning call (a throaty “reek-­reek-­ reek”) if another male approaches too closely (Palis, 2020b).

likely include snakes. Larvae are undoubtedly eaten by aquatic invertebrates. POPULATION BIOLOGY

No information is available. DISEASES, PARASITES, AND MALFORMATIONS

Biting midges of the genus Corethrella feed upon D. avivoca and may be attracted by the frog’s advertisement call (McKeever, 1977; McKeever and French, 1991). Paramecium-­like protozoans ­were observed feeding on eggs of this species in Mississippi (Tice et al., 2016). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. BREEDING SITES

Breeding occurs in riparian habitats, river bottomlands, and around ponds in cypress-­tupelo swamps. ­Water below calling sites is shallow, usually 0.1–0.7 m (mean 0.3 m) (Redmer et al., 1999a).

STATUS AND CONSERVATION

Villena et al. (2016) suggested populations in the South ­were declining based on probability of occurrence through time.

REPRODUCTION

Eggs are oviposited in packets of 3–15 eggs (Parker, 1951; Redmer, 1998b). Hellman (1953) estimated total clutch sizes of 567 and 720 eggs from 2 dissected individuals, and Redmer (1998b) estimated a clutch size of 150–180 from a single female observed over a 40 min period. Trauth and Robinette (1990) found a mean clutch size of 838 from 3 females and 315 from another female. It appears that females are capable of ovipositing several times during a breeding season. Eggs are oviposited in small clumps that quickly break apart and sink to the bottom of the substrate or adhere to nearby vegetation. Volpe et al. (1961) described hatching and early larval development in detail. Hatching occurs at ca. 40 hrs ­after deposition.

Adult Dryophytes avivoca, green phase. Photo: Kenny Wray

LARVAL ECOL­O GY

Larvae reared in the laboratory at 28.5–35.5°C required only 29 days to complete development. Recently transformed froglets mea­sure 9.4–13.2 mm SUL (Volpe et al., 1961). DIET

The diet consists of small invertebrates, particularly adult and larval beetles and lepidopterans. Other prey includes ants, treehoppers, leafhoppers, bark lice, assassin bugs, mites, and spiders (Jamieson et al., 1993; Redmer et al., 1999b). Dietary items reflect an arboreal existence. PREDATION AND DEFENSE

The dorsal coloration is cryptic and makes the frogs difficult to locate. Predators have not been reported, but

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Breeding habitat of Dryophytes avivoca. Photo: Stan Trauth

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234  Hylidae

Protection of riparian corridors is absolutely essential for this species, as it does not disperse widely away from bottomland swamps and riparian habitats (Burbrink et al., 1998). Bird-­voiced Treefrogs have colonized restored and constructed wetlands, but it may take several years for successful colonization to occur (Palis, 2012b, 2020b). Disturbances may or may not have significant impacts on populations of Bird-­voiced Treefrogs. Following Hurricanes

Ivan and Katrina in Louisiana, Bird-­voiced Treefrog numbers actually increased substantially in swamp habitats (Schriever et al., 2009). Florey and Mullin (2005) could detect no trends in D. avivoca populations in Illinois during surveys from 1986 to 1989. This species is considered Threatened in Illinois, although Palis (2021) has now documented its occurrence at 32 new locations, including swamp remnants, a stream channel, and human-­made ­water bodies.

Dryophytes chrysoscelis (Cope, 1880) Cope’s Gray Treefrog

(D. versicolor has a slow trill [17–35 notes/sec], whereas D. chrysoscelis is fast trilling [34–69 notes/sec]); cytology (cells of D. versicolor are larger with more nucleoli than ­those of D. chrysoscelis) (Cash and Bogart, 1978). ­There are no morphological characters related to body proportions or coloration that accurately separate ­these species (Matson, 1990). Adults. Cope’s Gray Treefrog is a small to medium-­sized frog with a distinctive lichen-­like dorsal pattern. The lichen coloration is composed of vari­ous gray to buff patches; sometimes the patches may be distinctively greenish. ­There is a con­spic­u­ous light patch under­neath the eye, and the eyes are prominent. The toes are tipped by con­spic­u­ous toe pads. The rear toes are partially webbed, but ­there is only a slight trace of webbing between the fin­gers. The concealed portions of the inner thigh, shanks, groin, and axilla are bright orange and unspotted. The undersides of the body are unpigmented, but the throat of the male is dark during the breeding season. A blue (axanthic) D. chrysoscelis has been reported from Minnesota (Oldfield and Moriarty, 1994). Juveniles are a yellowish to olive tan to gray dorsally, frequently with a dark band between the eyes. The belly is white to cream, with the dark-­colored intestinal vein obvious. Males are smaller than females, and the male’s vocal sac appears as a transverse fold when not inflated. Adults reach maximum size quickly and show ­little growth between breeding seasons (Ritke et al., 1990). Sexual maturity is reached in 2 years (Jensen et al., 2008). Size reports vary for ­Virginia, with Mitchell (1986) reporting males averaging 42 mm SUL and females 48 mm SUL, and Hoffman (1946) reporting males to reach sexual maturity at 34–40 mm SUL (mean 37.2 mm). In Wisconsin, males are 27–42 mm SUL (mean 35.2 mm SUL) (Jaslow and Vogt, 1977). In Nebraska, males are 31–47 mm SUL (mean 38 mm SUL) and females 40–50 mm SUL (mean 44 mm SUL) (Lynch, 1985), whereas in Tennessee males are 39–53 mm SUL (mean 45.9 mm SUL, mass 7.6 g) and females 45–62 mm SUL (mean 52.4 mm SUL, mass 11.2 g) (Ritke et al., 1990). Larvae. Hatching occurs at a total body length of approximately 4.1–4.7 mm in Florida, and by 19 days, the total

ETYMOLOGY

chrysoscelis: from the Greek chrysos meaning ‘gold’ and kelis meaning ‘spot’ or ‘stain.’ The name refers to the golden spots on the back of the thigh of this gray treefrog. The name Cope in the common name refers to famed herpetologist and paleontologist Edward Drinker Cope who described this species in 1880. NOMENCLATURE

Powell et al. (2016): Hyla chrysoscelis Fouquette and Dubois (2014): Hyla (Dryophytes) chrysoscelis Synonyms: Hyla femoralis chrysoscelis, Hyla versicolor chrysoscelis, Hyla versicolor sandersi ­There are many publications where the identity of the species of “Gray Treefrog” is uncertain between D. versicolor and D. chrysoscelis. For example, Fellers (1975) discusses intermale be­hav­ior and aggression in D. versicolor, but in a ­later paper (Fellers, 1979a), the characteristics described seem more associated with D. chrysoscelis (e.g., in intermale calling distances). For this reason, this information is included in the D. chrysoscelis account. In other situations, it may not be pos­si­ble to identify the species involved, especially when ranges overlap (e.g., Bowers et al., 1998; Anderson and Arruda, 2006; Swanson et al., 2019). Many characteristics (morphology, calling season, be­hav­ior, some life history traits, and desiccation rates) may be applicable to both species. It is best to check the context of the original citation for confirmation. IDENTIFICATION

This species is morphologically identical with D. versicolor. It can be differentiated by the following characteristics: chromosome number (D. versicolor is tetraploid [n = 48], whereas D. chrysoscelis is diploid [n = 24]); call rate

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Dryophytes chrysoscelis 235

length increases to 11.5 mm (Hellman, 1953). Jensen et al. (2008) report hatching at 6–7 mm TL in Georgia. The general body coloration may be several shades of brown, with numerous small black and gold flecks scattered across body and tail. Body coloration is not affected by the color of the ­water in which development occurs (Akers, 1997). Tail fins are basically colorless in terms of the background, but they are heavi­ly mottled with black pigment. ­There is a pair of preorbital stripes, but they are not distinctly outlined and only become apparent ­after 2 weeks. The venter is immaculate anteriorly with some gold flecking, and the intestines are readily vis­i­ble through the body wall. As the tadpole grows, the venter becomes more cream colored posteriorly with heavy intrusions of gold flecks. The iris is bright gold. Total tadpole lengths may reach 64–65 mm just before metamorphosis (Jensen et al., 2008), but the largest larva mea­sured by Wright (1929) in the Okefenokee Swamp was 46.6 mm TL. Tadpole descriptions are in Hellman (1953) and Altig (1970). ­There is a unique color morph of D. chrysoscelis tadpoles that appears only when predators are abundant at a breeding pond. Such tadpoles are longer with shallower bodies and develop bright red tail fins with dark margins. When predators, such as dragonfly larvae, are absent, the bright red coloration does not develop, and tail fins and bodies tend to be broader. The red coloration can be induced experimentally by raising larvae in the presence of predators (McCollum and Van Buskirk, 1996). Bright tail fins are seen only on the oldest and largest larvae, usually within about 2 weeks of metamorphosis. Eggs. Most descriptions of the eggs of Gray Treefrogs do not differentiate between D. chrysoscelis and D. versicolor. ­There appears to be some variation in egg size, number of eggs per packet, and total clutch size, but ­whether ­these variables are species specific or reflect intraspecific variation is unknown. The eggs of D. chrysoscelis likely are similar to ­those described for D. versicolor. Eggs are deposited initially in small floating packets of 15°C. Arboreal calling occurs from trees or shrubby vegetation bordering the breeding sites, where males call from perches located 0.5–6 m (usually 1–2 m) above the ­water level; perch dia­meters range from 1 to 30 cm (Godwin and Roble, 1983; Ritke et al., 1990). Perches tend to be horizontal or located at a 45° ­angle, but Cope’s Gray Treefrogs also can call from vertical tree trunks. In prairie regions, Cope’s Gray Treefrogs call from the ground next to breeding pools, where they have been observed calling from bare ground, grass, and floating algae. Males call from the same general area throughout the breeding season, although not necessarily from the same perch or site each night. In areas of sympatry, D. versicolor calls from higher locations than D. chrysoscelis (Johnson, 1966). As with Gray Treefrogs, dominant males are territorial and aggressively defend calling sites by kicking, shoving, head-­ butting, or jumping on the interloper. Dominant males space themselves at intervals of ca. 75 cm (Fellers, 1979a). Few (8 ha in Palis, 2007) and followed over a multiyear period, successful colonization is more likely to be documented. Fire may have variable effects on Cope’s Gray Treefrogs. ­Under experimental conditions, tadpoles grow faster on unburned leaf litter than on burned litter. Tadpoles in burned sites also tend to be smaller than in unburned sites, although survivorship does not appear to be affected (McDonald et al., 2018). Females also choose unburned litter over burned litter as oviposition sites. Still, McDonald et al. (2018) noted no differences in abundance or morphometric mea­sure­ments among adults in dif­fer­ent burn treatments, suggesting that adults are not affected by fire management. When options are available to adults, they tend to choose unburned plots over burned plots for breeding.

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As with most anurans, roads and highways cause considerable mortality. At high road densities, D. versicolor/D. chrysoscelis populations are negatively impacted at small local scales, but not over a more widespread regional scale (Marsh et al., 2017). Impervious surfaces do not seem to impact populations per se. Call surveys used to monitor this species can be effective if ­either 5 or 10 min listening

stops are used to rec­ord presence (Sargent, 2000; Burton et al., 2006). However, calls of this species have been misidentified for other species during call surveys (Lotz and Allen, 2007). A total of 7,268 Cope’s Gray Treefrogs ­were collected commercially in Florida from 1990 to 1994 (Enge, 2005a). This species is considered Endangered in New Jersey.

Dryophytes cinereus (Schneider, 1799) Green Treefrog

Naturalist (Anonymous, 2007), but albinos are unknown. The diploid chromosome number is 24 (Bushnell et al., 1939). Females tend to be only slightly larger than males based on both SUL and tibiofibula length (Gunzburger, 2006). Adults are 40% of known hybrids could be misclassified as pure parental species based on phenotype alone, had not ge­ne­tic analyses proved other­wise. However, ­there is no evidence that morphological asymmetry

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is greater in hybrids than in the parental species, suggesting that hybrids are not subject to decreased levels of developmental stability (Lamb et al., 1990). Hybrids may exhibit spectral variability in call characteristics, thus resembling ­either of the parental species, although the ranges of variation tend to overlap with D. cinereus (Gerhardt et al., 1980). Most introgressive hybridization occurs between male D. cinereus and female D. gratiosus. Hybridization experiments with other species (D. arenicolor, D. versicolor complex, Pseudacris clarkii) have not proven successful (e.g., Littlejohn, 1961a; Pierce, 1975). However, Gerhardt (1974b) reported a single laboratory cross between a female Dryophytes cinereus and a male D. andersonii that resulted in normally developing larvae, and Anderson and Moler (1986) reported a single hybrid between D. andersonii and D. cinereus from Florida. A putative natu­ral hybrid between D. cinereus and D. chrysoscelis was reported from Louisiana (Glorioso et al., 2015). ADULT HABITAT

When not at or near wetlands during the breeding season, Dryophytes cinereus lives in forested areas adjacent to freshwater breeding sites. They are frequently observed in upland hardwood forest, hardwood hammock, bottomland forest, old fields, ecotones between forested and open areas, broad-­leaved marshes, sawgrass marsh, sloughs, canals, wetland forest, woody shrub habitats along stream channels, ravines (Enge et al., 1996; Enge, 1998a; Enge and Wood, 1998; Meshaka et al., 2000; Donnelly et al., 2001; Meshaka and Layne, 2015), and riparian areas (Rudolph and Dickson, 1990; Burbrink et al., 1998; Muenz et al., 2006). For much of the year, adult habitat consists of the margins of the wetlands in which they breed. They move to mostly permanent lakes and ponds in spring (March–­April) and return to terrestrial feeding areas and refugia in the autumn, usually from September to November. Green Treefrogs are also associated with habitats that may be saline, such as mangrove forest (Carr, 1940a; Meshaka et al., 2000), coastal marshes (Allen, 1932; Burger et al., 1949; Werler and McCallion, 1951; White and White, 2007), brackish areas of the Florida Keys (Peterson et al., 1952), rivers with saltwater tidal fluxes (Dunn, 1937), salt marshes (Neill, 1958), and splash pools and marshes near Chesapeake Bay (Noble and Hassler, 1936; Cooper, 1953; Hardy, 1953) and Mobile Bay (Neill, 1958). TERRESTRIAL ECOL­O GY

Dryophytes cinereus usually becomes active in late spring in the north and remains active ­until October (e.g., Rossman, 1960; Meshaka et al., 2020c); even in ­Virginia, however, Green Treefrogs may be found as early as January (Meshaka

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et al., 2020c). As one proceeds south, the activity season is extended, and in the southern Coastal Plain and Florida, frogs are active year-­round. Green Treefrogs are primarily active at night, at dusk, and on cloudy or rainy days with high humidity. It is during ­these times when they are most likely to be feeding or moving to and from calling perches. During the day, they remain hidden in refuges or clinging to vegetation where they are exposed directly to the ele­ments. Freed (1980b) termed this basking be­hav­ior, and showed that growth rates are increased by basking. ­Under laboratory conditions, frogs spend considerable time basking, and may raise their body temperatures >6°C above nonbasking frogs. Individual frogs are often difficult to observe ­because of their green coloration, lack of movement, and the tucked-in position of the limbs. Indeed, Hobson et al. (1967) suggested that Green Treefrogs may be sleeping. Once 1 is seen, an observer may develop a search image and see dozens more attached to vegetation, especially cattail Typha stems, branches, or palm fronds. In extreme southern Florida, small individuals are frequently found arboreally in bromeliads (Neill, 1951). Green Treefrogs remain in the vicinity of the calling site throughout the breeding season. Spring and summer retreat and feeding sites are usually located in close proximity to the calling site. Daily movement from calling perches to the diurnal retreat site occurs from 01:00 to 03:00 and may involve traveling to and from cattail stems only a few meters away. Green Treefrogs are frequently associated with ­human habitations, and they readily take shelter near buildings or in pipes, gutters, electric or cable boxes, or other suitable places associated with anthropogenic structures (Goin, 1958). They may return to the same refuges ­after spending the eve­ning feeding, or they may choose other sites or move around from location to location. Occasionally, frogs may be active at temperatures approaching 4°C, but most activity occurs at temperatures >13°C. The CTmin is 3.6–4.6°C and the CTmax is 36.6°C (John-­Alder et al., 1988). In the southern portion of its range, D. cinereus never becomes dormant for long periods. During cold or dry weather, it shelters ­under coarse woody debris or boards, ­under bark, in sawdust piles, treehole crevices, or palm thatch, and around buildings or other protected places (Fontenot, 2011; Meshaka and Layne, 2015). The extent to which Green Treefrogs move away from breeding sites is unknown, although Dodd (1996) recorded D. cinereus 457–914 m (mean 545 m) from the nearest pos­si­ble breeding site in north Florida sandhills. In northern Florida, ­there appears to be a peak in movement to winter retreats in November, but even in winter frogs have been recorded moving as far as 238 m between retreat sites (Zacharow et al., 2003). In extreme south Florida, Green Treefrogs tend

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to occupy cypress and marsh habitats rather than prairie habitats during the dry season, but in the wet season such habitat differences are not apparent (Waddle, 2006). Capture probabilities are inversely correlated with marsh depth, suggesting that Green Treefrogs disperse during the wet season (summer) and are more concentrated in the dry season (autumn and winter). Dryophytes cinereus is not particularly ­adept at absorbing ­water from moist soil, which prob­ably inhibits its activity during drought or dry weather (Walker and Whitford, 1970). However, they are able to slow down evaporative ­water loss better than most nonarboreal species (Wygoda, 1984). To do this, Green Treefrogs engage in a complex series of movements whereby the limbs are used to wipe the head and body with an extraepidermal layer of mucous and lipids, which helps retard ­water loss (Barbeau and Lillywhite, 2005). Body wiping is impor­tant in allowing frogs to remain exposed in arboreal perches. Green Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Green Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

In the South, occasional calls are heard beginning in late February (Carr, 1940b), although the breeding season does not begin in earnest ­until ­later in April; choruses have been heard as late as early September (Bancroft et al., 1983; Dundee and Rossman, 1989; Gunzburger, 2006) or October (Einem and Ober, 1956; Meshaka and Layne, 2015) in Florida and Louisiana. In Texas, calling occurs from April to July (Car­ter et al., 2018). In the northern portion of the range, such as in Illinois and North Carolina, calling begins in May and extends to August (Garton and Brandon, 1975; Redmer et al., 1999a; Gaul and Mitchell, 2007). Environmental conditions such as temperature appear to control the onset of breeding. Arboreal or ground-­level calling occurs from shrubby vegetation surrounding a lake or pond or from emergent vegetation (to about 30 cm from the ­water’s surface) around a lake’s shoreline and shallows. In Florida, Bancroft et al. (1983) recorded calling from cattail, pickerel weed (Pontederia), and ­water hyacinth (Eichhornia). Green Treefrogs rarely call from the water-­shoreline interface or from ­water as does the closely related Barking Treefrog. They may occasionally call from such locations, however, if the shrubby area around the breeding site is removed or altered. In such cases, the habitat change and subsequent shift in calling site may cause

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250  Hylidae

them to come into contact and breed with D. gratiosus, resulting in hybridization (Lamb, 1987). A chorus of Green Treefrogs usually begins with 1 or a few males initiating a calling bout. More and more males join into a chorus, ­until virtually all males are calling and the pond or lake reverberates in a deafening cacophony of noise. A bout may continue for several minutes, followed by a sudden cessation of calling. A number of minutes might then pass in silence ­until the chorus bout repeats itself. The cycle of slow buildup to a full chorus followed by silence continues ­until the early morning hours. On a large lake, calling bouts among Green Treefrogs give the impression of waves of noise as the cycle repeats itself among resident subpopulations. Carr (1940a) noted that choruses of Green Treefrogs may continue unbroken for 10–12 km of river front. Calling begins in earnest at dusk and continues to ­after midnight. In South Carolina in June and July, most calling occurred from 22:00 to 01:30 (Mohr and Dorcas, 1999). Calls are also heard on overcast or rainy days. Male Green Treefrogs produce 2 types of calls during the breeding season, a mating call and a “pulsed” call. The mating call duration varies from 0.10 to 0.29 pulses/sec, with repetition rates of 0.27–1.1 sec (Blair, 1958a; Oldham and Gerhardt, 1975). The high frequency part of the mating call centers around 3.0–3.8 kHz, and the low frequency peak ranges from 0.7 to 1.250 kHz (Gerhardt, 1982). In playback experiments, females prefer calls with low frequencies of 0.8–1.0 kHz and high frequencies of 2.4–3.6 kHz (Gerhardt, 1987), preferences that are in­de­pen­dent of female body size. Female preference allows for species-­specific discrimination, so that although D. cinereus and D. gratiosus are often found calling from the same site, mating between the two does not normally occur ­unless the habitat has been altered or satellite males intercept a heterospecific female. The pulsed call is an agonistic or encounter call made by a dominant male ­toward a subordinate or nearby male. Males may occasionally produce a call that is intermediate between ­these two. The calling male ­will turn to face the direction of the intruder. The encounter call is then produced when the approaching male comes within ca. 170 cm of the primary calling male (Fellers, 1979a). Females are able to discriminate among the dif­fer­ent calls and are attracted primarily to the male’s breeding call (Oldham and Gerhardt, 1975). Indeed, her call discriminatory ability is graded in such a manner as to facilitate identification among calling males, rather than responding to categorical cues that would only identify a male as being a male (Gerhardt, 1978b). Calls are projected omnidirectionally, thus making it pos­si­ble to attract females no ­matter where they are located in relation to the calling male. Females, however, locate calling males by using lateral head movements, and by

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elevating and lowering the head as they move ­toward a calling male. ­These head movements help them locate arboreally calling males, perhaps by using a sound pressure gradient system similar to that used by insects to localize sounds (Gerhardt and Rheinlaender, 1982). This is impor­ tant, since large choruses (>200 calling males to literally thousands around large lakes) likely mask the acoustic signals of all but the closest males. Females thus choose from among the 3–5 males closest to her, essentially allowing her to select the best that is available close at hand (the “best of N”; Gerhardt and Klump, 1988). As might be expected ­under such circumstances, ­there is no evidence for size-­ assortative mating (Gerhardt, 1982, 1987). Satellite males frequently accompany calling male D. cinereus (to 18% at a Georgia study site, Perrill et al., 1978). A single calling male may have 1 to 3 satellite males, usually within 50 cm of the caller. Calling males attempt to thwart satellite males through encounter calls to warn the interloper away, but they may also resort to wrestling, head-­butting, or chasing the intruder away. If a calling male leaves or is removed from a pair-­wise confrontation, the satellite male ­will assume a mating call (Perrill et al., 1982). Calling males also may become satellites in the presence of another nearby calling male. Satellite males are occasionally successful in intercepting females approaching a calling male, a strategy known as sexual parasitism. The propensity of satellite males to intercept females has led some authors to suggest that the directional introgression seen in hybrid populations of D. cinereus and D. gratiosus results from satellite male D. cinereus intercepting D. gratiosus females on their way to the water/water-­shoreline calling sites of conspecific males (Lamb and Avise, 1986). As it is for other hylid treefrogs, calling is an energetically expensive activity for Green Treefrogs, one that is almost entirely aerobic and may exceed the level of energy required for locomotion (Prestwich et al., 1989). Calling requires 7 times the metabolic activity of a Green Treefrog at rest. This is not surprising, since Green Treefrogs can utter 3,500 calls/ hr (at 27°C) during peak calling per­for­mance. Axillary amplexus is initiated when a female approaches and touches a calling male or crawls over his back. Males stop calling ­after amplexus, but ­there is 1 report of a male calling while still in amplexus ­under natu­ral conditions (McCallum and Trauth, 2009). At her approach, he turns and ­faces the female, and his call rate and pitch increase. ­After she touches him, he immediately amplexes her. At this point, ovulation begins as denoted by muscular contractions along the female’s flank. Egg deposition occurs within 4–5 hrs. BREEDING SITES

Green Treefrogs breed around a wide range of mostly permanent wetlands, from ponds and swamps to the margins

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of large lakes. Abundance is positively associated with wetland size and depth when considered in a landscape of seasonally flooded ponds, and the most used breeding sites are ­those located within 50–100 m of forested habitat (Babbitt et al., 2006). In this latter study, pasture ponds ­were deeper, larger, of higher pH (6.0 vs. 5.3), and of greater conductivity (120 μs vs. 40 μs) than wetlands located within woodlands. Dryophytes cinereus readily uses anthropogenic sites, such as retention ponds, impoundments, golf course ponds, and drainage ditches (Lichtenberg et al., 2006; Dodd and Barichivich, 2007; Scott et al., 2008; Birx-­Raybuck et al., 2010). Reproduction may occur in small, isolated wetlands, such as freshwater marshes, cypress savannas, or cypress/gum ponds (Babbitt and Tanner, 2000; Eason and Fauth, 2001; Russell et al., 2002a; Surdick, 2005; Liner et al., 2008). Green Treefrogs prefer wetlands with substantial amounts of emergent vegetation, including ­water hyacinth (Kilby, 1936; Goin, 1943), bordering shrubs, and other types of low vegetation around the shoreline, from which they call. Sites with dense mats of floating and subsurface vegetation are preferred, as ­these sites provide protection for eggs and larvae. For example, ­these frogs are abundant in the wet prairies of the Okefenokee Swamp (Wright, 1932). As with many species, many potential breeding sites are not occupied annually for reproduction. For example, Babbitt et al. (2006) recorded breeding by D. cinereus at 45% of the 78 seasonally flooded wetlands surveyed on an agriculturally modified tract of land in southern Florida. The presence or absence of fish does not appear to influence the choice of breeding sites. REPRODUCTION

Reproduction takes place from early spring throughout the summer, when temperatures are >20°C. Eggs have a minimum temperature tolerance of 20°C and a maximum tolerance of 34–39°C (Ballinger and McKinney, 1966). Mated pairs are found in March in Alabama and Florida, although most breeding occurs from April to July or August throughout much of the species’ range (Wright, 1932; Carr, 1940a; Moulton, 1954; Mecham, 1960a; Mount, 1975; Gunzburger, 2006; Meshaka et al., 2020c). Breeding is nocturnal, and rainfall is not required to trigger it. However, the greatest amount of breeding occurs during warm rain events (e.g., Richmond and Goin, 1938). Diurnal breeding activity is also triggered during humid and overcast conditions. ­After the female initiates amplexus, the mated pair remains stationary for several hours before entering the ­water. Ovulation occurs 4–5 hrs ­after the female touches the male, leading most oviposition to occur during the early morning hours (02:00–04:00). Indeed, Garton and Brandon (1975) suggested auditory stimuli are responsible for initiating

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ovulation, w ­ hether or not a female enters into amplexus. Ovulation takes 15–35 minutes. Females depress their back and extend their limbs downward. This ­causes the vent (the opening to the cloaca) to rise and to be near the male’s vent. The male then fertilizes the eggs as they are extruded. Eggs (10–50 at a time) are sent backward from the female’s vent ­under the ­water’s surface, where they adhere to floating vegetation. The eggs must remain near the surface, as the warm ­waters of the pond and lake are often oxygen depleted even a short distance below the ­water’s surface. Hatching occurs in 2–3 days. Garton and Brandon (1975) provided a detailed description of mating be­hav­ior and oviposition. The degree of heterozygosity in D. cinereus is impor­tant to reproductive success, but it varies between the sexes (McAlpine, 1993). In wild-­caught females ­under laboratory conditions, clutch size and the number of offspring surviving through metamorphosis was correlated positively with heterozygosity, and the percentage of offspring that hatched from eggs showed a similar trend, although it was not statistically significant. In contrast, body size was not correlated with heterozygosity in ­either males or females, and the number of eggs that hatched was not correlated with the number of heterozygous loci in males. The total reproductive success of females was correlated with heterozygosity, but not with fitness traits such as body size (McAlpine, 1993). McAlpine and Smith (1995) concluded that multilocus heterozygosity was not a good indicator of ­either survival or mating success in Green Treefrogs. Clutch size ranges from 359 to 2,658 (mean 1,214) in Florida (Gunzburger, 2006); a mean of 700 in Illinois (Garton and Brandon, 1975); a mean of 790 in Georgia (Perrill and Daniel, 1983); a mean of 2,152 (range 1,348–3,946) in Arkansas (Trauth et al., 1990); a mean of 1,472 eggs, with high hatching success (mean 87%, range 33–100%) in South Carolina (McAlpine, 1993); a mean of 1,271 (range 689– 2,228) in ­Virginia (Mitchell and Pague, 2014); and, also in ­Virginia, a mean clutch size of 884 eggs (range 442–1,473) (Meshaka et al., 2020c). Some of the variation in the lit­er­a­ture reports of clutch size may result from small sample sizes and the confusion of singular clutch counts with the number of ova produced per season. Perrill and Daniel (1983) noted that female D. cinereus may deposit from 1 to 3 clutches per season, with a mean of 19.3 days between clutches. Single clutches ranged from 275 to 1,160 eggs (mean 275). Female size is positively correlated with clutch size and weakly correlated with the size of the amplexing male. LARVAL ECOL­O GY

Larvae hatch into shallow areas with dense submergent or floating vegetation, such as hyacinth mats in Florida. The dense vegetation provides both cover and food for the

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252  Hylidae

Tadpole of Dryophytes cinereus. Photo: Dirk Stevenson

Tadpole of Dryophytes cinereus with black-­spotted tail fins. Photo: Joe Mitchell

growing larvae. This is impor­tant since the late hatching time means that the vulnerable larvae ­will be exposed to predators that may have been in the wetland for a time and so be of a size to easily consume them. Babbitt et al. (2006) found a larval density of 0.078/m2, a value lower than most other summer season breeders. The duration of the larval period (normally 5–9 weeks) and the size of the larvae as they approach metamorphosis are dependent on food levels and the temperature at which development occurs, among other ­factors. ­Under experimental conditions, larvae raised at high temperatures (30°C) develop more quickly (24.4 days) but are smaller at metamorphosis (0.90 g) than larvae raised at lower temperatures (25°C, 1.18 g, 35.7 days). Likewise, D. cinereus larvae raised ­under high food availability are much larger (1.22 g) and complete metamorphosis sooner (29.4 days) than larvae raised ­under more restricted food intake conditions (0.67 g, 36.8 days) (Blouin, 1992a). Size at metamorphosis is

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correlated positively with the length of the larval period and larval growth rates, and ­there appears to be a ge­ne­tic component between the size at metamorphosis and larval growth rates (Blouin, 1992b). According to Wright (1929), the largest tadpoles grow to 40 mm TL just prior to metamorphosis. Newly transformed D. cinereus are 11.5–17 mm SUL (Wright, 1932). Dryophytes cinereus larvae occasionally are vulnerable to pond or lake drying, even though they prefer permanent ­water bodies for reproduction. ­Whether a pond dries slowly or rather quickly does not influence the duration of the larval period or the body size at metamorphosis per se ­under experimental conditions. This is not surprising given that this species normally does not face desiccating conditions and that se­lection may ­favor short larval periods for a species cohabiting with fish predators. However, larval density increases as ­water evaporates, and the increase in density may result in extended larval periods or in metamorphosis at a smaller size than normal (Leips et al., 2000). ­Under experimental conditions, the larval period lasts 35–58 days. In Illinois, Garton and Brandon (1975) report a larval period of 4–6 weeks, whereas in Georgia the larval period is 55–63 days (Wright, 1932). Green Treefrog larvae have been observed in ­water that is saline, to 8.3‰ (Diener, 1965). Normally lethal salinity levels are 8–12‰ (Schriever, 2007). Salinity decreases sperm motility and velocity, and weakly brackish pools have lower levels of oviposition than freshwater pools (Wilder and Welch, 2014). The developmental rate of tadpoles is inversely correlated to the salinity of the ­water. At 2‰, the larval period takes a mean of 25 days, whereas at 6‰, the mean is 30.8 days. Tadpoles that survive 8‰ salinity take a mean of 34 days to complete development. In addition, tadpoles developing at higher salinities have smaller body masses at metamorphosis (Schriever, 2007). ­These data suggest that salinity may have sublethal effects even among surviving larvae. ­Under normal circumstances, body mass and TL are positively correlated with the length of the larval period. DISPERSAL

Recently metamorphosed D. cinereus tend to remain in the immediate vicinity of the natal breeding site during their first winter period, although they have been recorded as far as 90 m from the breeding site into surrounding forest (in Jensen et al., 2008). The metamorphs grow quite rapidly, and some individuals approach adult size before they enter their first winter dormancy (Garton and Brandon, 1975). Wright’s (1932) interpretation of the relationship between size classes and age is prob­ably in error.

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DIET

Feeding occurs primarily during dusk and at night, as the frog is far more active nocturnally than during the day. During cloudy or rainy days of high humidity, the frogs ­will be diurnally active and more likely to feed ­later in the after­noon. Feeding occurs throughout the reproductive season, as mated pairs both contain food items (Kilby, 1945). The diet consists of prey indicative of the shrubs and herbs on which the species is found. Insect prey primarily includes flies, moths, crickets, dragonflies, butterfly adults and larvae, ants, bees and wasps, and beetles. Green Treefrogs are primarily insectivorous, although they readily consume mollusks (snails), spiders, mites, collembolans, millipedes, and phalangids (daddy-­longlegs). Inorganic debris and epidermis are found frequently in the digestive tract (Haber, 1926). Cannibalism also has been reported (Höbel, 2011). In frogs obtained from the Okefenokee Swamp in Georgia, Haber (1926) reported a high percentage (92%) of frogs with food in their digestive tracts, which included recently eaten insects (68%), orthopterans (24%), coleopterans (24%), lepidopterans (21%), hymenopterans (15%), hemipterans (13%), spiders (24%), and epidermis (35%). Many digestive tracts contained nematodes (41%) and parasitic protozoans (7%). In a Florida study, Green Treefrogs particularly ate larvae of lepidoptera (Spodoptera) and beetles (Chauliognathus), stink bugs (Euschistus), crawling flea beetles (Disonycha), spiders (Clubiona), and ants (Crematogaster); prey of 35 invertebrate families ­were included in the diet (Freed, 1982). The diet of the introduced population at Big Bend National Park consists of beetles, cockroaches, grasshoppers, crickets, ants, spiders, and even scorpions (Leavitt and Fitzgerald, 2009). Additional information on diet is contained in Haber (1926), Kilby (1945), and Brown (1974). Prey is selected based on size, activity, and frequency of occurrence (Freed, 1980a, 1982). Green Treefrogs have prey preferences, such as choosing flies (Musca) over mosquitoes; the more a prey item is likely to be active, the more likely it is to be eaten. Juveniles and adults have basically the same diet, but one attuned to the size of the frog. Green Treefrogs presumably locate invertebrates through the prey’s movement. In feeding ­trials, they did not use the acoustic signals of a potential prey, the cricket Achaeta domesticus, to guide their foraging movements (Höbel et al., 2014).

mus), and musk and mud turtles (Sternotherus, Kinosternon). Adult and juvenile Green Treefrogs are eaten by a wide variety of vertebrates and large spiders. Specific reports include spiders (Konvalina and Trauth, 2015, and references therein), killdeers (Schardien and Jackson, 1982), rat snakes (Pantherophis alleghaniensis) (Neill, 1951), black racers (Coluber constrictor), ribbon snakes (Thamnophis sauritus), ­water snakes (Nerodia sipedon), snapping turtles (Chelydra serpentina), herons, raccoons, and foxes (Wright, 1932; Mitchell and Anderson, 1994). Green Treefrogs are also attacked by the mosquito Uranotaenia lowii (Blosser and Lounibos, 2012). The best antipredator defense of postmetamorphic Green Treefrogs is their ability to blend in with vegetation and avoid diurnal movement over long periods. When disturbed, they may hop or jump away (a 4 g frog may hop 73 cm; John-­Alder et al., 1988), but their inner thighs do not contain the flash colors possessed by other hylids that might startle would-be predators. Contact with snakes may elicit body inflation (Marchisin and Anderson, 1978), and attacked individuals ­will use their limbs in an attempt to prevent ingestion (Höbel, 2011). ­There is no indication that their skin contains toxins or noxious substances that might deter predators. POPULATION BIOLOGY

Sexual maturity is reached in 1 year following metamorphosis (Garton and Brandon, 1975; Meshaka et al., 2020c). Meshaka et al. (2020c) reported that the smallest mature females yolking eggs ­were 32–33 mm SUL. Although the Green Treefrog is considered to be extremely abundant, ­there are no published data on population size. Literally thousands of Green Treefrogs can be heard in some areas calling on warm summer nights. In Louisiana, Pham et al. (2007)

PREDATION AND DEFENSE

Eggs have been killed by dense algal (Spirogyra-­like) mats in Mississippi (Tice et al., 2016). Larval predators include dragonfly naiads (Tramea, Anax), dytiscid larvae, notonectids, belostomatids (­giant ­water bugs), crayfish, fish (Centrarchus, Micropterus, Umbra, Fundulus), newts (Notophthal-

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Adult Dryophytes cinereus. Photo: Alan Cressler

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estimated ­there ­were 143 Dryophytes cinereus in a small urban office complex prior to metamorphosis. ­After metamorphosis, the population estimate was 446. COMMUNITY ECOL­O GY

Green Treefrogs breed l­ater in the season than many other species, and they prefer permanent bodies of ­water over temporary ponds. This may be ­because temporary ponds tend to build up a diverse array of predators as the season progresses, especially since temporary ponds lack fishes, which might other­wise keep many invertebrate predators in check. As such, variable predation pressure might be very impor­tant in determining both larval occupancy and reproductive success. Gunzburger and Travis (2005) showed that larval D. cinereus had low but constant survival rates when exposed to many types of predators and suggested that predation rate increased linearly with predator density. Not surprisingly, smaller tadpoles ­were more vulnerable to predators than medium to large tadpoles (Gunzburger and Travis, 2004), and tadpoles tended to decrease activity when exposed to potential predators (Anax, Lepomis, Notophthalmus) ­under experimental conditions (Richardson, 2001). In contrast to t­ hese results, Schiwitz et al. (2020) found no effect on larval activity in the presence of predator chemical cues or conspecific alarm cues. Green Treefrogs and Pine Barren Treefrogs have very similar calls, call from similar locations and microhabitats, and may occasionally be found at the same pond. Call rates of D. cinereus may be as fast as D. andersonii, but call durations and repetition rates are shorter in D. andersonii; lower peak frequencies are higher in D. andersonii compared with D. cinereus, but upper peak frequencies are lower in D. andersonii. Female D. cinereus are not attracted to calls of male D. andersonii. However, some female D. andersonii are attracted to the calls of male D. cinereus. Differences in female call discrimination capability could lead to interspecific hybridization if both species called from the same location (Gerhardt, 1974b). The presence of Rhinella marina larvae did not affect the growth, development, or survivorship of larval Dryophytes cinereus ­under experimental conditions (Smith, 2005a). DISEASES, PARASITES, AND MALFORMATIONS

The fungus Basidiobolus ranarum has been reported from Dryophytes cinereus in 6 of 14 animals examined in Missouri/Arkansas (Nickerson and Hutchison, 1971). Amphibian chytrid fungus (Bd) is reported from Alabama, Georgia, Mary­land, North Carolina, Oklahoma, Texas, and ­Virginia (Villamizar-­Gomez et al., 2016; Watters et al., 2016; Chiari et al., 2017; Marhanka et al., 2017; Tupper et al., 2017; Fuchs et al., 2018; Rivera et al., 2019; Lentz et al., 2021). However, Brannelly et al. (2012) found no evidence

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of Bd in wild populations from Louisiana, nor ­were they able to show any adverse effects of laboratory-­induced infection on body condition or growth of Green Treefrogs. They concluded that this species is not susceptible to Bd. Ranavirus is reported from D. cinereus in Georgia (Rivera et al., 2019), North Carolina (Lentz et al., 2021), and Oklahoma (Davis et al., 2019; also see Smith et al., 2019). An unidentified Myxidium (a myxozoan parasite of the gallbladder) was observed in tadpoles from the southeastern United States (Green and Dodd, 2007), and unidentified nematodes and protozoans (Opalina) have been reported from the gut (Haber, 1926). The trematodes Gyrodactylus sp. and Polystoma nearcticum have been observed on larvae (Green and Dodd, 2007) and adults (Brooks, 1979), and the nematode Cosmocercella haberi on adults (McAllister et al., 2008). Haber (1926) also reported ectoparasitic mites. Unidentified leeches ­were found on a D. cinereus in ­Virginia (Mitchell, 2012). Biting midges of the genus Corethrella feed upon D. cinereus and may be attracted by the frog’s advertisement call (McKeever, 1977; McKeever and French, 1991). Höbel and Slocum (2010) reported a Green Treefrog with malformed vocal and dorsal sacs. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. Aluminum at sublethal concentrations slows the growth rate of D. cinereus larvae, and the effect is more pronounced at a low pH (4.5) than at a higher pH (5.5). The LC50 (96 hr) is 277 μg/L at a pH of 4.5. Larval mortality increases with increasing aluminum concentration at low pHs, but not at higher pHs. High aluminum concentrations result in a reduced body size, which in turn makes larvae swim at a slower speed and increases the likelihood of predation (Jung and Jagoe, 1995). However, it is not just the smaller body size that ­causes a reduction in swimming speed; swim speed reduction is a direct result of the aluminum itself. Other contaminants. Green Treefrogs in a contaminated nuclear power fa­cil­i­ty outflow swamp had radiocesium levels at a mean of 204.2 pCi g-1 dry weight (Dapson and Kaplan, 1975). The biological half-­life of radiocesium is 30.1 days at 20–30°C. pH. Green Treefrogs are found in Carolina bays at pHs as low as 4.3 and in farm ponds with pHs of 7.9 (Jung and Jagoe, 1995). The lower limit of pH tolerance is 4.2, and Green Treefrogs are found commonly in ponds with a pH >4.5 (Eason and Fauth, 2001). Salinity. The mean LC50 value for early-­stage embryos is 5.2‰, 8.0‰ for late-­stage embryos, and 9.2‰ for larvae. Salinity is lethal at 8–12‰. Salinity levels affect body mass and larval period (see Larval Ecol­ogy), and may have sublethal effects for surviving larvae (Schriever, 2007). In experimental ­trials, larval D. cinereus had 100% survival at salinities ≤ 5 ppt. However, no larvae survived at salinities of

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14 or 16 ppt over a 72 hr period (Brown and Walls, 2013). Only 3.3% survived at 12 ppt. Chemicals. Bennett et al. (1983) reported no significant amounts of the pyrethroid insecticide fenvalerate in D. ­cinereus living in the vicinity of a cotton field sprayed 5 days previously. However, they only examined 1 frog. The insecticide carbaryl inhibits oviposition, but only when freshly dosed (Wilder and Welch, 2014). Petroleum crankcase oil had no effect on Green Treefrog hatching success, growth, or successful metamorphosis at concentrations of 0–55 mg/L (Mahaney, 1994). At 100 mg/L, tadpole growth was slowed considerably, although not at lower concentrations, and no successful metamorphosis occurred. However, oil concentrations decreased through time as experimental mesocosms received rainfall and oil was chemically and biologically degraded; thus, the adverse effects of intermediate concentrations of oil ­were masked by dilution. High oil concentrations also reduced the amount of floating algae, but not the overall algal standing crop. As a result, Mahaney (1994) suggested that Green Treefrog larvae would not metamorphose successfully at breeding sites contaminated by >50 mg/L of oil, since growth would be so delayed that winter low temperatures would inhibit thyroxin output and prevent complete metamorphosis. STATUS AND CONSERVATION

Dryophytes cinereus is a widespread and abundant species and, based on lit­er­a­ture accounts and laboratory experiments, McCoy et al. (2021) reported D. cinereus is not particularly sensitive to environmental change. Populations appear to be expanding in Illinois, the Piedmont of the South, in the South in general, and along the eastern seaboard (Platt et al., 1999; Rice et al., 2001; Friebele and Zambo, 2004; Florey and Mullin, 2005; Villena et al., 2016).

Habitat of Dryophytes cinereus. McIntosh County, Georgia. Photo: C.K. Dodd, Jr.

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They are common in rural areas and are able to persist in some abundance in disturbed (urbanized, silvicultural, agricultural) settings (Brown, 1974; Wilson and Porras, 1983; Delis et al., 1996; Surdick, 2005). For example, Pieterson et al. (2006) recorded a substantial increase in call detection over a 5 yr period in southwest Florida, despite intensive urbanization from 2000 to 2004. However, roads can cause significant mortality (Smith et al., 2005). Marsh et al. (2017) found that ­there was a negative relationship between the distribution of D. cinereus and primary/ secondary road density at a distance of 5 km, but that distribution was not affected by impervious surface cover. In contrast to population expansion in the north, Green Treefrogs may be displaced by the invasive Cuban Treefrog where they co-­occur in the Southeast, even in natu­ral protected areas, leading to extensive changes in the structure of treefrog communities (see Osteopilus septentrionalis account). They are eaten by this invasive species, out competed for refugia, and the tadpoles of Cuban Treefrogs outcompete Green Treefrog larvae for resources. Once Cuban Treefrogs move in, Green Treefrog abundance decreases. Changes in vegetation structure surrounding breeding ponds may lead to a greater potential for hybridization between D. cinereus and D. gratiosus. In par­tic­u­lar, a surrounding border of arboreal calling sites is necessary for Green Treefrog reproduction so that they do not come into contact with Barking Treefrogs; clearing vegetation from the shoreline and borders of ponds and lakes is not conducive to Green Treefrog reproduction. In the South, ­these frogs readily survive prescribed burns and recolonize burned wetlands, as long as retreat sites are available (Schurbon and Fauth, 2003; Langford et al., 2007). ­Because of their propensity for using cavities as retreats, the use of ground-­or tree-­placed PVC pipes has proven effective for monitoring this species and examining its movement patterns (Boughton et al., 2000; Zacharow et al., 2003; Waddle, 2006; Campbell et al., 2010). The lack of observations or captures of few of ­these frogs during extended surveys may reflect sampling biases (e.g., use of inappropriate techniques) rather than scarcity. Researchers in monitoring studies show that survivorship decreases by as much as 11% when 3 or 4 toes ­were clipped during mark-­ recapture studies, although toe-­clipping did not influence capture probability (Waddle, 2006). In terms of wetland mitigation mea­sures, Green Treefrogs readily colonize newly created ponds and wetlands (Merovich and Howard, 2000; Palis, 2007; Shulse et al., 2010; Drayer et al., 2020). Disturbances may or may not have significant impacts on populations of Green Treefrogs. Following Hurricanes Ivan and Katrina in Louisiana, Green Treefrog numbers plummeted in swamp, marsh, and levee habitats, presumably due

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to saltwater intrusion (Schriever et al., 2009). In contrast, populations can recover quickly in some coastal areas that receive hurricane overwash with resulting temporary increases in salinity (Gunzburger et al., 2010). In noncoastal habitats, canopy gaps may increase insect prey abundance, and Green Treefrogs may be more abundant in open-­ canopied situations than in closed-­canopy forest habitats (Horn et al., 2005). Presumably the frogs are in the open-­ canopy gaps ­because of better feeding opportunities.

Green Treefrogs may use restored wetlands. Mitchell (2016) found this species at 8% of agriculturally restored ponds in the Mid-­Atlantic region, and they have been found commonly in wetlands restored ­under the auspices of the Wetland Reserve Program (Walls et al., 2014a). Green Treefrogs have occasionally turned up in prepackaged salads sold in grocery stores (Hughes et al., 2019b). A total of 31,265 D. cinereus ­were exported from Florida from 1990 to 1994 for the pet trade (Enge, 2005a).

Dryophytes femoralis (Bosc, 1800) Pine Woods Treefrog

Alabama frogs, Brown (1956) gave a size range of 28.2– 40.5 mm SUL (mean 33.5). Larvae. Very small larvae are brownish yellow with a distinct lateral stripe and sharply bicolored tail musculature. As the tadpole grows, the dorsolateral body stripe is lost, and the tail stripe becomes more distinctive. The body assumes a dark olive to black coloration with a light (often bright yellow) venter. A pale postorbital stripe is pre­sent, but ­there is no interorbital bar. The tail musculature is distinctly striped, and the tail fins are flecked or blotched, although a clear area remains near the tail musculature. ­There is a well-­developed clear flagellum at the tip of the tail. Larvae exposed to certain predators develop orange-­red coloration on the tail fins and have deeper bodies and shorter tails than larvae not exposed to predators. Tadpoles reach about 22–38 mm TL (Brown, 1956; Jensen et al., 2008). Siekmann (1949) described the tadpole and Altig (1972a) described early development. Eggs. The eggs are brown dorsally and yellow ventrally. The vitellus is 0.8–1.2 mm in dia­meter, the inner envelope is 1.4–2 mm in dia­meter, and the outer envelope is 4–8 mm in dia­meter (Livezey and Wright, 1947; Delis, 2001). This outer envelope is loose, indistinct, and sticky. Eggs are deposited in a circular or elliptical mass 1.5–10 cm × 10–18 cm, which floats on the surface. Altig (1972a) noted that egg masses covered ca. 45 cm2.

ETYMOLOGY

femoralis: from the Latin femuralis meaning ‘pertaining to the hindleg.’ The name likely refers to the bold markings on the back of the thighs. NOMENCLATURE

Powell et al. (2016): Hyla femoralis Fouquette and Dubois (2014): Hyla (Dryophytes) femoralis Synonyms: Auletris femoralis, Calamita femoralis IDENTIFICATION

Adults. Pine Woods Treefrogs are mostly gray, tan, or reddish brown with distinct, bark-­like dark dorsal markings, with broad heads and short, rounded snouts. Some individuals adopt a greenish coloration when on a green background. Frogs are often lighter with more uniform markings at night than during the day, but ­there is considerable variation among individuals. ­There is a dark mark between the eyes that is variable in shape. A thin black line may be pre­sent on the upper jaw. Venters are white and unspotted. The rear of the thighs is marked by bright yellow-­orange spots. Toe pads are apparent but are not as con­spic­u­ous as in many other Dryophytes. Males have a dark vocal sac located more posteriorly than other Dryophytes; the sac forms a transverse fold pectorally when collapsed. According to Brown (1956), females are more reddish brown and males are more often gray than vice versa. Males are smaller than females (Delis, 2001); central Florida males are 21–36 mm SUL (mean 27.1 mm) and females are 22–38.5 mm SUL (mean 31.1 mm). In north Florida, frogs are 19–40 mm SUL (mean 32.1 mm) (O’Neill, 1995). South Florida males average 30.7 mm SUL and females 34.2 mm SUL (Duellman and Schwartz, 1958). For

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DISTRIBUTION

Dryophytes femoralis occurs on the Atlantic Coastal Plain from southeastern ­Virginia throughout much of Florida (to Miami-­Dade and Collier counties), then west to the Florida Parishes of Louisiana. Isolated populations occur in west central Alabama and in the Coosa Valley of central Alabama. ­There is a single, questionable rec­ord from Calvert County, Mary­land (Fowler and Orton, 1947; Cooper, 1953). The species occurs on Roanoke Island, North Carolina (Gaul and Mitchell, 2007), Harris Neck, Isle of Hope, St. Catherine’s, Sapelo, and Cumberland islands, Georgia (Martof, 1963; Laerm et al., 2000; Shoop and Ruckdeschel, 2006; Dodd and

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Moler (1986) reported a single hybrid between D. andersonii and D. femoralis from Florida. ADULT HABITAT

Distribution of Dryophytes femoralis

Barichivich, 2007; O’Hare and Madden, 2018), and Marco and St. George islands, Florida (Duellman and Schwartz, 1958; Irwin et al., 2001). Impor­tant distributional references include: Alabama (Mount, 1975; Redmond and Mount, 1975), Florida (Means and Simberloff, 1987; Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007), South Carolina (Dodd and Barichivich, 2017), and ­Virginia (Tobey, 1985; Mitchell and Reay, 1999). FOSSIL REC­O RD

Pleistocene fossils of D. femoralis are reported from northern Florida (Holman, 2003). Holman (2003) provided characters of the ilium that separate this species from other hylids. SYSTEMATICS AND GEOGRAPHIC VARIATION

Dryophytes femoralis is most closely related to D. squirellus, D. gratiosus, and D. cinereus (Hedges, 1986). In the laboratory, D. femoralis hybridizes successfully with D. arenicolor, D. cinereus, D. avivoca, D. gratiosus, D. squirellus, and D. chrysoscelis (Mecham, 1965; Fortman and Altig, 1973; Pierce, 1975), although success depends on the sex of the parents (Mecham, 1965). Fortman and Altig (1973) described the hybrid tadpoles. Jensen et al. (2008) reported natu­ral hybrids with D. chrysoscelis, and Anderson and

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As its name implies, the Pine Woods Treefrog is closely associated with the ­great pine forests that once covered the Southeastern Coastal Plain. ­Today the species is found in remnants of ­these habitats, as well as in mixed-­pine deciduous hardwood forests, including mesic and xeric hammocks, bottomland and slope forest, creek swamps, dome swamps, cypress bays, pine flatwoods, and sphagnum bogs (Carr, 1940a; Enge et al., 1996; Enge and Wood, 1998, 2000, 2001; Smith et al., 2006; Chandler et al., 2015a; Erwin et al., 2016). The unifying theme in the habitat is the presence of pine trees, although ­these frogs may be found on live oaks and magnolias (Harper, 1932). Occasional individuals are found among palmetto fronds. The species does not venture into the open wetlands favored by other Southeastern hylids, although it may occur in shrubby wetlands surrounding wet prairies. For example, Babbitt et al. (2005) found no D. femoralis in wetlands >150 m from the nearest woodlands. TERRESTRIAL ECOL­O GY

Pine Woods Treefrogs are active year-­round, weather permitting (O’Neill, 1995). This species is largely arboreal on pine and hardwood trees, although individuals may be found on the ground as they make their way to and from breeding sites. Movement patterns and distances traveled are largely unknown, but in Florida sandhills, Dodd (1996) found D. femoralis 42–815 m (mean 317 m) from the nearest ­water body. During emigration away from a pond, recent metamorphs tend to move directly ­toward the nearest surrounding forest (Dodd and Anderson, 2018). They frequent tree canopies, but ­little is known of their activity in high branches. Pine Woods Treefrogs take refuge in treeholes, cracks in trees, titi knotholes, and ­under bark (Harper, 1932). They also are found in the structures of old, dilapidated wood buildings—­such as cabins—­and in pine stumps. Harper (1932) speculated that only 1 frog occupied each tree, but this has not been confirmed. They sit with their limbs folded into the body to minimize moisture loss. Pine Woods Treefrogs engage in a complex series of movements whereby the limbs are used to wipe the head and body with an extraepidermal layer of mucous and lipids which helps to retard ­water loss (Barbeau and Lillywhite, 2005). Body wiping is impor­tant in allowing frogs to remain exposed in arboreal perches. As a result, evaporative ­water loss is significantly lower in D. femoralis than in many terrestrial anurans (Wygoda, 1984). Pine woods Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moon-

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light when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Pine Woods Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling normally occurs from April to September when air temperatures are >21°C, although advertisement calls have been heard as early as February in north and central Florida (Carr, 1940b; O’Neill, 1995; Delis, 2001). In south Florida, calling occurs from March to October (Duellman and Schwartz, 1958; Babbitt and Tanner, 2000; Meshaka and Layne, 2015), whereas in Georgia calls are heard from March to September (Jensen et al., 2008). ­There are 2 distinctive calls. One is made from the trees (to ca. 9 m in height) and occurs throughout the warm activity season, whereas the other is a more typical male advertisement call. Canopy calls occur both day and night, especially before, during, and ­after summer rain showers. At such times, the pine woods can erupt in choruses of D. femoralis. Males call from tree trunks at 1–4 m above the ground surface, in trees located in or around breeding pools; they may also call from tufts of grass near the shore. They wedge the rear part of their body into the tree bark and extend the front legs fully while calling. This allows expansion of the large vocal sac, which can be nearly as large as the male’s body. At breeding ponds, males produce an advertisement call from logs and floating debris, or from the pond margin on the ground. The advertisement call can be deafening, with Harper (1932) noting that one may find “the incessant din quite oppressive to the auditory organs.” Harper (1932) further noted that the “kek-­kek-­kek-­kek” call can be repeated for as long as 11 min without pausing. At the end of a series of “keks,” a male may even increase the rapidity of the call. The call has a dominant frequency of 4,800 cps, a duration of ca. 2.35 sec, and a repetition rate of 12.2 notes/ sec (Blair, 1958a). Large choruses (>50 calling males) are not uncommon (Delis, 2001). Most advertisement calling occurs at night. BREEDING SITES

Breeding occurs in temporary ponds, woodland pools, and isolated freshwater marshes scattered throughout the pine woods and uplands. Sites include cypress ponds, roadside ditches, rain pools in fields, branch and creek swamps, Carolina Bays, depression marshes, cypress savannas, and cypress-­gum ponds (Harper, 1932; Eason and Fauth, 2001; Greenberg and Tanner, 2005b; Liner et al., 2008). Fish should be absent, but they occasionally may be pre­sent (Holbrook

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Eggs of Dryophytes femoralis. Photo: Ronn Altig

and Dorn, 2016). In south Florida, breeding habitats are associated positively with the presence of nearby woodlands (within 20 m) and negatively with low pH and the presence of fish (Babbitt et al., 2006). Not all breeding sites may be used in any 1 year. For example, Babbitt and Tanner (2000) found D. femoralis tadpoles in 5 of 12 potential breeding sites in south Florida over a 21 month sampling period. According to W. Brode (in Neill, 1958), they breed in brackish ­water around Bay St. Louis, Mississippi. REPRODUCTION

Although calling can be prolonged, most breeding occurs in early summer, especially ­after heavy rainfall and during moderate temperatures. Eggs are deposited in a small surface film containing 17–68 or more eggs per mass (Wright, 1932; Altig, 1972a), or just below the ­water’s surface attached to vegetation or debris. A female deposits several masses, however, so the total fecundity was estimated at 500–800 eggs per female; this estimate was based on counts of single ovaries of only 2 females and extrapolated (Wright, 1932). Delis (2001) found clutch sizes of 205–1,948 (mean 924) in Florida and Mitchell and Pague (2014) reported clutch sizes of 701–2,086 (mean 1,290) in ­Virginia. Clutch size is not correlated with SUL, but it is positively correlated with clutch mass. Fertility is about 73%, with 97% of clutches showing at least some development (Delis, 2001). Eggs cannot survive dehydration, but they can withstand heat shock as high as 42°C; survivorship decreases at 8°C (Delis, 2001). Hatching occurs in 3 days. LARVAL ECOL­O GY

As with other hylids, the conditions faced by Pine Woods Treefrog larvae in breeding ponds have varied effects on

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The larval period is estimated at 40–70 days (Wright, 1932). The maximum tadpole size is 33–36 mm TL, and newly metamorphosed froglets are 11.5–14 mm SUL based on a sample size of 4 (Wright, 1932). Greenberg et al. (2017b) recorded emigration in August–­September and stated that juveniles ­were 40% of known hybrids could have been misclassified as pure parental species based on phenotype alone, had not ge­ne­tic analyses proved other­wise. However, ­there is no evidence that morphological asymmetry is greater in hybrids than in the parental species, suggesting that hybrids are not subject to decreased levels of developmental stability (Lamb et al., 1990). Hybrids may exhibit spectral variability in call characteristics, thus resembling ­either of the parental species, although the ranges of variation tend to overlap with D. cinereus (Gerhardt et al., 1980). Most introgressive hybridization occurs between male D. cinereus and female D. gratiosus. In the laboratory, D. gratiosus hybridizes successfully with D. chrysoscelis (Moore, 1955; Mecham, 1965; Fortman and Altig, 1973). Fortman and Altig (1973) described the hybrid tadpoles. Crosses with D. avivoca are generally unsuccessful (Mecham, 1965). Crosses with ♂ D. femoralis are unsuccessful, but the reciprocal cross is highly successful. Crosses with ♀ D. squirellus are unsuccessful, but the reciprocal cross is highly successful (Mecham, 1965). Individuals from south Florida do not have the prominent white line along the upper lip posterior to the tympanum that is seen in northern populations. ­There are a number of other minor color variations between south Florida frogs and their counter­parts in the Carolinas and Louisiana (Duellman and Schwartz, 1958). ADULT HABITAT

Dryophytes gratiosus is found in a variety of Coastal Plain and inland habitats, including pine flatwoods, longleaf pine

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sandhills, xeric hammock, mixed-­pine deciduous forest, and basin and depression marshes (Carr, 1940a; Enge and Wood, 1998, 2000, 2001; Chandler et al., 2015a). Habitats should contain temporary wetlands for breeding. They are occasionally found in suburban areas and along riparian stream corridors in habitats being developed (Barrett et al., 2016). TERRESTRIAL ECOL­O GY

During the day, adults normally are found in the surrounding forest, often in the canopy, and move to the margins of ponds and wetlands at dusk or ­after sunset. Males arrive before females. Individuals are frequently observed clinging vertically to vegetation surrounding a pond or wetland during the day. The limbs are tucked into the body to minimize moisture loss, and the eyes are closed tightly. Barking Treefrogs in such a position are often quite exposed to sunlight and the ele­ments. ­After finishing calling, males ­either return to the forest canopy or remain at the breeding site, presumably to forage. All movements to and from breeding ponds occur at night (Todd and Winne, 2006). Barking Treefrogs engage in a complex series of movements whereby the limbs are used to wipe the head and body with an extraepidermal layer of mucous and lipids, which helps retard ­water loss (Barbeau and Lillywhite, 2005). Body wiping is impor­tant in allowing frogs to remain exposed in arboreal perches. As a result, evaporative ­water loss is significantly lower in D. gratiosus than in many terrestrial anurans (Wygoda, 1984). ­After the breeding season, Barking Treefrogs seek shelter in dense forested habitats within proximity to wetland breeding sites. They take up residence in trees at heights >2.5 m above the ground, but ca. one-­third of the frogs are found buried under­ground in shallow pits or burrows covered by soil or pine ­needles; ­these individuals may have been found as they sheltered during postbreeding emigration. In a radio-­telemetry study in the late fall and winter, Delis et al. (2020) recorded daily movements of 0.4–17.2 m (range 0–287.5 m), although most movements ­were of short distance (19.5°C; warm rains stimulate extensive breeding activity. In northern Florida, Brugger (1984) found choruses from March to September, with most frogs calling from roadside ditches from May to September rather than from the semipermanent ponds that ­were frequented ­earlier in the season. In the spring prior to breeding (April in north Florida), males call from high in the trees (personal observation). Males also have a “rain call” that is made from the tree canopy throughout the warm months, especially during high humidity and precipitation; the species is sometimes referred to as the “rain frog.” The function of the rain call is not known. Males have a typical advertisement call made from the margins of breeding ponds and pools. Calling is primarily an aerobic activity and is energetically costly for a calling male. Calls are produced at the rate of 6,000/hr with a mean duration of 0.2 sec at dominant frequencies of 1,200 and 2,950–3,457 Hz (Blair, 1958a; Prestwich et al., 1989). The fundamental frequency is ca. 133 cps, and ­there is a large series of harmonics ranging up to 7,500 cps (Blair, 1958a). As with other

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hylids, call characteristics vary with temperature and are primarily aerobic (Brugger, 1984). The trunk musculature associated with calling makes up 10% of the total body mass. Calls are produced in an omnidirectional pattern. Calling D. squirellus males are territorial and have an aggressive encounter call if an intruder approaches too closely. The minimum distance between calling males is ca. 110 cm, and males are usually evenly spaced. The encounter call is similar to the advertisement call but has an increased repetition rate, made by shortening the interval between calls (Fellers, 1979a). If an intruder continues to approach, a wrestling match ensues. Satellite males may be nearby (usually within 50 cm; Brugger, 1984), but females bypass ­these males as they approach a calling male. The female initiates amplexus by touching the calling male (Brugger, 1984). Amplexus is axillary. Males and females may remain in amplexus on the bottom of a pond for more than 2 hrs prior to egg deposition, which takes 35–65 minutes to complete (Brugger, 1984). The SULs of amplexed pairs are not correlated, suggesting random mating rather than se­lection based on size.

REPRODUCTION

Breeding occurs throughout the warm spring and summer months, and males may amplex more than 1 female during the breeding season (Brugger, 1984). Mitchell (2014) reported that the smallest mature male on the Delmarva Peninsula was 27 mm SUL and the smallest gravid female was 31 mm SUL. Gravid females ­were found as late as 19 October on the Delmarva Peninsula (Mitchell, 2014). Eggs are deposited singly and are scattered on the substrate or pasted onto grass blades (Brugger, 1984). Other reports have eggs being deposited in a film or in small clumps (Wright, 1932). It might be that eggs are initially oviposited in a film or in small clumps that quickly break apart and submerge. Eggs may cling to one another. Wright (1932) reported clutches of 942 and 972 eggs, whereas Brugger (1984) recorded a mean of 1,059 eggs for 5 clutches observed in nature. In additional laboratory experiments, Brugger (1984)

BREEDING SITES

Breeding occurs in small temporary pools, ditches, and shallow, grassy wetlands in fields, and is associated with summer precipitation. Squirrel Treefrogs are frequently found in Carolina Bays, depression marshes, cypress savannas, pools in pastures, and cypress-­gum ponds (Babbitt et al., 2005; Liner et al., 2008). They are capable of breeding in very small pools, and almost any available small ­water body may contain larvae. Duellman and Schwartz (1958) even reported calling from a prairie adjacent to a marine canal. Open-­canopied wetlands are favored (Binckley and Resetarits, 2007). They do not normally breed in pools containing fish (e.g., Binckley and Resetarits, 2002), but when they do (usually only on nights with large numbers of breeding individuals; Binckley and Resetarits, 2008), larval densities are much lower than in pools without fish. In south Florida, breeding habitats are associated negatively with the presence of nearby woodlands (preferred 21–100 m) and positively with conductivity (Babbitt et al., 2006). Although fairly ubiquitous, not all breeding sites may be used in any 1 year. For example, Babbitt and Tanner (2000) found D. squirellus tadpoles in 11 of 12 potential breeding sites in south Florida over a 21 month sampling period. Frog populations calling from semipermanent ponds are larger than ­those calling from roadside ditches (Brugger, 1984). Brugger (1984) also found that about 25% of adults returned to the same breeding site in successive years, but that interpond movements ­were quite frequent.

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Eggs of Dryophytes squirellus, Alachua County, Florida. Photo: C. K. Dodd, Jr.

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272  Hylidae

Tadpole of Dryophytes squirellus. Photo: Steve Bennett

recorded a mean of 900 eggs per clutch (range 361–2,003). In ­Virginia, Mitchell and Pague (2014) gave a mean of 1,076 (range 392–2,081). Female body size may be correlated with clutch size. Hatching occurs in 36–48 hrs. LARVAL ECOL­O GY

Larvae of D. squirellus are often the most abundant tadpoles in small temporary pools. They are active foragers, even in the presence of invertebrate predators. Survival, however, is greater in habitats with complex vegetative structure than in less complex habitats (Babbitt and Tanner, 1997). Tadpoles become more difficult to find and predators have reduced foraging success when cover is available. In the presence of dragonfly (Anax, Pachydiplax, Tramea) nymphs, D. squirellus larvae decrease activity levels in laboratory ­trials, but in the presence of newts (Notophthalmus) or predatory fish (Lepomis), activity actually increases (Richardson, 2001; McCoy and Bowker, 2008). Larvae are likely to encounter dragonfly nymphs in their breeding ponds, but not the latter predators. Environmental conditions affect larval growth, size at metamorphosis, and the duration of the larval period. For D. squirellus, low temperatures during development increase the duration of the larval period but have no effect on size at metamorphosis (Blouin, 1992a). In general, however, D. squirellus larvae are as active, have slower growth rates, and metamorphose at smaller sizes than other temporary pond–­developing hylids (Richardson, 2001). Larvae are not infrequently found in coastal areas affected by salt spray or occasional overwash. According to W. Brode (in Neill, 1958), they breed in brackish ­water around Bay St. Louis, Mississippi. Webb (1965) observed tadpoles in a shallow pool with a salinity of 4.7 ppt in coastal North Carolina. The larval period is unknown, but possibly 40–50 days or more; newly transformed froglets are 11–15 mm TL (Wright, 1932; Mitchell, 2014).

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Adult Dryophytes squirellus, green phase. Photo: Aubrey Heupel

DIET

Dryophytes squirellus likely consume a wide variety of mostly arboreal insects. Duellman and Schwartz (1958) found beetles, small crayfish, a spider, an ant, and a cricket in a few south Florida Squirrel Treefrogs. Both males and females appear to feed during the breeding season (Brugger, 1984). PREDATION AND DEFENSE

This species is cryptically colored and blends in well with its background. The ability to change color allows it to blend in with a variety of background substrates. Predators likely include a wide range of vertebrates. Larvae are readily eaten by some native fish (Baber and Babbitt, 2003). POPULATION BIOLOGY

Breeding choruses can be small or large, depending on the breeding site used. Brugger (1984) found as many as 220 chorusing males in a single semipermanent pond, whereas roadside ditches contained 7–48 chorusing males. Brugger (1984) also found that operational sex ratios around a pond changed nightly, with more males arriving several days ­after a heavy rain. Immediately ­after a rain, the sex ratio was 2.6 males per female, whereas several days ­later it was 33 males per female. DISEASES, PARASITES, AND MALFORMATIONS

Biting midges of the genus Corethrella feed upon D. squirellus and may be attracted by the frog’s advertisement call

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(McKeever and French, 1991). It is also attacked by the mosquito Uranotaenia lowii (Blosser and Lounibos, 2012). An unidentified myxozoan parasite (Myxidium sp.) has been reported from Georgia (Green and Dodd, 2007). Squirrel Treefrogs are parasitized by the nematode Gyrinicola batrachiensis (Pryor and Greiner, 2004). As of 2007, amphibian chytrid had not been reported from this species (Rothermel et al., 2008), but it was found ­later in Georgia (Rivera et al., 2019). Ranavirus also was found in 1 individual from that state. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Salinity. In experimental t­ rials, larval D. squirellus had 100% survival at salinities ≤5 ppt. However, no larvae survived at salinities of 14 or 16 ppt over a 72 hr period (Brown and Walls, 2013). At 10 ppt, >80% of larvae survived. STATUS AND CONSERVATION

This species is widespread and common, and even may be expanding its range into the Piedmont and elsewhere (e.g., Pague and Mitchell, 1987). Based on lit­er­a­ture accounts and laboratory experiments, McCoy et al. (2021) reported that D. squirellus is not particularly sensitive to environmental change, although Villena et al. (2016) suggested populations in the South ­were declining based on probability of occurrence through time. The species tolerates urban, suburban, and agricultural areas fairly well (Delis et al., 1996; Hanlin et al., 2000; Surdick, 2005; Muenz et al., 2006), and is frequently seen at night feeding on insects while perched on buildings and lighted win­dows. They ­will use restored wetlands, such as ­those in the Wetland Reserve Program (Walls et al., 2014a). Still, Pieterson et al. (2006) noted declines from 2000 to 2004 based on call monitoring surveys in heavi­ly urbanized southwest Florida, and occupancy is reduced in silvicultural areas (Surdick, 2005). On Kiawah Island, South Carolina, Squirrel Treefrogs ­were found only in low development areas (Hanson and McElroy, 2015). Initial studies of the effects of well draw-­down zones in west central Florida noted ­little effect on tadpoles or call-­survey indices (Guzy et al., 2006). Squirrel Treefrogs may be displaced by the invasive Cuban Treefrog, even in natu­ral protected areas, leading to extensive changes in the structure of treefrog communities (see Osteopilus septentrionalis account). They are eaten by this invasive species,

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Breeding habitat of Dryophytes squirellus. Charlton County, Georgia. Photo: C.K. Dodd, Jr.

outcompeted for refugia, and the tadpoles of Cuban Treefrogs outcompete Squirrel Treefrog larvae for resources. Once Cuban Treefrogs move in, Squirrel Treefrog abundance decreases. Disturbances may have significant impacts on ­populations of Squirrel Treefrogs. Following Hurricane Dennis in Florida, Squirrel Treefrogs had not recolonized wetlands inundated with hurricane overwash 3 yrs following the storm (Gunzburger et al., 2010). However, Squirrel Treefrogs readily populate forest habitats managed by prescribed fire (Langford et al., 2007). Roads may impact Squirrel Treefrogs (Dodd et al., 2004), but the small size of this species makes it difficult to assess mortality, as carcasses are quickly obliterated or scavenged. Although numbers may change annually, only small numbers of D. squirellus may be captured in drift fences around breeding ponds (Gibbons and Bennett, 1974; Bennett et al., 1980; Enge and Marion, 1986; Dodd, 1992; Russell et al., 2002a; Greenberg and Tanner, 2005b; Baxley and Qualls, 2009), presumably ­because they can easily cross a fence. This species is better sampled using PVC pipes placed ­either in the ground or on nearby trees (Zacharow et al., 2003; Means and Franz, 2005; Campbell et al., 2010). Waddle (2006) found ­little effect of toe-­ clipping on ­either survival or capture probabilities of Squirrel Treefrogs. A total of 5,362 D. squirellus ­were exported from Florida from 1990 to 1994 for the pet trade (Enge, 2005a).

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274  Hylidae

Dryophytes versicolor (LeConte, 1825) Gray Treefrog Rainette versicolore ETYMOLOGY

versicolor: From the Latin word versi meaning ‘changing’ or ‘variable,’ and the Latin word color meaning ‘color.’ The name refers to the ability of individuals to change color. NOMENCLATURE

Powell et al. (2016): Hyla versicolor Fouquette and Dubois (2014): Hyla (Dryophytes) versicolor Synonyms: Calamita verrucosus, Dendrohyas versicolor, Dryophytes versicolor, Hyla phaeocrypta, Hyla richardii, Hyla verrucosa, Hyla versicolor phaeocrypta ­There are many publications where the identity of the species of “Gray Treefrog” is uncertain between D. versicolor and D. chrysoscelis (see D. chrysoscelis account for example; Roble, 1979). Therefore, it may not be pos­si­ble to identify the species involved in lit­er­a­ture discussions of Gray Treefrog biology, especially when ranges appear to overlap (e.g., Anderson and Arruda, 2006; Swanson et al., 2019). Many characteristics (morphology, calling season, be­hav­ior, some life history traits) may be applicable to both species. It is best to check the context of the original citation for confirmation. IDENTIFICATION

This species is morphologically identical to D. chrysoscelis. It can be differentiated by the following characteristics: chromosome number (D. versicolor is tetraploid [n = 48], whereas D. chrysoscelis is diploid [n = 24]); call rate (D. ­versicolor has a slow trill [17–35 notes/sec], whereas D. chrysoscelis is fast trilling [34–69 notes/sec], depending on temperature); and cytology (cells of D. versicolor are larger with more nucleoli than ­those of D. chrysoscelis) (Cash and Bogart, 1978). ­There are no morphological characters related to body proportions or coloration that accurately separate ­these species (Matson, 1990). Adults. The Gray Treefrog is a small to medium-­sized frog with a distinctive lichen-­like dorsal pattern; they appear to reach a larger size than their ­sister taxon D. chrysoscelis. The ground color is brownish to gray to greenish with a distinct lichen-­like pattern. The lichen coloration is composed of vari­ous gray to buff patches; sometimes the patches may be distinctively greenish. ­There is a con­spic­u­ous light patch or spot under­neath the eye, and ­there may be a dark crescent dorsally ­behind the eyes. All toes are tipped by con­spic­u­ous toe pads; a mucous layer is produced by the toe pad surface

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cells which ­causes the pads to become sticky and allows the frog to climb vertically (Green, 1981b). The rear toes are partially webbed, but ­there is only slight webbing between the fin­gers. Dorsally, the rear legs have a dark banding pattern. The inner thigh is bright yellow orange to orange and unspotted. Bellies are unpigmented, but the throat of a calling male is black. Males generally are smaller than females. Sizes also vary geo­graph­i­cally and perhaps from 1 pond to another. In Maine, adult males range in size from 50 to 54 mm SUL (­Sullivan and Hinshaw, 1992) with females to 60 mm SUL (Hunter et al., 1999); in ­Virginia, adults are 44–51 mm SUL (mean 47.7 mm) (Hoffman, 1946); in Connecticut, males are 36–51 mm SUL (mean 46.3 mm SUL) and females 43– 60 mm SUL (mean 49.9 mm SUL) (Klemens, 1993); in Pennsylvania, males are 35.9–56 mm SUL (means 40– 47.5 mm) and females 45.8–55.5 mm (mean 51.3 mm) (Meshaka et al., 2015d); and in Wisconsin, adults are 40–46 mm SUL (mean 42.6 mm SUL) (Jaslow and Vogt, 1977). Mean sizes of males in Ohio ranged from 43.2 to 46 mm SUL, and females 48.9–52.6 mm SUL, depending upon pond sampled (Gatz, 1981b). In Rhode Island, males averaged 46 mm SUL and females 48 mm SUL (Raithel, 2019). Gray Treefrogs change from dark to light (and vice versa) depending on temperature and light conditions. At night, melanophores in the skin contract, producing a light-­colored frog. As temperatures decrease or as ambient light increases, the melanophores expand, producing a darker-­colored frog (Edgren, 1954). Thus, a dark frog may change to a very light lichen-­colored pattern rather quickly (Babbitt, 1937). Metamorphs are usually green, gradually changing into the adult shades of gray and brown. The bright light spot ­under the eye is con­spic­u­ous and should aid in the identification of recent metamorphs. In coloration, they readily match the substrates surrounding a breeding site, such as grass, tree bark, or soil substrate. The skin may be completely smooth or granulated. As metamorphs age, ­there is less of a tendency to be green. Confusion over the color pattern may have led Allen (1868) to rec­ord this species as D. squirellus in Mas­sa­chu­setts, a place far from the distribution of the Squirrel Treefrog. Larvae. The description of the tadpole of D. versicolor is essentially the same as D. chrysoscelis; Altig (1970) does not differentiate the species. Gills are completely absorbed by 6 days ­after hatching. The general body coloration may be several shades of brown to olive green, with numerous, small, black and gold flecks scattered across the body and tail. The tail musculature does not have dorsal ­saddles. Tail fins may be colorless or blotched. ­There is a pair of preorbital

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stripes, but they are not distinctly outlined and only become apparent 2 weeks ­after hatching. The venter is immaculate anteriorly with some gold flecking, and the intestines may or may not be readily vis­i­ble through the body wall. As the tadpole grows, the venter posteriorly becomes more cream colored with heavy intrusions of gold flecks. The iris is bright gold or bronze. Maximum size is 42 mm TL. Descriptions of larvae and their mouthparts are in Hinckley (1880, 1881), Wright (1914, 1929), Babbitt (1937), Walker (1946), Gosner and Black (1957b), and Altig (1970). Hypomelanistic tadpoles from the Gray Treefrog complex have been reported from ­Virginia (Gibson et al., 2020). ­There is a unique color morph of D. versicolor tadpoles that appears only when predators are abundant at a breeding pond. Such tadpoles are longer with shallower bodies and develop bright red tail fins with dark margins. When predators, such as dragonfly larvae, are absent, the bright red coloration does not develop, and tail fins and bodies tend to be broader. The red coloration can be induced experimentally by raising larvae in the presence of predators (McCollum and Van Buskirk, 1996; Relyea, 2018). Bright tail fins are seen only on the oldest and largest larvae, usually within about 2 weeks of metamorphosis. Eggs. Most descriptions of the eggs of Gray Treefrogs do not differentiate between D. chrysoscelis and D. versicolor. ­There appears to be some variation in egg size and total clutch size, but ­whether ­these variables are species specific or reflect intraspecific variation is unknown. When deposited, eggs are drab in coloration and roughly 1.1–2 mm (mean 1.7 mm) in dia­meter (4–8 mm with the envelope, mean 5.2 mm). ­After a few hours, the vegetal pole becomes white to cream or yellow, the extent of which increases with development. ­There is only a single jelly capsule surrounding the egg (illustration in Tyler, 1994). Eggs may be oviposited singly or in small packets of 4–40 eggs along the surface of submerged vegetation (depth 0.2–0.4 cm) or at the edge of the breeding site. Packets may be separated by 15–30 cm (Wright, 1914; Livezey and Wright, 1947). The total egg complement per female is approximately 1,000–2,600 eggs. Hatching occurs from 2 to 5 days ­after deposition. Egg descriptions are in Wright (1914), Livezey and Wright (1947), and Tyler (1994). DISTRIBUTION

Determination of the distribution of this species is compounded by confusion with D. chrysoscelis. Many publications make no distinction between the 2 (e.g., Bragg, 1943a; Suzuki, 1951; Smith, 1961; Tobey, 1985; Dundee and Rossman, 1989; Hoberg and Gause, 1992; Phillips et al., 1999; Davis and Menze, 2000; Dixon, 2000; Hulse et al.,

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Distribution of Dryophytes versicolor

2001; Minton, 2001; Sievert and Sievert, 2006; Niemiller and Reynolds, 2011; Tipton et al., 2012), and ­later distributional publications often refer to the “Gray Treefrog complex” rather than differentiate between the 2 morphologically identical forms. In some places, Gray Treefrogs are not sympatric, whereas in ­others they overlap to a greater or lesser extent. In determining distribution, I generally followed the range map in Holloway et al. (2006). To be absolutely certain about identification, it is best to carefully listen to the calls (slow trills vs. fast trills), obtain a karyotype (diploid vs. tetraploid), or examine cell size (see above). The Gray Treefrog is found in Canada from western and southern New Brunswick and adjacent northern Maine, where it is expanding its range (McAlpine et al., 2009), and southern Québec and Ontario west to southern and western Manitoba and eastern Saskatchewan. One population is known from western Nova Scotia. Isolated populations appear in southern and west central Ontario. Pure populations of D. versicolor occur southward to southern New Jersey and Pennsylvania, west central ­Virginia, eastern West ­Virginia, central Ohio, Indiana, and Illinois westward to northeast Missouri, and in scattered areas in eastern Wisconsin north to northern Minnesota and southern Manitoba. Isolated D. versicolor populations occur in northwestern Kentucky and west Tennessee, western Wisconsin, northeastern Iowa, South Dakota (Kiesow, 2006), northwestern Arkansas, and east central Oklahoma. The species overlaps with D. chrysoscelis in a broad area from

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276  Hylidae

Mary­land and south central ­Virginia to southern Ohio, and westward in an arc all the way from eastern Kansas to east Texas and western Louisiana. Dryophytes versicolor also is found in Breckenridge, Hardin, and ­Meade counties, Kentucky (Burkett, 1989) and in Warren and Caswell counties, North Carolina (Dorcas et al., 2007). Additional mixed-­ species populations occur widely in eastern North Dakota, Wisconsin, Minnesota, and Iowa. In Wisconsin, D. chrysoscelis is more prevalent in the southeastern prairie regions of the state than D. versicolor (Vogt, 1981; Casper, 1998). The distribution pattern suggests that the polyploid D. versicolor may be adapted to colder and drier conditions (more northern, at higher elevations) than D. chrysoscelis, and adaptation to harsher conditions may have played a role in the evolution and establishment of polyploidy in this species complex (Otto et al., 2007a). Dryophytes versicolor may be able to tolerate more variation in climatic conditions, leaving most of the range of the progenitor species in more southerly areas of lower elevation, where climate extremes are not as common. The model used by Otto et al. (2007a), however, did not include northwestern populations of D. chrysoscelis, hence the applicability of the climate model origin may be ­limited to the northeastern tetraploid lineage of Holloway et al. (2006). Gray Treefrogs occur on the island of Ile Perrot, Québec (McCoy and Durden, 1965), Long Island, New York (Overton, 1914), Staten Island, New York, but nearly extirpated (Nicholls et al., 2017), the Elizabeth Islands of Mas­sa­chu­setts (Lazell, 1974), Walpole Island in Lake St. Clair (Woodliffe, 1989), islands at the Lake Ontario end of the St. Lawrence River, the Apostle Islands in Lake Michigan, Isle Royal in Lake Superior, and islands in Georgian Bay of Lake Huron (Hecnar et al., 2002). Hecnar et al. (2002) found them on ca. 23% of 107 islands surveyed. In New Hampshire, they occur from 75 to 195 m in elevation (Oliver and Bailey, 1939). Impor­tant distributional references include: Arkansas (Black and Dellinger, 1938; Trauth et al., 2004), Connecticut (Klemens, 1993; Klemens et al., 2021), Delaware (Zweifel, 1970a; Otto et al., 2007a), eastern Canada (Bleakney, 1958a; Logier and Toner, 1961), Illinois (Brown and Brown, 1972b), Iowa (Hemesath, 1998; Oberfoell and Christiansen, 2001), Kansas (Hillis et al., 1987; Collins, 1993; Collins et al., 2010), Kentucky (Burkett, 1989), Louisiana (Boundy and Carr, 2017), Maine (Hunter et al., 1999; Lindemann and O’Brien, 2019), Manitoba (Preston, 1982; Taylor, 2009), Mary­land (Noble and Hassler, 1936; Harris, 1975; Otto et al., 2007a), Mas­sa­chu­setts (Lazell, 1974), Michigan (Bogart and Jaslow, 1979), Minnesota (Oldfield and Moriarty, 1994; Moriarty and Hall, 2014), Missouri (Johnson, 2000; Daniel and Edmond, 2006), New Brunswick (Cox,

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1898, 1899; Bleakney, 1954; McAlpine, 1997a; McAlpine et al., 1980, 1991, 2009), New Hampshire (Oliver and Bailey, 1939; Taylor, 1993), New Jersey (Schwartz and Golden, 2002), New York (Gibbs et al., 2007), North Carolina (Dorcas et al., 2007), North Dakota (Wheeler and Wheeler, 1966), Ohio (Walker, 1946; ­Little et al., 1989; Pfingsten, 1998), Ontario (Johnson, 1989; MacCulloch, 2002; Weller, 2009; Choquette and Jolin, 2018), Pennsylvania (­Little et al., 1989), Québec (Bider and Matte, 1996; Desroches and Rodrigue, 2004), Rhode Island (Raithel, 2019), Saskatchewan (Taylor, 2009), South Dakota (Kiesow, 2006), Texas (Flury, 1951; Hardy, 1995), Vermont (Andrews, 2001, 2019), ­Virginia (Hoffman, 1946; ­Little, 1983; Mitchell and Reay, 1999; Otto et al., 2007a), West ­Virginia (­Little, 1983; Green and Pauley, 1987; ­Little et al., 1989), and Wisconsin (Jaslow and Vogt, 1977; Vogt, 1981; Casper, 1996; Mossman et al., 1998). FOSSIL REC­O RD

­ here is no way to differentiate members of the T D. versicolor/D. chrysoscelis complex osteologically, so ­there is no way to differentiate their fossils. Fossil members of the complex are found in Pleistocene (Irvingtonian) deposits from Nebraska and West ­Virginia. The complex also is found in Pleistocene (Rancholabrean) sites from Georgia, Kansas, Pennsylvania, and Texas (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

Previous suggestions that polyploidy in Gray Treefrogs was produced via a single speciation event among consistent progenitors are not supported based on a suite of analyses of molecular and call advertisement data. Dryophytes versicolor arose though polyploidy via several speciation events from D. chrysoscelis-­like diploid ancestors and 2 other now extinct lineages of treefrogs. According to Holloway et al. (2006), ­these 3 groups then freely interbred to produce the single species, D. versicolor. ­There are 4 distinct haplotype lineages, with D. avivoca also involved in the evolution of the tetraploid D. versicolor, although not in all 4 lineages. ­Today, however, gene flow is restricted between D. avivoca and the Gray Treefrog complex. Despite the dif­fer­ent origins of the lineages of tetraploid D. versicolor, the lineages are not reproductively isolated from one another. According to Keller and Gerhardt (2001), the reduction in pulse rate in D. versicolor is the direct consequence of ploidy level. Geographic variation in albumins (Maxson et al., 1977; Maxson and Maxson, 1978; Ralin, 1978), call characteristics (Gerhardt, 1974a; Ralin, 1968, 1977; Bogart and Jaslow, 1979), and allele frequencies (Ralin and Selander, 1979; Romano et al., 1987) undoubtedly stem from the dif­fer­ent evolutionary histories of the progenitor species as well as

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comparisons not corrected for temperature (Johnson, 1966). Members of the D. versicolor/D. chrysoscelis complex are most closely related to D. andersonii and D. avivoca (Hedges, 1986). Laboratory crosses between D. versicolor and D. chrysoscelis produce high rates of mortality (Johnson, 1959, 1963; Ralin, 1976). Crosses between female D. versicolor and male D. squirellus, Hyliola cadaverina, and Pseudacris fouquettei hatch and may produce a tiny percentage of metamorphs. Crosses between female D. versicolor and male D. cinereus, Pseudacris streckeri, P. clarkii, and Hyliola regilla do not reach metamorphosis; crosses between Dryophytes arenicolor or female D. cinereus and male D. versicolor are likewise unsuccessful (Pierce, 1975; Ralin, 1976). Flury (1951) documented much interpopulation variation in color pattern and morphometrics within the D. versicolor complex in Texas. However, ­there was no evidence of variation, even within dif­fer­ent biotic provinces, suggesting that dif­fer­ent taxa might be involved. ADULT HABITAT

Preferred habitats include southern mesic hardwoods, southern and northern lowland forest, boreal forest, and northern mesic and dry-­mesic hardwoods (Vogt, 1981). In the upper Midwest, D. versicolor is considered a forest species, whereas D. chrysoscelis is more associated with prairies, grassland, and oak savanna habitats (Vogt, 1981; Oldfield and Moriarty, 1994; Knutson et al., 2000). Price et al. (2004) suggested that this species preferred “highly irregular cover class patches dispersed throughout large areas of forest.” As Gray Treefrogs normally do not venture far from breeding ponds, habitat variables influencing distribution apply both to breeding and nonbreeding habitats. In much of their range, they prefer sites with forest cover nearby (>40 m from a breeding pond; Herrmann et al., 2005), but not necessarily directly surrounding a pond. Along the ­Great Lakes, for example, Gray Treefrogs are positively associated with the type of wetland and the relative area of grass and sedge cover, but negatively associated with the extent of forest cover within 100 m of breeding ponds; however, occupancy is positively associated with the amount of forest cover within 500 m of the pond. Within 3,000 m, ­there is a negative correlation between the amount of urban habitat and recreational grassland. In Ontario, Gray Treefrog abundance was negatively associated with mining activities, particularly in relation to the alteration of vegetative structure (Sasaki et al., 2015). Although pre­sent at mine tailing sites in Sudbury, Ontario, biomass and abundance was decreased in comparison with unaffected sites (Leduc et al., 2012).

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TERRESTRIAL ECOL­O GY

Gray Treefrogs grow quickly and show ­little growth ­after attaining adult sizes. Sexual maturity is reached in 2 yrs. According to Babbitt (1937), metamorphs grow to >25 mm SUL before undergoing their first overwintering season. However, Wright (1932) recorded them at 20–30 mm SUL at 1 yr of age (mean 25 mm), 30–41 mm SUL at 2 yrs of age (mean 35 mm), and 41–51 mm SUL (mean 45 mm) thereafter. Differences could reflect differences in growth rates. During the first season following metamorphosis, Gray Treefrogs prob­ably spend most of the time near the forest floor (Roble, 1979). As they get older, they ascend into the deciduous forest canopy where they spend most of the time in cracks, crevices, tree holes, woodpecker holes, and other types of cavities. Gray Treefrogs have been found as high as 20.1 m off the ground in pine trees in northern Wisconsin (Laughlin et al., 2017). They generally remain within ca. 200 m of the breeding pond, with most frogs living in close proximity to the pond; the maximum recorded distance a frog moved from a pond was 330 m in Missouri (Johnson, 2005). Females occupy areas farther out from the pond than males, and thus move over greater distances. Gray Treefrogs spend considerable time moving back and forth from breeding ponds to terrestrial/arboreal sites away from the pond, primarily to feed. The juxtaposition between forest cover and breeding pond is thus very impor­ tant in population per­sis­tence. In Missouri, Johnson (2005) found that ­there was significant variation in population ge­ne­tic structuring, such that populations >30 km apart ­were identifiable. ­Under 3 km, however, significant structuring was not evident. Johnson (2005) hypothesized that Gray Treefrogs ­were structured as classic metapopulations at regional distances, but that populations inhabiting a localized series of ponds formed a “patchy” metapopulation structure whereby some cohesiveness was established through interpond movement. Thus, populations are structured differently depending upon regional scale of distance and adjacent habitats. Gray Treefrogs become active in spring, with the timing dependent on weather conditions. In Arkansas and Texas, individuals have been observed in early March (Trauth et al., 1990; Perez et al., 2021), and it seems likely that they become active ­earlier when temperatures permit. The time they cease activity varies by latitude. In Mas­sa­chu­setts, New York, Ohio, Rhode Island, and Connecticut, for example, they are active ­until October (Hinckley, 1880; Wright, 1914; Walker, 1946; Klemens, 1993; Raithel, 2019). Dryophytes versicolor is active at temperatures of 15–34°C in New Jersey, with a CTmax of 38.7°C (John-­Alder et al., 1988). However, they call at temperatures as low as 8°C in Ontario, and it is probable that they are active at temperatures lower than 15°C in other parts of their range.

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Adults are able to survive freezing temperatures (5–7 days at -6°C) through the use of glycerol as a cryoprotectant (Storey and Storey, 1986), although subadults use both glycerol and glucose to survive freezing temperatures (Schmid, 1982; Storey and Storey, 1985). Costanzo et al. (1992) noted, however, that glucose was the sole cryoprotectant in Indiana populations of D. versicolor, so ­there may be some regional variation in the physiological means by which this species survives freezing temperatures. In addition, Gray Treefrogs mobilize glycerol differently from year to year (Layne and Stapleton, 2009). The body temperature stabilizes at 0°C with the heart continuing to beat at 5 beats per minute when the frog is frozen; the body temperature rises slowly in recovery, with an increase in heart rate as the body thaws (Layne and Lee, 1995). As much as 35–41.5% of the frog’s body ­water may be contained as ice at such temperatures (Schmid, 1982; Storey and Storey, 1985), although it may reach 50% depending on temperature, freeze conditions, and season (Layne and Lee, 1989). Gray Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Gray Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling in Gray Treefrogs begins in spring and may extend throughout the summer months; most reproduction occurs during the early part of the breeding season. On Cape Cod, the peak duration of the calling season lasts only 15 days (Cook et al., 2011). The timing of calling varies by latitude and environmental conditions. For example, calling occurs from late March to July in Rhode Island (Anonymous, 1918; Raithel, 2019), in early to late May in Mas­sa­chu­setts, Pennsylvania, and Ontario (Allen, 1868; Hinckley, 1880; Bertram and Berrill, 1997; Meshaka et al., 2015d), extends to June in Maine and Québec (­Sullivan and Hinshaw, 1992; Lepage et al., 1997), to July in Ontario and Wisconsin (Piersol, 1913; Bishop et al., 1997; Mossman et al., 1998), and to early August in New Hampshire and Pennsylvania (Oliver and Bailey, 1939; Meshaka et al., 2015d). In Texas, Gray Treefrogs call from March to September (Wiest, 1982; Perez et al., 2021). Some lit­er­a­ture rec­ords may confuse breeding calls with rain calls (see below). Males call from ground-­level or low, shrubby vegetation surrounding a pond at ambient temperatures ranging between 13–26°C in Missouri (Gerhardt, 1978a), although some males call from emergent vegetation within a pond. In

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Ontario, males did not call at ambient temperatures 4 months but not permanent) to long (permanent wetlands) duration (Herrmann et al., 2005). Not surprisingly, therefore, wetlands used for breeding by Gray Treefrogs usually contain high invertebrate richness and abundance (Babbitt et al., 2003). Treefrog presence tends to be negatively associated with the presence of predaceous fish,

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although they may be found inhabiting ponds containing fish (Babbitt et al., 2003; Baber et al., 2004), particularly nonpredatory fish (Hecnar, 1997; Hecnar and M’Closkey, 1997b). In experimental ­trials, predaceous fishes easily eliminated tadpoles of D. versicolor, and fish abundance was directly correlated with predation rate (Kurzava and Morin, 1998; Smith et al., 1999). When fish, such as the green sunfish (Lepomis cyanellus), are introduced to a breeding site, Gray Treefrogs tend to dis­appear (Sexton and Phillips, 1986). In Ontario, Hecnar and M’Closkey (1996b) found that Gray Treefrogs preferred sites with low conductivity. As with most anurans, not all available breeding sites are used in any 1 year, and occupancy can change from 1 year to the next (Hecnar and M’Closkey, 1996a). In New Hampshire, for example, 41% of the potential breeding ponds surveyed by Herrmann et al. (2005) ­were occupied by D. versicolor, whereas 15 min (Licht, 2003) and 80% of 2–4-­week-­old larvae ­after 5–7 days of exposure (Crump et al., 1999). The jelly envelope absorbs about 80% of the incoming UV radiation (280– 320 nm). UV light may interact with toxic substances to increase their lethal effects. Plant extracts. In mesocosm experiments, ­there was no effect on survivorship when larval D. versicolor ­were raised on Chinese tallow (Triadica sebifera) leaf litter instead of leaf litter from native tree species (Cotten et al., 2012). Gray Treefrogs had slight but increased growth in tallow litter compared with oak litter, but not in maple litter. Tadpoles ­were more developed at all mea­sure­ment stages when raised in tallow litter compared with oak or maple litter treatments, but not in Gosner stage in the overall treatment. STATUS AND CONSERVATION

Gray Treefrogs are widely distributed throughout their range, and most populations appear stable, with no evidence of decline at a landscape scale (Weller and Green, 1997; Mossman et al., 1998; Casper, 1998; Brodman et al., 2002; Brodman, 2003; Weir et al., 2014). A few reports suggest localized extinctions or declines or that the species is uncommon (Mierzwa, 1998; Choquette and Jolin, 2018). However, habitat loss has taken its toll on individual populations of the Gray Treefrog complex, and both species are susceptible to adverse effects from urbanization (Babbitt, 1937; Klemens, 1993; Lehtinen et al., 1999; Knutson et al., 2000; Picone, 2015), prolonged drought, and roads. For example, in Ontario, Gray Treefrog abundance is inversely related to the volume of traffic on roads within 200 m of breeding ponds (Eigenbrod et al., 2008, 2009). Other road mortality reports are in Ashley and Robinson (1996) and Cunnington et al. (2014); numbers reported killed are often small, prob­ably reflecting almost immediate obliteration rather than a small amount of mortality. At high road densities, D. versicolor/D. chrysoscelis populations are negatively impacted at small local

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Breeding habitat of Dryophytes versicolor. Photo: Jennifer Anderson-­Cruz

scales, but not over a more widespread regional scale (Marsh et al., 2017). Impervious surfaces do not seem to impact populations per se. Gray Treefrogs also may be adversely affected, at least on a short-­term basis, by natu­ral disturbances such as catastrophic flooding (Grant et al., 2015, 2018. Gray Treefrogs may persist in the vicinity of many types of adjacent land uses, even in fragmented habitats, as long as sufficient forested habitat remains around breeding sites. Gray Treefrogs also may survive provided that riparian buffer zones are maintained (Rudolph and Dickson, 1990). The abundance of members of the Gray Treefrog complex is reduced in agricultural vs. natu­ral habitats, but Gray Treefrogs still use ponds located in agricultural landscapes quite frequently (Knutson et al., 2004; Anderson and Arruda, 2006). For example, Kolozsvary and Swihart (1999) reported Gray Treefrogs to be ubiquitous in 30 forest patches of vari­ous sizes and degrees of isolation in the agricultural midwestern United States.

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288  Hylidae

Gray Treefrogs naturally occupy restored wetlands (Kline, 1998; Lehtinen and Galatowitsch, 2001; Foster et al., 2004; Brodman et al., 2006; Shulse et al., 2010; Baecher et al., 2018), such as ­those created at former mine sites (Myers and Klimstra, 1963; Lacki et al., 1992). In the Prairie Pothole Region, they occupy restored conservation grasslands (Ba­las et al., 2012). In addition, they may be candidates for inclusion in large-­scale restoration programs; Gray Treefrogs have been successfully translocated to areas within the Gateway National Recreation Area, mostly using larvae for release (Cook, 2008). Based on an analy­sis of movement patterns, protecting a 60 m area around the breeding site may allow Gray Treefrog populations to persist when surrounding habitats are disturbed (Johnson and Semlitsch, 2003; Johnson, 2005). Gray Treefrogs also occupy retention ponds in urban areas, although the proportion of impervious land surface is negatively correlated with occupancy (Simon et al., 2009). However, salts tend to accumulate in such ponds and make them less than optimal as breeding sites inasmuch as survival is negatively correlated with conductivity (Brand et al., 2010). Exposure to retention-­pond sediments reduces embryo survival but does not affect larval survival. Instead, surviving larvae tend to develop faster and reach larger sizes at metamorphosis (Brand et al., 2010). McCarthy and Lathrop (2011) noted that while calls ­were heard at retention ponds, successful breeding did not occur, perhaps ­because of the presence of fish. Gray Treefrogs tend to deposit more eggs in open-­canopied breeding sites than ­those located in forested or selectively cut habitats (Hocking and Semlitsch, 2007). Areas near a forest–­open area ecotone appear favored, even when the breeding site is located 50 m from the forest edge. Thus, this species tends to do well in areas that ­were previously clearcut, as long as source habitats are located nearby. Isolation diminishes the potential of wetlands to serve as good breeding sites in clearcuts. In silvicultural operations, it is best

to leave a mosaic of habitats in order to benefit Gray Treefrog populations, particularly wetlands located in open-­canopied sites near the ecotone of forest and clearcut. Gray Treefrogs also are found in power-­line rights-­of-­way, which provide a broad extent of ecotonal habitats (Fortin et al., 2004b). Traffic noise does not appear to inhibit calling by this species. Vargas-­Salinas et al. (2014) found that Gray Treefrogs, a species with a high call peak frequency, called randomly with regard to traffic noise intensity in eastern Ontario. When Gray Treefrogs call in the vicinity of roads with low volumes of traffic noise, their call rates are significantly lower than when they are in the presence of high volumes of traffic noise. Call amplitudes and dominant frequencies do not change with noise volume (Cunnington and Fahrig, 2010). Traffic noise also elicits an immediate response in calling frogs to lower the call rate, but ­there is no effect on call frequency or call amplitude compared with calls immediately before exposure to traffic noise. However, traffic noise does not interfere with mate attraction in this species (Cunnington and Fahrig, 2013). This plasticity in call characteristics allows the frogs to communicate acoustically despite the potential interference of traffic noise. Gray Treefrogs commonly are monitored using call surveys (Bishop et al., 1997; Bonin et al., 1997a; Lepage et al., 1997; Mossman et al., 1998; de Solla et al., 2005). As the number of sampling nights increases, the more likely it is that Gray Treefrogs ­will be detected. De Solla et al. (2005) determined that 10 sampling nights ­were needed in order to achieve detection probabilities >80% in Ontario, and Lepage et al. (1997) reported that it would take 334 listening stations to estimate presence at ±5% accuracy, and 1,716 stations to estimate abundance at ±10% accuracy. Based on call surveys over a 4 yr period (1986–1989), populations of the Gray Treefrog ­were considered stable in Illinois (Florey and Mullin, 2005); in upstate New York, they ­were considered to be increasing (Gibbs et al., 2005).

Dryophytes wrightorum (Taylor, 1939) Arizona Treefrog

Fouquette and Dubois (2014): Hyla (Dryophytes) wrightorum Stebbins (2003): Hyla eximia Synonyms: Hyla eximia [in part], Hyla eximia wrightorum, Hyla regilla wrightorum

ETYMOLOGY

wrightorum: This species is named in honor of Albert Hazen Wright (1879–1970) and Anna Allen Wright (1882–1964) in recognition of their pioneering work on frogs. NOMENCLATURE

McGinnis and Stebbins (2018): Hyla wrightorum

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IDENTIFICATION

Adults. The Arizona Treefrog is bright green or brown as an adult with a dark line extending from the snout through the eye onto the side of the body. The posterior half of the lower jaw is darkly pigmented, and the posterior side of the femur

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is lightly and evenly pigmented. The underside of the thigh and groin are orange or gold with a greenish tinge. Throats of males are dull greenish and tan, whereas female throats are white. At transformation, metamorphs are brown but change to a green coloration within a day or 2 ­after the tail is resorbed. Males averaged 41.3 mm SUL (range 37– 47 mm) in 1 Arizona study (­Sullivan, 1986c) and 37.3 mm SUL (31–45 mm) in another (Renaud, 1977). Chapel (1939) noted adults 25–48 mm SUL. Goldberg (2018g) recorded males from 31 to 41 mm SUL (mean 36.9 mm) and females from 36 to 44 mm SUL (mean 40 mm) based on museum specimens from Arizona. Larvae. Newly hatched larvae are yellow brown dorsally and yellow ventrally. Dusky pigmentation develops early on the body and tail fin. Mature larvae are brown and have a deep, globose belly, wide head, blunt snout, and lateral eyes; venters become dusky. The dorsal tail fin is low, and ­there are dark reticulations on both the posterior dorsal and ventral fins. Maximum length is 38 mm TL. Note that descriptions of the larvae in early publications (e.g., Wright and Wright, 1949) may include information pertaining to D. arenicolor. Zweifel (1961) provided figures of larvae at vari­ous stages of development and diagrams of the mouthparts. Eggs. The eggs have not been described but presumably are similar to ­those of D. arenicolor. DISTRIBUTION

Dryophytes wrightorum occurs in the mountains of central Arizona and western New Mexico. Isolated populations

occur in extreme southeastern Arizona (Huachuca Mountains and Canelo Hills) and in the Sierra Madre Occidental of northern México. A rec­ord from 37.7 km north-­northeast of Yuma likely represents an introduction (Vitt and Ohmart, 1978), and a rec­ord from Santa Fe, New Mexico (Van Denburgh, 1924; Wright and Wright, 1949), represented a point of shipment but not a collection locality. Impor­tant distributional references include: Van Denburgh (1924), Degenhardt et al. (1996), Gergus et al. (2004), Brennan and Holycross (2006), Bezy and Cole (2014), Murphy (2019), and Holycross et al. (2021). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

The Arizona Treefrog was described by Taylor (1938) as distinct from the Mexican species D. eximius to which it previously had been referred. It was ­later considered a subspecies of Hyliola regilla (as Hyla regilla, Jameson et al., 1966; Jameson and Richmond, 1971), but this arrangement is not supported by allozyme and mtDNA data and analyses of advertisement calls (Case et al., 1975; Gergus et al., 2004). ­There are no fixed allozymes among Dryophytes wrightorum populations, and the amount of ge­ne­tic variation among populations is small. Dryophytes wrightorum is a member of the D. eximius species group, a primarily tropical assemblage from México. The species is closely related to D. eximius (whose call is very similar) and less so to D. arenicolor (whose call is rather dif­fer­ent) (Bryson et al., 2010, 2014). Gene capture and mitochondrial introgression are common within the D. eximius species group. Dryophytes arenicolor populations from the ­Grand Canyon contain introgressed mtDNA identical with D. wrightorum (Bryson et al., 2014). ­Under laboratory conditions, crosses between members of the Gray Treefrog complex (D. chrysoscelis/versicolor) or D. squirellus and D. wrightorum ­were largely unsuccessful, although some eggs produced larvae, which then failed to develop. Many structural abnormalities ­were evident (Littlejohn, 1961a). Klymus et al. (2010) found evidence of past mtDNA introgression between D. wrightorum and D. arenicolor in the region of the ­Grand Canyon. ADULT HABITAT

Distribution of Dryophytes wrightorum

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Arizona Treefrogs are found in oak–­Ponderosa pine and spruce–­Douglas fir forests above 1,520 m (Chapel, 1939; Aitchison and Tomko, 1974). Other dominant trees include juniper, pinon, Mexican white pine, and white fir. The species occurs along small streams, in wet meadows and cienegas, and in roadside ditches. Williams and Chrapliwy (1958)

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290  Hylidae

Adult Dryophytes wrightorum, color variation. Photo: Cecil Schwalbe

heard them at 2,438 m in Coconino County, Arizona. Aitchison and Tomko (1974) observed them at elevations of 2,133–2,560 m near Flagstaff, and Degenhardt et al. (1996) recorded them at 2,000–2,750 m in New Mexico. Murphy (2019) gives a range of 900–2,900 m in Arizona. TERRESTRIAL ECOL­O GY

Arizona Treefrogs may be found in the surrounding forest before and ­after the breeding season (Chapel, 1939), but ­little is known of their movements. They are found both on the forest floor and in trees. Chapel (1939) noted 1 individual 23 m off the ground. Arizona Treefrogs are likely photopositive in their phototactic response as is D. eximius, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are likely sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Arizona Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

The call of D. wrightorum is rapidly pulsed and similar but of lower dominant frequency than that of D. squirellus from the eastern United States. It has been likened to a low-­ pitched metallic trill, but ­Sullivan (2019) noted that it is more like a rapidly pulsed “quack.” The advertisement call has a dominant frequency of 1,900–2,200 cps at air temperatures of 6–16°C; ­there are 100–120 pulses/sec with a mean duration of 0.15–0.17 sec (Blair, 1960b). In other studies (Renaud, 1977; ­Sullivan, 1986c), dominant frequencies w ­ ere 1,600–2,300 cps, pulse rates ­were 77.5–156 pulses/sec, and call durations ­were 0.12–0.24 sec. Dominant frequency is negatively correlated with male SUL (­Sullivan, 1986c). Gergus et al. (2004) noted that temperature exerted

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an impor­tant influence on call variables, and ­Sullivan (1986c) noted that Renaud’s (1977) inability to correlate pulse rate with body temperature was inconsistent with most studies on temperature effects. Whereas Gergus et al. (2004) found similar pulse rates to Blair (1960b), the dominant frequency dropped to near 1,600 cps. Choruses are not prolonged, but form ­after each rainfall event and last 2–4 nights. Chorusing occurs all night but is greatest before midnight. Although the size of the male often influences female choice in anurans, this does not appear to be the case in D. wrightorum. ­There are no significant differences in the SULs of mated and unmated males, nor is ­there a correlation between the size of males and females in amplexus (­Sullivan, 1986c). The species calls most often from the ground but has been heard calling from high in the trees in Arizona (Rorabaugh and Lemos Espinal, 2016). BREEDING SITES

Breeding sites mostly consist of large, shallow, grassy rainwater pools formed a­ fter summer rains, but they also breed in stock tanks and slow-­moving ephemeral streams that lack aquatic predators. Chapel (1939) noted that they congregated prior to breeding by lakes, ponds, streams, wells, and in any area likely to hold ­water once rainfall began. REPRODUCTION

Based on histology, Goldberg (2018g) noted that males and females are capable of breeding from June to August in Arizona. The smallest mature male was 31 mm SUL, whereas the smallest mature female was 36 mm SUL. Goldberg (2018g) also suggested that oviposition may occur more than once during the breeding season. Reproduction commences with the first summer rains from June to August, the exact date depending on weather conditions. Eggs are attached in small loose clusters to vegetation in shallow ­water. Eggs may be swept away ­after sudden rainstorms. ­There is no information available on clutch size or other aspects of reproduction. However, the mean clutch size of the closely related species D. eximius is reported as 851.1 eggs (range 508–1,476) (Hernández-­Salinas et al., 2018), so the clutch size of D. wrightorum may be similar. LARVAL ECOL­O GY

At hatching, larvae are 4.9–5.2 mm TL. They develop in shallow pools and have been found in streams, perhaps washed in from nearby pools ­after summer storms. Mortality occurs when w ­ ater evaporates. Larvae forage in vegetation in the warmest part of the pools, and Chapel (1939) noted that they aggregate to feed in ­great numbers around cow manure. However, Murphy (2019) reports larvae being observed from 21 June to 6 November, with metamorphs in

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September and October. The duration of the larval period is 6–11 weeks, and larvae grow to 38 mm TL (Rorabaugh and Lemos Espinal, 2016). Newly transformed froglets are 10–13 mm SUL. DIET

Very few data are available on diet. Chapel (1939) noted beetles, a spider, an earthworm, a fly, and “ips” (bark beetles—an insect pest) from a small number of individuals. PREDATION AND DEFENSE

This species is protected to some extent by its camouflage coloration. Predation of D. wrightorum has been recorded by American Bullfrogs, and bullfrogs are considered a threat to this species (Jones and Timmons, 2010). The species also has a noxious skin secretion that ­causes burning to the eyes and likely the mucous membranes (in Degenhardt et al., 1996). Postmetamorphs are eaten by Garter Snakes (Thamnophis cyrtopsis, T. elegans, T. eques; Drost, 2020; Jones and Hensley, 2020; Jones et al., 2020) and the larvae by Garter Snakes and predaceous aquatic insects.

Breeding habitat of Dryophytes wrightorum. Photo: Cecil Schwalbe

(Sredl and Collins, 1992). ­These species occasionally occur together in permanent ponds, and the larval periods overlap to the extent that large salamander larvae could cause significant predation on the small frog larvae. DISEASES, PARASITES, AND MALFORMATIONS

No information is available.

Arizona Treefrogs are parasitized by the cestode Cylindrotaenia americana and the nematodes Cosmocercella haberi and Physaloptera sp. (Goldberg et al., 1996c).

COMMUNITY ECOL­O GY

SUSCEPTIBILITY TO POTENTIAL STRESSORS

Dryophytes wrightorum and D. arenicolor frequently are found at the same breeding sites in the mountains of central Arizona. Differences in calls, breeding seasons (although they do overlap), habitats, and responses to rainfall normally are sufficient to prevent hybridization, although Klymus et al. (2010) found evidence of past introgression between them in at least 1 region. ­Under experimental conditions, larval salamanders (Ambystoma tigrinum) severely reduce survivorship of larval D. wrightorum, but had no effect on mass at metamorphosis, length of the larval period, or growth rates

No information is available.

POPULATION BIOLOGY

Hyliola cadaverina (Cope, 1866) California Treefrog ETYMOLOGY

cadaverina: from the Latin cadaver meaning ‘corpse,’ and -­ina meaning ‘having the appearance of.’ The name refers to the pale coloration of the species.

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STATUS AND CONSERVATION

Arizona Treefrogs in the Huachuca Mountains are in danger of extirpation and have been identified as needing urgent conservation management. Status and trends of other populations have not been reported. It is not known ­whether this species ­will use newly constructed ponds in terms of management, but playing conspecific chorus sounds does not increase the likelihood of colonizing artificial ponds (Buxton et al., 2018).

NOMENCLATURE

McGinnis and Stebbins (2018): Pseudacris cadaverina Fouquette and Dubois (2014): Pseudacris (Hyliola) cadaverina Synonyms: Hyla affinis [in part], Hyla arenicolor [in part], Hyla cadaverina, Hyla californiae, Hyla nebulosa [in part], Pseudacris cadaverina The name Hyla cadaverina was assigned to the species by Cope (1866) as a replacement name for Hyla nebulosa

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Hallowell, 1859, since the name H. nebulosa was already occupied by a South American frog. Gorman (1960) incorrectly stated that Cope (1866) used Hyla cadaverina as a replacement name for H. affinis and cited a publication title by Cope that is non­ex­is­tent. Gorman (1960) then described this species as Hyla californiae and noted confusion in the identification of specimens recorded as Hyla arenicolor from California in 1875. In the early lit­er­a­ture (e.g., Storer, 1925; Slevin, 1928; Stebbins, 1951), accounts of H. arenicolor (now Dryophytes arenicolor) may include intermingled information on H. cadaverina. IDENTIFICATION

Adults. This is a small to medium-­sized (to about 51 mm SUL) treefrog. The gray to sandy dorsum is slightly blotched or spotted, and the skin appears somewhat warty (more evenly spaced on males than females); ­there are no parallel dorsal stripes. This coloration blends well with the rocks on which the frog sits, gray being common in granite areas with sandy or light brown colors on sandstone backgrounds. This species does not have a black or brown lateral stripe through the eye. The toes have larger discs on the ends of the digits, and the webbing, while reduced from that of aquatic species, is more extensive than that of the Pacific Treefrog. Venters are white with the undersides of the legs, groin, and lower abdomen a lemon yellow. Males have a dusky or yellowish throat and are smaller than females. Females are larger than males. The mean male size ranged from 27.9 to 36.4 mm SUL at 26 populations; females averaged 34.5–42.4 mm SUL at 10 populations (Ball and Jameson, 1970). Frogs from xeric habitats are larger than ­those from mesic habitats (Ball and Jameson, 1970). ­These authors provided additional morphological mea­sure­ments. Goldberg (2017d) reported males 27– 42 mm SUL (mean 32.9 mm) and females 30–48 mm SUL (mean 38.4) from Riverside County, California. Larvae. Pigmentation is from light brown to dark brown. Golden flecks may be evident dorsally and laterally, and ­these may increase during ontogeny. Larval H. cadaverina are more elongate than larval H. regilla, and the tail fins are not as high. The greatest tail fin depth occurs well ­behind the anus, down about half or more of the length of the tail. The body tapers slightly from a point immediately ­behind the eyes ­toward the posterior part of the body when viewed dorsally, and it is more flattened than the body of H. regilla larvae. The dorsal tail fin tapers to a point at its tip, and the ventral tail fin is somewhat parallel to the body when viewed laterally. The eyes are small and entirely dorsally oriented when viewed from above. The dorsal part of the body is pigmented, but the venter and ventral tail fin are much less

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so, especially as the tadpole grows. When viewed dorsally, the tail musculature has regularly spaced transverse light bars or chevrons, and ­these become more pronounced with size. The dorsal tail fin is mottled. The tail musculature is heavi­ly pigmented, much more so than the dorsal tail fin. The intestines are vis­i­ble through the ventral body wall as the tadpole grows but may not be vis­i­ble during early growth stages. Gaudin (1964, 1965) provided descriptions of larvae and larval ontogeny and illustrations of larvae and mouthparts. Eggs. The eggs of H. cadaverina are bicolored, dark dorsally and white ventrally. Eggs are deposited singly, and each egg has a single gelatinous envelope. The vitellus is 1.8–2.4 mm in dia­meter (mean 2.07 mm), and the gelatinous envelope is 3.9–5 mm in dia­meter (mean 4.4 mm) (Storer, 1925; Gaudin, 1965). Although the eggs are deposited singly, they tend to stick together. DISTRIBUTION

The California Treefrog occurs from the lower Salinas River canyon and associated tributaries in central California to about 29°N latitude in Baja California, México. The eastern limits of its range are the western fringes of the Mojave Desert northwest of the Salton Sea. Impor­tant distributional references include Storer (1925), Gorman (1960), Lemm (2006), and Flaxington (2021).

Distribution of Hyliola cadaverina

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FOSSIL REC­O RD

­There are no fossils known for this species. SYSTEMATICS AND GEOGRAPHIC VARIATION

Although described as Hyla cadaverina, this species was transferred to the genus Pseudacris by Hedges (1986), based on an analy­sis of allozyme phylogeny. Hedges (1986) further recognized the distinctiveness of a western clade of Pseudacris that included P. regilla and P. cadaverina. Cocroft (1994) suggested that a generic reallocation was unnecessary based on morphological data, and recommended that cadaverina remain in Hyla. Da Silva (1997) used additional morphological information to return cadaverina to the genus Pseudacris. Moriarty and Cannatella (2004) ­later examined 12S and 16S mtDNA from 38 populations of chorus frogs from throughout North Amer­i­ca. They concluded that the California Treefrog was a member of the West Coast Chorus Frog clade, along with P. regilla. The recognition of a monophyletic West Coast clade is supported by immunological (Maxson and Wilson, 1975), allozyme (Hedges, 1986), morphological (Cocroft, 1994; Da Silva, 1997), nDNA, and mtDNA data (Moriarty and Cannatella, 2004; Barrow et al., 2014). An analy­sis of the ­family Hylidae by Duellman et al. (2016) further recognized the West Coast Chorus Frog clade of Pseudacris as a distinct evolutionary lineage and transferred both cadaverina and regilla to the genus Hyliola. The current distributional pattern of Hyliola cadaverina was established during the Pleistocene. The center of speciation was the eastern Transverse Ranges of southern California. ­There are 3 major haplotype groupings (northern, central, southern), with a major break within the Transverse Range, as populations ­were fragmented by mountains and watersheds. ­Limited amounts of gene flow are evident, with some desert populations having been isolated over a considerable period of time (Phillipsen and Metcalf, 2009). Hybridization is rare in nature, except perhaps in 1 general area. Brattstrom and Warren (1955) reported a pos­si­ble H. regilla × H. cadaverina hybrid from San Diego County, California. Gorman (1960) noted that hybrids seemed “especially plentiful” in central Los Angeles County. Phillipsen and Metcalf (2009) did not mention any evidence of hybridization. Laboratory crosses with Dryophytes arenicolor ­were not successful (Pierce, 1975). ADULT HABITAT

Hyliola cadaverina prefers shady, rocky areas with granite boulders adjacent to swift-­moving permanent streams and spring-­fed warm ­water ponds. It occupies rocks within the swift current, or forages among rocks and rock crevices immediately adjacent to shallow ­water. The banks bordering

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streams are often steep. This species is seldom found very far from ­water, although Cunningham (1955b) reported an adult 46 m from a stream down a rodent burrow and a juvenile 69 m from ­water. The species occurs from near sea level to 2,290 m (Lemm, 2006). TERRESTRIAL ECOL­O GY

Hyliola cadaverina is active both by day and by night, although ­there is conflicting information in the lit­er­a­ture about ­whether it is primarily diurnal or nocturnal (Cunningham, 1955b; Lemm, 2006). Activity occurs from mid-­February through early October, and activity is influenced by temperature and moisture. At some locations, individuals may be active year-­round to some extent. Populations near the California coast are more likely to be active for a longer period than are populations in the cooler high deserts. Individuals are tolerant of desiccation, and can lose up to 35% of their body ­water before succumbing to moisture loss (Cunningham, 1955b). They can also store up to 25% of their body weight in the form of dilute urine (McClanahan et al., 1994). California Treefrogs do not venture far from ­water, often sitting on boulders in the sun but within a quick jump of the ­water. While sunning, adult body temperatures may be considerably higher than ambient air and ­water temperatures. For example, Brattstrom (1963) recorded body temperatures of 20.8–26.2°C for frogs sitting in the direct sun when the ­water temperature was 21°C and the air temperature 19.8°C. California Treefrogs can maintain their body temperature at 12) pools and 20°C), they ­will move away from the pond to find cooler vegetation, move to areas where the ­water is cooler, such as streams fed by springs or seeps, or hide in cool cracks, crevices, or animal burrows. Hyliola regilla lose ­water slowly and can tolerate ca. 51% loss of body ­water; rehydration is rapid (Claussen, 1973a). Much activity of H. regilla during the nonbreeding season occurs nocturnally, as nighttime has more favorable temperatures and humidity than daytime. However, recent metamorphs disperse diurnally, and their body temperatures may reach 34°C. The metamorphs control their body temperature by moving to favorable microclimates and avoiding direct solar insolation when too warm, or by moving into sunny areas during cool weather. The CTmax is 36–37.7°C for adults, whereas the CTmin is -1 to -2°C (Cunningham and Mullally, 1956). Pacific Treefrogs are capable of using glucose as a cryoprotectant to survive bouts of freezing weather (Croes and Thomas, 2000). Freezing, especially in the fall, ­causes an increase in plasma glucose production from glycogen stored in the liver. As the winter season progresses, glycogen stores are depleted, thus reducing the effectiveness of responding to subfreezing temperatures. Pacific Treefrogs appear to be photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing

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illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Pacific Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males enter breeding sites as many as several weeks prior to females and may call in substantial choruses. At the beginning of the season, most calling occurs during the day. However, as the season progresses and temperatures rise, most chorusing occurs ­after dark. Then, males call in full chorus for up to 2 hrs; a second bout of strong chorusing may occur for 2 hrs ­after sunrise. Chorusing normally occurs at ­water temperatures of 8–19°C, despite reports suggesting calling does not occur below 9.8°C (Brattstrom and Warren, 1955). In fact, calls have been heard at ­water temperatures as low as 0–3.8°C (Cunningham and Mullally, 1956; Snyder and Jameson, 1965; Rombough and Trunk, 2019b) and air temperatures of -2.9–0.5°C (Schaub and Larsen, 1978; Rombough and Trunk, 2019b). Calling males even have been reported to sing from ­under deep snowbanks and in lakes nearly completely covered in ice (Grinnell and Storer, 1924; Grinnell et al., 1930). ­Water temperatures >20°C inhibit chorusing (Brattstrom and Warren, 1955). Calling often occurs in bouts; that is, a chorus ­will call for several minutes, be ­silent for several minutes, then start again. Calls may be heard at any time of the day, but peak chorusing occurs several hours ­after dark. At least some males are highly territorial and ­will return to a favored calling site if displaced (Perrill, 1984). Certain call characteristics, such as dominant frequencies, call repetition rates, and call duration vary significantly among populations (Snyder and Jameson, 1965); call characteristics in the south tend to be more variable than ­those in the north. ­These differences appear to correlate well with the dif­fer­ent phylogenies of the vari­ous populations, perhaps lending credence to taxonomic recognition. The characteristics of the mating call of H. regilla have been described by a number of authors (Snyder and Jameson, 1965; Ball and Jameson, 1966; Littlejohn, 1971; Allan, 1973; Straughan, 1975; Rose and Brenowitz, 2002), sometimes drawing dif­fer­ent conclusions from one another as to the distinctiveness of the call of this species from that of H. cadaverina. Littlejohn (1971) demonstrated, however, that ­there are differences in call duration, pulse repetition rate, and dominant frequency between ­these species, especially where they are sympatric. Indeed, Straughan (1975) suggested that pulse repetition rate was the only variable necessary for call discrimination. In H. regilla, the call duration averages 237.7 ms, the pulse repetition rate averages 66.2 pulses/sec, and the dominant frequency ranges from 2,350 to 2,650 Hz at 16°C (Littlejohn, 1971).

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Male Pacific Treefrogs have both a mate attractant call and an encounter call, that is, a call made upon the approach of a conspecific male. The advertisement call is of 2 types. In stable choruses in which the number and position of calling males is constant, they produce a diphasic advertisement call. The diphasic call consists of 9–13 pulses followed ­after 50 ms by 3–4 pulses. The monophasic advertisement call is a longer series of 20–25 pulses uttered at the approach of a female. Since females prefer a monophasic call to a diphasic call (Straughan, 1975), and in contrast to the conclusions of Allan (1973), males switch from a diphasic advertisement call to a monophasic call at the approach of a female. A second type of call is the encounter call. This is an aggressive call and serves to notify an intruder that he is encroaching upon a resident male (Awbrey, 1978; Whitney, 1980). Both the resident and the intruder may use the encounter call—­the resident to warn the intruder and the intruder to notify the resident of his intent to approach. If the intruder does not retreat, a fight (head-­butting, wrestling) between males may ensue. One function of the encounter call, therefore, is to maintain spacing, presumably preventing calling males from being too close to one another and thus reducing the chance of attracting a female (Whitney, 1980). A resident male ­will issue his encounter call when the intruder’s encounter call exceeds a certain amplitude. Presumably, the level of call amplitude provides a resident with information about the intruder’s distance. This helps to maintain calling male spacing patterns in stable choruses by notifying the intruder not to come closer. However, if chorus size increases, spacing patterns may change. A resident male may then respond differently to an intruder’s advertisement call by tolerating a higher amplitude than he would normally tolerate. The aggressive threshold for advertisement calls thus varies with the amplitude of the neighbor’s call (Rose and Brenowitz, 1991). Indeed, a resident may tolerate an amplitude 4 times that which he normally would, a response known as an accommodation. An aggressive response threshold (that is, the amplitude at which the encounter call is made) thus ­will differ between calling males in a dense population and ­those in a sparsely distributed chorus (Rose and Brenowitz, 2002). Such accommodation may occur rapidly. Plasticity in aggressive thresholds appears to affect responses to encounter calls much more strongly than responses to advertisement calls (Rose and Brenowitz, 1997). Pacific Treefrogs cannot use call duration to differentiate advertisement calls from encounter calls; if males did so, the advertisement call would sound similar to the encounter call, especially since both types of calls are uttered at the same rate of about 100 cps. Females also prefer the advertisement call to a male’s encounter call (Brenowitz and Rose, 1999). Therefore, the plasticity in male accommodation (that is,

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changing the threshold for aggression) allows males to maximize their time in producing advertisement calls and minimize time spent in aggressive encounters, especially in response to changes in male spacing patterns at a chorus. Males tend to space themselves at regular intervals around a breeding site. In British Columbia, Whitney (1980) recorded regular spacing by calling males about 50 cm apart, but males may call at 20–30 cm from one another when the density of calling males at a breeding site is greater. If additional males are added to a well-­established chorus, fewer males call than in a control situation (Whitney and Krebs, 1975b). This suggests that ­there may be a maximum density of chorusing males at a breeding site, at least for established choruses. If other males encroach within 50 cm, resident males issue the encounter call. Females approach a calling male, almost to the point of touching him, before he ceases calling and attempts amplexus. Females tend to prefer males that initiate calling bouts (Whitney and Krebs, 1975a) and males that are large (Benard, 2007). Certain males not only initiate calling bouts, but also tend to call longest within a calling bout; they also call at a faster rate, are louder, and are more likely to call outside the bout when other males are not calling. ­These cues provide a receptive female with extensive information about the fitness of the calling males. The male that can expend the most energy on calling is likely the most successful. However, the risk of predation is substantial during the mating season, and smaller males have a better chance of surviving attacks by ­giant ­water bugs than larger males (Benard, 2007). Thus, ­there is a trade-­off between female choice and predation risk with regard to male body size. Male body condition is not correlated with ­either mating success or predation risk (Benard, 2007). Males appear to adopt 1 of 3 strategies at a breeding site: become a calling male in a fixed location within the chorus; become a satellite male; become an opportunist and switch back and forth between calling and satellite status. Up to 17% of males at a chorus may be satellite males; that is, they do not call but position themselves near a calling male. In this way, they may be able to intercept a female moving ­toward the calling male. Satellites are occasionally successful (Perrill, 1984). Satellite males may become calling males, and ­these males may switch back and forth from calling to satellite from 1 night to the next (Perrill, 1984). ­These opportunistic males also may be successful in obtaining mates. Calling males may see satellite males as females and attempt to amplex them (Whitney, 1981). If a male is clasped by another male, he ­will give a trilling call that serves to identify his sex and ­causes the clasping male to release him (Brattstrom and Warren, 1955; Allan, 1973).

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Male Pacific Treefrogs sometimes call during off-­season rains or at other times during the nonbreeding season. Calls may be uttered from seasonal retreat sites when the frogs are nowhere near their normal breeding sites. Brattstrom and Warren (1955) observed males in January calling from small holes. The frogs would sing a few notes, then shimmy back down the hole to safety, only to emerge and call a few minutes ­later. The holes ­were about 50 cm in depth and ­were sometimes occupied by a Common Side-­blotched Lizard (Uta stansburiana). The purpose of ­these calls, if any, is unknown. BREEDING SITES

Pacific Chorus Frogs breed in slow-­moving streams (12°C and 28°C resulted in substantial embryonic mortality (mean 85%, range 70 to 97%), whereas embryos developing at 29°C in California experienced normal development with hatching successes of 80–100% (Brown, 1975a).

normally exposed to a wide range of temperatures daily and seasonally (Cunningham and Mullally, 1956). In one extreme, larvae ­were recorded in the desert in somewhat saline ­water at 33.4°C (Brues, 1932). Acclimation can raise the lethal temperature to 36°C. However, compared with desert and tropical species, larval H. regilla are not particularly heat resistant or tolerant. Aggregations of Pacific Treefrog larvae have been reported to form intermittently, such as Brattstrom and Warren’s (1955) observations of an aggregation of 150–180 tadpoles swimming in a circle about 61 cm in dia­meter. Most tadpoles faced in the same direction in ­water 7.5 cm in depth. Larvae do not show a preference for kin in forming an aggregation (O’Ha­ra and Blaustein, 1988). Such aggregations may serve thermal, feeding, or defensive functions. The larval period is 2–3 months (Jones et al., 2005), and tadpoles may reach 45–55 mm TL (Nussbaum et al., 1983). At metamorphosis, young Pacific Treefrogs are 11.2–16.5 mm SUL (Stebbins, 1951; Jameson, 1956; Gaudin, 1965).

LARVAL ECOL­O GY

Larvae are herbivorous, feeding particularly on filamentous green algae. In studies of growth rates, Kupferberg et al. (1994) found that larvae grew fastest on a diet of Cladophora (a filamentous green alga that has diatom epiphytes). Larvae grew at lower rates on filamentous green algae that did not have diatom epiphytes (e.g., Mougeotia), and slowest on flocculent detritus and ambient seston. Larvae fed other algae (Zygnema, Oedogonium) had between 40 and 53% survivorship compared with 85% for larvae fed Cladophora with epiphytes. Since body mass and length of the larval period are inversely correlated, a high-­quality diet reduces the amount of time spent as a tadpole. Kupferberg et al. (1994) suggested that alteration of streams by ­human activities could affect the resulting algal communities, which in turn affects the quality of the food available to anuran larvae. Larvae prefer the deeper ­water (to 1.2 m) of ponds and pond outflows to shallow ­water. Larvae can tolerate ­water to 34°C without acclimation (Brown, 1969), and they are

Tadpoles of Hyliola regilla. Photo: Breck Bartholomew

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DIET

Larval Pacific Treefrogs are omnivorous, feeding on filamentous algae and associated diatom epiphytes and bacteria suspended in the ­water. Adult Pacific Treefrogs eat a variety of small invertebrates, including beetles (larvae and adults), midges, tabanid flies, crane flies, true bugs, leafhoppers, ants, parasitic wasps, ichneumons, spiders, mites, pillbugs, isopods, snails, dragonfly naiads, and adult and larval flies (Needham, 1924; Tanner, 1931; Brattstrom and Warren, 1955; Johnson and Bury, 1965). Feeding occurs above ­water, and ­these small treefrogs may climb 1.2 m into bushes and branches as they forage. They also make use of floating vegetation as platforms for ambush, and ­will sit motionless in the shade of vegetation ­until an insect lands nearby. PREDATION AND DEFENSE

The eggs and larvae of Pacific Treefrogs are palatable and do not appear to have any chemical defenses (Licht, 1969a). They are preyed upon by backswimmers, predaceous ­water bugs, and salamander larvae (Ambystoma gracile, Taricha granulosa) (Peterson and Blaustein, 1991). It is not surprising, therefore, that this species generally avoids breeding in permanent ponds, especially ­those with fish. In experimental ­trials, for example, mosquitofish reduced tadpole survival significantly (Kerby et al., 2012), and their presence, as well as that of other nonnative fish or American Bullfrogs, was strongly negatively correlated with occupancy by H. regilla in natu­ral habitats (Preston et al., 2012). Larvae tend to form aggregations in response to the presence of chemicals emanating ­either from potential Garter Snake predators or from conspecifics attacked by Garter

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Snakes (DeVito et al., 1999). Presumably, ­these aggregations minimize the potential for any 1 tadpole to be eaten by the snake. Once metamorphosed, however, juveniles do not form aggregations (as do some toads), regardless of ­whether a predator is pre­sent. Larval H. regilla tend to avoid areas where conspecifics and even other anuran larvae have been attacked by predators such as predaceous diving beetles (Dytiscus). They presumably are able to detect larval damage-­release chemicals in the ­water. Adams and Claeson (1998) showed that larval Pacific Treefrogs avoided traps where damaged larvae ­were pre­sent, and that it made no difference ­whether the injured larvae ­were conspecifics or Rana. Adult and juvenile Pacific Treefrogs are cryptic on many types of substrates, blending especially with green and tan backgrounds in spring and with autumnal reds and russets in the summer and fall. Color morph frequencies change with seasonal changes in background color, suggesting se­lection ­toward seasonal substrate matching. The dorsal patterns of lines and spots further aid in crypsis. When disturbed by movement, Pacific Treefrogs ­will cease calling or jump away from a potential predator. Dill (1977) suggested a slightly leftward bias in jumping preference, with frogs generally jumping within a 70° arc from the frog’s initial bearing. Defensive be­hav­ior also may involve remaining motionless so as not to draw the attention of a predator. An adult was reported to become immobile in response to being poked while sitting on a cactus (Banta, 1974), and Brattstrom and Warren (1955) noted that if an adult was tossed into the ­water and landed on its back, it would remain motionless, with its limbs folded into the body, and float. A ­great many predators prob­ably prey on Pacific Treefrogs, including invertebrates, snakes, birds, and mammals. Specific references include: ­giant ­water bugs (Benard, 2007), American Bullfrogs (Lithobates catesbeianus), Belted Kingfishers (Megaceryle alcyon), herons, egrets, Mallards (Anas platyrhynchos), Red-­tailed Hawks, robins, Garter Snakes (Thamnophis elegans, T. sirtalis), raccoons, skunks, opossums, river otters, domestic ­house cats (Felis sylvestris), and dogs (Canis familiaris) (Storer, 1925; Grinnell et al., 1930; Arnold and Wassersug, 1978; Nussbaum et al., 1983; Jennings et al., 1992; Weitzel and Panik, 1993; Werner et al., 2004; Rombough and Bradley, 2010; Rombough and Trunk, 2019a). In addition to native predators, introduced American Bullfrogs, mosquitofish (Gambusia affinis), and other nonnative fish readily consume Pacific Treefrog larvae and may have an effect on stream-­dwelling populations (Goodsell and Kats, 1999; Kerby et al., 2012; Preston et al., 2012). Larvae also are eaten by predaceous aquatic insects, other anurans (Rana muscosa), and Garter Snakes (T. gigas) (Pope, 1999; Ersan et al., 2020).

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POPULATION BIOLOGY

Adult population size varies from a few dozen to several hundred individuals at a breeding site. For example, Weitzel and Panik (1993) estimated an adult annual breeding population of 60 at a small pond in northwestern Nevada, and Schaub and Larsen (1978) estimated the size of the male breeding population at study sites in Idaho as 360 individuals 1 year and 160 the next. Jameson (1957) estimated ­there ­were 484 adult males at a pond in Oregon in April. At a pond in California, Jameson and Pequegnat (1971) estimated a population size of about 322 frogs in the first year (range 245–435 as the season progressed) and 366 during the second year (313–435 as the season progressed) of a 2 yr study. Most H. regilla mate only once in their lifetime, but a small percentage return to a breeding site in more than 1 year. Jameson and Pequegnat (1971) estimated an annual mortality rate of 90%, making this species essentially an annual species with a short life-­span (3 years or less, Jameson, 1956). A total of 38 of 373 (10.2%) marked frogs ­were recaptured 1 year ­later at Jameson’s (1957) Oregon site, and Schaub and Larsen (1978) recaptured 13.8% of adults marked 1 year during the following year. Jones et al. (2005) reported that most Pacific Treefrogs do not reach maturity ­until their second year (presumably the first spring ­after metamorphosis). Males prob­ably remain at a breeding site for days to weeks, although perhaps not throughout the entire breeding season. In Idaho, Schaub and Larson (1978) recorded male stays from an average of 10–13 days, depending upon year surveyed. Hyliola regilla males may mate with multiple females, and Perrill (1984) recorded successful amplexus by males on consecutive nights. Even ­after mating with several

Adult Hyliola regilla. Photo: Aubrey Heupel

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females, males ­will continue calling throughout the peak breeding season. Females may remain only as long as it takes to oviposit their complement of eggs. The size of the male does not appear to influence mating success, and ­there is no size-­assortative mating between males and females. This might be expected of a species that normally has only 1 chance at reproduction and for which size differences are not very ­great between males and females. For ­those frogs that do breed more than once, subsequent breeding may take place at a dif­fer­ent pond than the pond originally selected (Storer, 1925). Since the availability of breeding sites varies annually, a lack of site philopatry allows the frogs to breed at what­ever sites are available. It seems probable that long-­term per­sis­tence at any 1 location is precarious, as environmental perturbations could easily eliminate populations, at least temporarily. A cycle of per­sis­tence for a number of years followed by temporary extirpation appears to have occurred at an isolated population in northwestern Nevada, where successful reproduction occurred during 12 of 15 years the population was studied (Weitzel and Panik, 1993). In the other 3 years, the population was wiped out by severe flooding, only to be rapidly recolonized. Indeed, the cycle of successful reproduction–­ extirpation appears to have been repeated frequently over a period of 84 years due to flooding, desiccation, and extreme fluctuations in ­water temperature. The red-­green color frequencies of the metamorphs do not match the color frequencies of ­either the parental population or the population that breeds the following year. ­These results suggest that polymorphism is maintained by differential se­lection among color types and that this se­lection pressure varies seasonally and by location (Jameson and Pequegnat, 1971), perhaps in much the same way color morph frequencies vary in Boreal Chorus Frogs. Jameson and Pequegnat (1971) further speculated that the plasticity in color morph frequencies among and within populations allowed adaptation to rapidly changing environmental conditions. COMMUNITY ECOL­O GY

Pacific and California Treefrogs occupy the same general regions and stream drainages but tend to partition the habitat according to specific microenvironmental preferences (e.g., along slow-­moving vs. fast-­moving ­waters). However, as drought conditions cause habitats to dry in summer, individuals of ­these 2 species may be brought into close contact. Despite this, ­there is ­little evidence of hybridization between them, even though they may si­mul­ta­neously use the same ­waters for reproduction. Despite assertions in Ball and Jameson (1966), differences in call characteristics provide a premating isolating mechanism to prevent introgression between ­these closely related taxa.

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Pacific Treefrogs appear to be impor­tant to the distribution of garter snakes, particularly Thamnophis elegans. Indeed, the presence of H. regilla is a major indicator of the presence of this snake (Knapp, 2005). Anurans are a major portion of the diet of T. elegans, and snakes appear to congregate along streams and ponds when metamorph Pacific Treefrogs are beginning their dispersal (Arnold and Wassersug, 1978). The distribution of and declines in garter snake populations may be correlated with the presence or absence of H. regilla populations. Garter snakes also may have a secondary effect on the activity and be­hav­ior of larval Pacific Treefrogs. When given a choice, larvae feed preferentially on the green alga Cladophora, which may contain diatom epiphytes, rather than lower nutritional quality filamentous algae such as Mougeotia, which does not contain epiphytes. However, larvae switch to Mougeotia in the presence of garter snakes. This is ­because Mougeotia forms a floating cloud-­like structure with numerous hiding locations, whereas Cladophora forms a much more exposed floating mat. If larvae are forced to remain in the Mougeotia, their growth rate declines and the larval period may increase ­because of the decrease in available nutritional resources (Kupferberg, 1997b). Larval life history characteristics can also be affected by other anuran larvae. In a California river, for example, the presence of American Bullfrog larvae decreased the size at metamorphosis of Pacific Treefrog larvae by 16% (Kupferberg, 1997a, 1997b). Kupferberg (1997a) attributed the effect to competition for algae between ­these species. The decreased size at metamorphosis in turn could have adverse impacts on this species, as fitness may be a function of size at metamorphosis. Thus, even though tadpole survivorship was not affected, larvae of the nonindigenous American Bullfrog could have long-­term indirect detrimental impacts on populations of Pacific Treefrogs. Despite the results from Kupferberg et al. (1994) and Kupferberg (1997b), Adams (2000) and Pearl et al. (2005b) could not demonstrate any direct effects of American Bullfrogs on Pacific Treefrog survivorship or presence in the Pacific Northwest. They noted that Pacific Treefrogs ­were virtually absent from permanent ponds, but attributed their absence to the presence of predaceous exotic sunfish in the permanent habitats. Adams (2000) suggested that gradients in habitat characteristics may be more impor­tant than broad-­scale characterizations in assessing the effects of nonindigenous species on distributional patterns. Still, indirect effects may play a critical role by altering food availability, access, and be­hav­ior, as noted by Kupferberg (1997b). For example, Preston et al. (2012) found that the presence of bullfrogs decreased occupancy by H. regilla and reduced larval growth rates, but not survivorship.

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Many species of anuran larvae are able to detect the presence of potential predators within their environment. Pacific Treefrog larvae, for example, can detect chemicals emanating from the native Redside Shiner (Richardsonius balteatus) and reduce their activity accordingly. Tadpoles can also detect odors of nonnative rainbow trout and, even when exposed to trout odors as embryos, grow more slowly and take refuge more often than controls (Garcia et al., 2019). They are able to detect alarm chemicals from injured hetero-­ and conspecifics and avoid them, presumably reducing their chance of a predatory encounter (Adams and Claeson, 1998), and are more likely to take refuge than controls even ­after being exposed to ­these cues only as embryos (Garcia et al., 2019). However, they do not respond to chemicals from certain nonnative fish (Lepomis) or crayfish (Procambarus) (Pearl et al., 2003). This inability to detect nonindigenous predators exposes them to predation and may help to limit their distribution to temporary wetlands. It also helps explain why the introduction of nonindigenous species has a detrimental effect on the treefrogs. This is not to say that Pacific Treefrogs do not respond to introduced predators in other ways, however. Larval abundance and egg mass counts are lower in streams possessing nonindigenous crayfish (Riley et al., 2005), and Pease and Wayne (2014) noted that tadpoles that inhabited streams in California where the crayfish Procambarus clarkii was introduced had shallower and narrower tails than tadpoles from streams without this predator. In contrast to the results above, Pease and Wayne (2014) found that tadpoles from both crayfish-­inhabited and non-­inhabited streams ­were less active than normal when exposed to crayfish chemical cues. However, tadpoles with deeper tails survived better than tadpoles with narrow tails in laboratory ­trials, a seemingly contradictory result from field observations. Narrow tails may have other as yet unmea­sured advantages in natu­ral populations. The presence of grazing H. regilla tadpoles may have effects on the periphyton community within a wetland, at least for the duration of the developmental period. In field experiments, Kupferberg (1997c) demonstrated that larval Pacific Treefrogs could cause as much as an 18% decrease in area-­specific primary production due to grazing on periphyton. Grazing had no effect on periphyton biomass-­specific productivity. Community effects ­were based on consumption, rather than through the recycling of nutrients. The interplay between H. regilla and its parasite Ribeiroia ondatrae is complex. The presence of other predators of the parasite (mollusks, zooplankton, fish, larval insects, newts) can reduce the infectious stage of this trematode by 62–93% (Orlofske et al., 2012). Both newts (Taricha) and damselfly larvae reduced infection of H. regilla tadpoles by ca. 50%, ­either by direct predation or by serving as alternate hosts.

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Mosquitofish, on the other hand, are predators of the parasite and may adversely impact tadpoles, but do not serve as hosts for the parasite. Thus, host-­parasite interactions in natu­ral habitats have impor­tant implications for pathogen transmission and population effects. Pacific Treefrogs are found in the same habitats as other frogs, including Northern Red-­legged Frogs. An unusual form of mortality was reported by Rombough (2020) whereby adult male H. regilla apparently became trapped in the jelly of R. aurora egg masses and drowned. Dead frogs ­were found on the pond bottom covered by ­water mold, but this occurred only ­after death and was not responsible for the mortality. DISEASES, PARASITES, AND MALFORMATIONS

A few morbidity events have been recorded for the Pacific Treefrog. Ranavirus is known to have caused morbidity in northern Idaho (Russell et al., 2011). Green et al. (2002) recorded a hypopigmentation and gigantism event in Oregon that involved about 5% of the observed larvae of Pacific Treefrogs and Western Toads. Another morbidity event involved dermal metacercariae and possibly Bd affecting 4 larval species. In neither report is it pos­si­ble to discern ­whether ­these abnormalities or pathogens affected all the species listed from the par­tic­u­lar event. Batrachochytrium dendrobatidis (Bd) was found in museum specimens collected from California from the 1970s to the 2000s (Padgett-­Flohr and Hopkins, 2009), and is pre­sent in this species at both montane and coastal sites in California (Fellers et al., 2011; Piovia-­Scott et al., 2011; Ecoclub Amphibian Group et al., 2016) and in Washington (Gaulke et al., 2011). Larvae experimentally infected with Bd show no evidence of behavioral fever or altered thermoregulation (Han et al., 2008), and indeed seemed ­little affected by the pathogen (Blaustein et al., 2005b). However, high and fluctuating temperatures have negative effects on the survival of this species when exposed to Bd (Rumschlag et al., 2014). In another study, Bd infection resulted in negative impacts on tadpole activity (reduced by 43%), survival to metamorphosis (by 8%), time to metamorphosis (by 2.5 days), and time of tail absorption (by 0.5 days), with marginal effects on mass at metamorphosis (Kleinhenz et al., 2012). However, recent metamorphs from ­these experimental Bd-­inoculated larval ­trials did not test positive for Bd. In experimental ­trials, Bd had no interactive effects with sublethal pesticide mixtures (involving atrazine, acetochlor, glyphosate, 2,4-­D, carbaryl, chlorpyrifos, endosulfan, permethrin) on survivorship or on infection loads, and Bd did not affect larval survival in the absence of the pesticide mixture (Jones et al., 2017; but see Kleinhenz et al., 2012). Parasites include the trematodes Alaria sp., A. mustelae, Clinosternum sp., Distoichometra bufonis, and Megalodiscus

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microphagus (Goldberg and Bursey, 2001b, 2002a; Zamparo and Brooks, 2005); the cestode Distoichometra bufonis (Koller and Gaudin, 1977); the nematodes Cosmocercoides variabilis, Oswaldocruzia pipiens, Physaloptera sp., Rhabdias sp., and R. ranae (Koller and Gaudin, 1977; Goldberg and Bursey, 2001b, 2002a); and the acanthocephalid Centrorhynchus californicus (Millzner, 1924). Limb malformations have been reported commonly in Pacific Treefrogs. Both field and laboratory studies have linked ­these malformations with interference in development by the dige­ne­tic trematode Ribeiroia ondatrae (Sessions and Ruth, 1990; Johnson et al., 1999, 2002; Green et al. 2002; Blaustein and Johnson, 2003; Johnson and Sutherland, 2003; Lannoo, 2008; Roberts and Dickinson, 2012). Malformations include missing limbs (ectromely) and digits (ectrodactyly), extra digits (polydactyly) and limbs (polymely), femoral projections, bony triangles (taumely), missing eyes (anophthalmy), abnormal jaws (mandibular hypoplasia), unusual skin webbings, and ­others. Hind limbs are affected far more often than forelimbs. In heavy experimental infestations, nearly 3 abnormalities per abnormal frog ­were recorded (Johnson et al., 1999). Other reports of malformations are found in Hebard and Brunson (1963), Miller (1968), Reynolds and Stephens (1984), and Lannoo (2008). Cercariae of R. ondatrae ­were reduced by 60% in experimental ­trials by predators such as mosquitofish, newts (Taricha), damselfly nymphs, and clam shrimp (Orlofske et al., 2012). The presence of certain predators (damselfly nymphs) and alternative hosts (newts) also reduced transmission rates to H. regilla, although mosquitofish did not. Other predators had no effect, such as dragonfly nymphs, backswimmers, and clams. ­These results suggest that certain predators may act as alternative hosts to the cercariae, whereas other predators do not. In effect, parasite-­host interactions are complex, and ­these interactions result in varied infection intensities of anuran larvae. The pathogenic ­water mold Saprolegnia had no effect on the survival of H. regilla larvae in experimental ­trials (Romansic et al., 2006). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. Mercury has been found in H. regilla from California at concentrations of 0.063–0.22 μg/g wet weight (mean 0.12) (Hothem et al., 2010). pH. Acidification does not appear to have affected the distribution of H. regilla in the Sierra Nevada Mountains of California (Bradford et al., 1994a). Nitrates, nitrites, and ammonia. Prolonged exposure to ammonium and nitrate compounds from agricultural runoff could have detrimental effects on H. regilla prior to metamorphosis. Mortality of H. regilla larvae is complete at

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concentrations of 15,000 mg/L N ­after a 10 day exposure to urea. ­Little mortality occurs at concentrations of 6,000 mg/L N or less (Schuytema and Nebeker, 1999a; Romansic et al., 2006). The LC50 (10 day) was estimated at 7,105–9,921 mg/L N for Pacific Treefrog larvae. For embryos, the LC50 (10 day) ranged from 25 to 32.4 mg/L NH4-­N, whereas the LC50 (10 for sodium nitrate was 578 mg/L NO3-­N (Schuytema and day) Nebeker, 1999b). Ammonium sulfate, another component of fertilizers, also has detrimental effects on larval growth rates and body weight, but not so much on survival; larvae are more sensitive to ammonium sulfate during the early stages of larval development than at ­later stages (Nebeker and Schuytema, 2000). The median lethal concentration of nitrite for larvae is 1.23 mg N-­NO2/L at 15 days, 3.6 at 7 days and >5.5 at 4 days (Marco et al., 1999). ­These results suggest that ­there may be sublethal effects on Pacific Treefrog larvae from ammonium fertilizers if exposure occurs throughout the developmental period. Hyliola regilla larvae are not particularly sensitive to the effects of nitrates alone, however. Chemicals. Pesticides pose a serious threat to Pacific Treefrog populations. At high elevation sites in the Sierras, Smalling et al. (2013) recorded 9 pesticides and 3 pesticide degradates in samples collected in 2009–2010. The fungicides pyraclostrobin and tebuconazole and the herbicide simazine ­were frequently detected in frog tissues, with maximum concentrations of 64–363 μg/kg wet weight. The insecticide/metabolites p.p’-­DDT and p.p’-­DDE ­were the most frequently detected pesticides from all sites examined. ­These authors also reported diazinon, 3,4-­DCA, 3,5-­DCA, bifenthrin, carbofuran, iprodione, myclobutanil and propyzamide from H. regilla. With the exception of diazinon and DDT, ­these pesticides had never been recorded from frog tissue. Additional pesticides ­were found in ­water and sediments from frog-­inhabited sites. The biological effects on frogs of the fungicides, in par­tic­u­lar, are unknown. Cholinesterase activity in tadpoles is depressed in populations exposed to organophosphorus pesticides (e.g., malathion, chlorpyrifos, diazinon), as ­these pesticides bind with cholinesterase to inhibit neural functioning. Cholinesterase of Pacific Treefrogs is particularly depressed in the Sierras of California, east of areas with heavy pesticide use as well as in areas downwind of the agricultural San Joaquin Valley. In such areas, >50% of the population had organophosphorus pesticide residues, with concentrations as high as 190 ppb (Sparling et al., 2001). In experimental ­trials, diazinon alters tadpole be­hav­ior by reducing the amount of time they spend in refuges, but does not affect survivorship at 0.5 or 1.0 mg/L (Kerby et al., 2012). Counterintuitively, greater activity by tadpoles exposed to both fish and diazinon had no effect on survivorship, since fish activity and attack rates ­were also reduced in the presence of the pesticide.

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Chlorpyrifos (to 17.4 ppb in larvae) and chlorothalonil (to 47.7 ppb in larvae and 4.5 ppb in eggs) have been found in H. regilla from natu­ral populations in California (Datta et al., 1998). Larvae experimentally exposed to field concentrations of chlorpyrifos, methyl parathion, temephos, fenthion, and malathion experienced a decreased thermal tolerance compared with controls. Malathion was least toxic, whereas chlorpyrifos and methyl parathion ­were the most active toxicants (Johnson, 1980). Methyl parathion and malathion also resulted in depressed activity in comparison with controls. The insecticide endosulfan is highly toxic to Pacific Treefrogs even at low concentrations, with an LC50(4 day) of 21.4 ppb (Jones et al., 2009). Endosulfan ­causes slight decreases in larval survival, larval deformities (kinked tails, loss of pigmentation), and changes in the be­hav­ior of survivors at environmentally relevant doses (Westman et al., 2010); it does not affect egg development, however. At concentrations >50 ng/L, azinphosmethyl had decreased survivorship, but no changes in be­hav­ior ­were evident (Westman et al., 2010). Kleinhenz et al. (2012) could not demonstrate any additive effects of the pesticides chlorpyrifos, endosulfan (I and II), diazinon, or malathion on Bd-­ infected larvae that could not already be attributable to Bd infection alone. The insecticide carbaryl actually tends to inhibit the effects of the Bd fungus. The time to tail resorption decreases in H. regilla tadpoles when they are exposed to carbaryl, and it counteracts the effects of Bd to extend tail resorption time (Rumschlag et al., 2014). Malathion has no effect on tail resorption regardless of Bd exposure. Heavy concentrations of noncholinesterase-­inhibiting pesticides have been found in the agricultural areas of California. ­These include endosulfan (in as many as 86% of the frogs sampled at some populations), 4,4’-­dichlorodiph enyldichloroethylene, 4,4’-­DDT and 2,4’-­DDT (DDTs in as many as 40% of the frogs sampled), and α-­ and γ-­ hexachlorocyclohexane in their tissues (Sparling et al., 2001). Organochlorine pesticide residues (PCBs, DDE, DDT, toxaphene) have been recorded in eggs, tadpoles, and adults from natu­ral populations in California (Datta et al., 1998; Sparling et al., 2001; Angermann et al., 2002). Concentrations ranged as high as 258 ppb DDE and 229 ppb PCBs on the campus of the University of California at Davis (Datta et al., 1998). Toxicant levels are much higher at low elevations in the Sierras than at high elevations, but the presence of ­these toxicants at high elevations at long distances from their point of origin suggests significant transport on wind and air currents. In a series of transplant experiments, Cowman (2005) examined pesticide uptake at 3 high elevation national parks

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in California. She found that DDE was pre­sent in 97% of her samples from Yosemite, 84% from Sequoia, and 15% from Lassen Volcanic national parks. Total endosulfans ­were detected in 3% of the Sequoia frogs, 9% of the Lassen Volcanic frogs, and 24% of the Yosemite frogs. Organophosphorus pesticides ­were not detected. Contaminants ­were accumulated directly as the tadpoles fed. Cowman (2005) further demonstrated chromosomal breakage in juvenile Pacific Treefrogs, mostly at Yosemite and Sequoia, which are closer to areas of high agricultural contamination than is Lassen Volcanic. Sublethal effects of pesticide exposure included shorter SULs, longer developmental periods, and lower survivorship at metamorphosis, especially at Sequoia National Park, than at the control site. Aerially transported pesticides likely have had severe impacts on frog populations in the Sierra Nevada Mountains. Other contaminants include the organophosphorus pesticides Guthion (lethal to tadpoles at 8.7–9.7 mg/L) and Guthion 2S (lethal to tadpoles at 1.4–1.5 mg/L) (Schuytema et al., 1995). Guthion affects total length, hind limb length, and mass at 3.6 mg/L. Guthion 2S is significantly more toxic than Guthion, and it has greater adverse effects on larval growth at lower concentrations than Guthion (Nebeker et al., 1998). The herbicide Roundup® is toxic to larvae, with an LC50 (24 hrs) of 0.43 mg/L (King and Wagner, 2010). The LC50 decreases at 15 days to 1.30 mg/L. Salinity. Pacific Treefrogs are found in somewhat saline habitats, such as at Newport Back Bay and Alkaline Marsh in California. Frogs found in ­these habitats are more tolerant of salinity than frogs originating in freshwater habitats, suggesting a population-­based adaptation to salinity tolerance (Weick and Brattstrom, 2020). At Newport Back Bay, most frogs ­were tolerant of 240 mM/L (40% of sea ­water) of NaCl for 72 hrs, whereas most frogs from freshwater Trabuco Canyon experienced some mortality at 120 mM/L (20% of sea ­water) NaCl and complete mortality at 180 mM/L (30% sea ­water) NaCl. Frogs from saline habitats osmoregulate by mainstreaming plasma sodium and increasing plasma urea. Urea is particularly impor­tant in maintaining internal osmoconcentration. UV light. Eggs and larvae exposed to ambient UVB radiation, as well as enhanced UVB radiation 15% and 30% above the ambient level, showed no differences from controls in terms of hatching success. Enhanced levels of UVB radiation reduced larval survivorship by 18.4%, but controls and ambient levels of UVB radiation had no effect on larval survivorship (Ovaska et al., 1997). Other studies have found similar results regarding development and hatching success (Blaustein et al., 1994b; Anzalone et al., 1998; Vredenburg et al., 2010a), and together suggest that H. regilla is relatively tolerant of UVB radiation. However, UVB may have effects

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on larvae when in combination with other sublethal stressors, such as nitrate. For example, larval mass is reduced when ­these stressors are combined in field experiments at low elevations; at high elevations, the combination reduces larval survivorship (Hatch and Blaustein, 2003). Neither of the stressors had an effect on larvae by themselves. Larval H. regilla do not avoid areas of high UV light ­either in the field or ­under experimental conditions. Instead, they tend to choose warmer ­water regardless of UV light levels (Bancroft et al., 2008). Eggs of Pacific Treefrogs have a relatively high content of photolyase, an enzyme known to be impor­tant in repairing UVB radiation damage to DNA (Blaustein et al., 1994b). STATUS AND CONSERVATION

Like all species of North American anuran, Pacific Treefrogs have dis­appeared from wetlands lost to development, agriculture, drainage, or habitat modification. In urban areas, remnant treefrog populations are particularly vulnerable, especially in semidesert and desert regions, and the species does not do well in urban landscapes where habitats are fragmented (Delaney et al., 2021). For example, treefrogs have virtually dis­appeared from the San Francisco Bay region, except in remnant parks and preserves (Banta and Morafka, 1966), although they still are pre­sent in some areas around greater Portland, Oregon (Holzer, 2014). Roads also may take a heavy toll (e.g., Watson et al., 2003; Crosby, 2014), and the frog may (Crosby, 2014) or may not (Schuett-­Hames et al., 2019) use underpasses when available to cross roads. Fencing does seem to reduce occurrence on roads, although frogs may still bypass single or double fencing (Crosby, 2014). Still, the species is not vulnerable on a landscape scale, particularly away from ­human population centers. Many species and populations of anurans have dis­ appeared from the high mountains of California during the

Habitat of Hyliola regilla. Photo: Chris Brown

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past few de­cades. However, ­these declines do not appear to have affected the presence of H. regilla at historically identified sites, although ­there may have been a decrease in abundance at many of the sites (Drost and Fellers, 1996). The species is still widely distributed in the Sierra Nevada, where it was found at 95% of sites where the species historically occurred (Brown et al., 2014). Exotic trout (Oncorhynchus, Salvelinus, Salmo) and other nonnative fish species have a negative impact on the presence of H. regilla in the high mountains of California and Oregon and in the Willamette Valley of Oregon (Matthews et al., 2001; Bull and Marx, 2002; Knapp, 2005; Pearl et al., 2005b; Welsh et al., 2006; Preston et al., 2012). Pacific Treefrogs are 2.4–3 times more likely to be found in ­waters without trout as opposed to ­those with trout (Matthews et al., 2001; Welsh et al., 2006). Wetland and riparian modification have undoubtedly affected many populations. For example, Banta (1961) noted that populations of Pacific Treefrogs had dis­appeared along the Colorado River as a result of dam building and reservoir filling. In mesocosm experiments, nonnative mosquitofish decreased the availability of zooplankton (resulting in slower larval development and smaller sizes) and caused considerable damage to Pacific Treefrog larvae (Preston et al., 2012). Pacific Treefrogs may be among the first amphibians to recolonize habitats affected by major catastrophic disturbances. ­After the 1980 eruption of Mt. St. Helens, Pacific Treefrogs ­were breeding successfully within the blast area by 1985. By 2000, they ­were pre­sent at more than 100 ponds. They currently are the most common amphibian throughout the area (Karlstrom, 1986; Crisafulli et al., 2005). In a similar vein, they readily occupy open-­canopied wetlands in commercial silvicultural forest and in early succession stages of habitats previously clearcut or in a sapling stage (Raphael, 1988; Bosakowski, 1999). As forests mature into old growth, abundance decreases with canopy closure. Hyliola regilla also ­will readily colonize artificial ponds (Monello and Wright, 1999; Holzer, 2014). In the Okanagan Valley of British Columbia, constructing breeding ponds and restoring wetlands allow for per­sis­tence and connectivity among populations, where Pacific Treefrogs colonized 15 of 21 ponds and produced metamorphs in 7 from 2007 to 2014 (Ashpole et al., 2018). Concern has been expressed about how climate change ­will affect many frog species. In experimental ­trials, O’Regan et al. (2014) showed that H. regilla displayed a degree of phenotypic plasticity in response to warming and artificial drying (altering ­water depth) in mesocosms. Warming decreased the time to metamorphosis, but survivorship increased. Tadpoles from permanent pools and temporary pools had similar growth rates and time to metamorphosis,

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but larvae from temporary pools ­were smaller and had only marginally greater survival rates. ­These mixed results suggest that warming and drying effects are additive or antagonistic, not synergistic. Pool warming and drying enhanced by climate change may not always be harmful to this species, but the mixed results also show that antagonistic responses are difficult to predict. Laboratory ­trials have further documented increases in developmental rates and body mass with increasing temperatures, and the authors have noted that plasticity in traits affecting larval fitness may not serve to buffer the effects of a warming climate (Thurman and Garcia, 2017). The optimum temperature range when this species showed maximum weight gain and developmental rate was 22–25°C, above which per­for­mance plateaued. Pacific Treefrogs have occasionally turned up in prepackaged salads sold in grocery stores (Hughes et al., 2019b).

Efforts to exclude them from agricultural fields in California include the installation of aluminum exclusion barriers with a horizontal lip to prevent frogs from trespassing the fence (Hughes et al., 2021). This species is not considered to be of conservation concern in Canada (Weller and Green, 1997) or in any US state.

Pseudacris brachyphona (Cope, 1889) Mountain Chorus Frog

Ground coloration ranges through several shades of tan to brown. Adults have a generally broad head, more rounded snout, and longer hind legs than sympatric Pseudacris. Toes have distinct disks with minimal webbing. ­There is a con­spic­u­ous triangle between the eyes when viewed dorsally (apex pointing posteriorly), and ­there is a broad stripe extending posteriorly from the tympanum that arches inward ­toward the midline of the dorsum. ­These stripes may connect, giving a cruciform appearance similar to that of P. crucifer. Some individuals have a spot above the vent. In addition, a few individuals have an irregular spotting pattern without the 2 dorsolateral stripes. Dorsal patterns are somewhat more variable in P. collinsorum (by having a greater frequency of broken stripes or lacking a dorsal pattern altogether) than in P. brachyphona. The venter is cream colored and usually unmarked, although some individuals have a few dark speckles. The undersides of the limbs have a yellowish coloration, which is most pronounced in breeding individuals. The male’s vocal sac is dusky to nearly jet black. Newly transformed froglets have the adult pattern. ­These species might sometimes be confused with P. ­crucifer, especially in individuals where the dorsolateral stripes fuse to form a cruciform pattern. However, the skin of P. brachyphona is rougher than that of P. crucifer, the toe disks are smaller, the species frequently has a yellowish coloration on the hind limbs, and the call is distinctive between ­these species (Netting, 1933). ­These characters presumably separate P. collinsorum from P. crucifer as well. ­There is a degree of sexual dimorphism, with males being generally olive brown and females being lighter and more

Pseudacris collinsorum Ospina, Tieu, Apodaca, and Lemmon, 2020 Collinses’ Mountain Chorus Frog ETYMOLOGY

brachyphona: from the Greek word brachys, meaning ‘short,’ and phōnē, meaning ‘voice.’ The name refers to the short trilling call of this species. collinsorum honors Joseph and Suzanne Collins for their lifelong efforts in promoting the science of herpetology and the conservation of amphibians and reptiles, particularly in Kansas. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pseudacris) brachyphona Synonyms: Chorophilus feriarum brachyphonus Pseudacris collinsorum was described in 2020 based on recognition of the southern populations of P. brachyphona as being distinct from northern populations. As such, lit­er­a­ture references to populations south of the Tennessee River system refer to P. collinsorum. IDENTIFICATION

Adults. ­These are essentially morphologically identical small frogs, with adults averaging approximately 30 mm SUL.

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COMMENT

The call of the Pacific Treefrog is one of the most recognized frog calls in the world. This species’ call is invariably used as background in Hollywood movies and tele­vi­sion programs, regardless of where the movie or program is supposed to take place. Listen for its repetitive “kreek-ek” call, especially in older movies whenever a frog call is used to enhance the “naturalness” of a scene.

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reddish brown. Females are also larger than males. In a large sample from Kentucky, the adult mean was 29.8 mm SUL, with a range of 25–32.2 mm (Barbour, 1957); on Big Black Mountain, males averaged 30 mm SUL (range 27–33 mm) and females 34.8 mm SUL (range 31–37 mm) (Barbour, 1953). Male P. brachyphona averaged 24.6 mm SUL in Pennsylvania; females in this latter population averaged 30.3 mm SUL (Hulse et al., 2001). In Ohio, males ­were 24.5–30 mm SUL and females 26–34 mm SUL (Walker, 1946). Males of P. collinsorum ranged from 24 to 32 mm SUL (mean 27 mm) in North Carolina (Schwartz, 1955). Larvae. The larvae of ­these species are small and deep bodied, and reach a length of about 25–30 mm SUL prior to metamorphosis (19 mm when rear legs become evident). The eyes are located dorsolaterally. The spiracle is not obvious, and it is located on the left side of the body. The body is generally black to dark brassy brown dorsally, and the venter is dark brown with numerous iridescent bronze specks. The tail fin is low and has scattered pigment dots, and ­there is a small amount of pigment on the lower edge of the tail musculature. ­There are 2 rows of labial teeth above the mouth (termed anterior) and 3 rows below the mouth (termed posterior). Additional descriptions of the tadpole are in Green (1938) and Altig (1970). Eggs. The vegetal pole of the eggs of P. brachyphona is creamy white to buff, whereas the dorsal animal pole is brown. Eggs have a single gelatinous envelope and range from 6 to 8.5 mm (mean 7 mm) in total dia­meter. The vitellus is 1.6 mm. The envelopes of adjacent eggs adhere to one another providing a degree of cohesion to the loosely formed mass. Descriptions of eggs are in Green (1938). Presumably, the eggs of P. collinsorum are similar. DISTRIBUTION

The Mountain Chorus Frog is found from southwestern Pennsylvania (as far north as Jefferson County), through the Allegheny Mountains and Cumberland Plateau of West ­Virginia, southeast Ohio, eastern Kentucky (and extending west along the Green River and Rolling Fork drainages to Edmonson and Jefferson counties), east central Tennessee (Cumberland Mountains, Blue Ridge Mountains), and southwest ­Virginia (Iron Mountain). Populations are found in Ohio (Carroll and Jefferson counties) and the Panhandle of West ­Virginia (Berkeley County). Only a single population remained in Mary­land as of 1999 of nine historically known populations (Forester et al., 2003), and the Mary­land herpetological atlas proj­ect could not confirm the continued presence of this species in the state (Cunningham and Nazdrowicz, 2018). The species occurs from 365 m to 1,220 m in elevation (Green, 1938; Barbour, 1953; Hoffman, 1981).

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Distribution of Pseudacris brachyphona and Pseudacris collinsorum

The range of P. collinsorum includes most of northern and central Alabama (south of the Tennessee River and north of the Fall Line, including the Talladega Uplands), extreme northeast Mississippi, and into Georgia and extreme southwestern North Carolina (Cherokee County; Floyd and Kilpatrick, 2002). Populations are found as far south as southeastern Alabama. Suggestions by Neill (1954) that the species might occur in the Florida Panhandle have not been verified. Impor­tant distributional references for t­ hese species include: range-­wide (Lemmon et al., 2007b), Alabama (Mount, 1975; Redmond and Mount, 1975; Graham, 2010; Alabama Herp Atlas, courtesy of David Laurencio), Georgia (Martof and Humphries, 1955; Williamson and Moulis, 1994; Jensen et al., 2008; Graham, 2010), Kentucky (Barbour, 1957, 1971), Mary­land (Harris, 1975; Cunningham and Nazdrowicz, 2018), Mid-­Atlantic (Beane et al., 2010), Mississippi (Ferguson, 1961), North Carolina (Schwartz, 1955; Dorcas et al., 2007), Ohio (Walker, 1946; Pfingsten, 1998; Davis and Menze, 2000), Pennsylvania (Hulse et al., 2001), Tennessee (Gentry, 1955; Barbour, 1956; Redmond and Scott, 1996; Wilmhoff et al., 1999; Niemiller and Reynolds, 2011), ­Virginia (Hoffman, 1955, 1981; Tobey, 1985; Mitchell and Reay, 1999), and West ­Virginia (Green and Pauley, 1987). FOSSIL REC­O RD

No fossils have been referred to ­these species.

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PSEUDACRIS BRACHYPHONA AND PSEUDACRIS COLLINSORUM 313

SYSTEMATICS AND GEOGRAPHIC VARIATION

Pseudacris brachyphona was described by Cope (1889) as Chorophilus feriarum brachyphonus, but its specific status was not recognized ­until Walker’s (1932) examination of additional individuals from Ohio. Although no subspecies have been described, ­there is a distinct difference in mitochondrial 12S and 16S rRNA genes between northern and southern populations (Lemmon et al., 2007b). The northern clade includes populations from Tennessee, Kentucky, and West ­Virginia northward, whereas the southern clade includes populations in North Carolina, Georgia, Alabama, and presumably Mississippi. The southern populations have now been described as a separate species, P. collinsorum (Ospina et al., 2020). Based on a cladistic analy­sis of morphology and karyology, Cocroft (1994) suggested that P. brachyphona was more closely related to P. feriarum and P. kalmi than to other Pseudacris, but this arrangement is not supported by molecular data. Instead, P. brachyphona and P. collinsorum are most closely related to P. brimleyi (Moriarty and Cannatella, 2004; Barrow et al., 2014; Banker et al., 2020). ­These latter 3 species diverged from a common ancestor during the Pliocene (4.6 mya), perhaps as a consequence of interspecific competition with a Nigrita clade ancestor (Lemmon et al., 2007a). Together, they are the ­sister clade of the Nigrita clade (nigrita, feriarum, triseriata, kalmi, fouquettei, clarkii, maculata), which together form the Trilling Chorus Frog clade (Barrow et al., 2014). This large clade shares a similar albumin phylogeny (Maxson and Wilson, 1975), and all members possess a cuboidal intercalary cartilage (Paukstis and Brown, 1987). Natu­ral hybridization occurs with P. feriarum in northeast Mississippi (sensu P. collinsorum) and south central Kentucky (sensu P. brachyphona), and with P. triseriata from central Kentucky (Lemmon et al., 2007b). Tadpoles can be produced in laboratory crosses between P. brachyphona and P. triseriata, and between ♀ P. crucifer and ♂ P. brachyphona. Crosses ­were not ­viable between ♂ P. crucifer and ♀ P. brachyphona (Green, 1952). In contrast, Mecham (1965) reported almost the exact opposite results to Green (1952) involving P. brachyphona and P. crucifer. Laboratory experiments also produced ­viable tadpoles in crosses with P. nigrita, P. feriarum, P. ornata, and P. brimleyi (Mecham, 1965). In most cases, metamorphosis was completed successfully and frogs grew normally, appearing intermediate in phenotype between the parental species. Lemmon et al. (2007b) found molecular evidence of crosses in nature between P. brachyphona and P. triseriata and P. feriarum. ­There may be regional or population variation in the extent of the X-­pattern on the dorsum. About 50% of

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P. collinsorum had fused stripes into ­either an X-­or a U-­pattern in northern Georgia (Martof and Humphries, 1955). In Ohio, however, Walker (1932) reported that only about 20% of P. brachyphona had fused dorsolateral stripes. In West ­Virginia, 55% of 1,251 P. brachyphona examined had a crescent pattern, 35% had a cruciform (fused) pattern, and 10% had no dorsal markings (Green, 1969). ­There is no correlation between dorsal markings and sex or size. ADULT HABITAT

Mountain Chorus Frogs and Collinses’ Mountain Chorus Frogs are associated almost entirely with deciduous woodlands, and individuals are not usually found away from woodland habitats. Pseudacris brachyphona tends to inhabit the Eastern cool temperate ruderal/deciduous, South Central mesophytic, and Allegheny/Cumberland dry oak forests, whereas P. collinsorum inhabits mixed evergreen and deciduous forests, with pines being a common ele­ment in ­these forests (Ospina et al., 2020). The amount of tree cover appears impor­tant to ­these species, regardless of ­whether they are in upland or valley habitats. When areas are cleared, ­these species are replaced by P. feriarum. Hulse et al. (2001) noted that Mountain Chorus Frogs are sometimes seen moving ­after summer rains, and that they are found ­under surface debris in the leaf litter. TERRESTRIAL ECOL­O GY

Walker (1932) rec­ords P. brachyphona as occurring in “deep woods” during the summer. Upon emergence, they may spend several weeks in transit to breeding sites, often pausing at small wetlands along the way (Green, 1952). ­After breeding, they return to terrestrial sites farther away from the early season breeding sites but may pause along the way and breed ­later in the summer at ­these intermediate wetlands. Green (1952) recorded marked individuals traveling 610 m between terrestrial overwintering sites and breeding ponds, and 1,220 m between breeding ponds in successive years. This is a terrestrial species, with no evidence of climbing. Small adult male P. brachyphona grow more rapidly than larger frogs, and growth essentially ceases in the largest individuals. A 23 mm frog might be expected to increase from 3 to 9 mm, whereas a 30 mm male might grow 0–4 mm in a year (Green, 1964). The oldest frogs even appear to shrink by 1–2 mm as they reach the largest size classes. Like most frogs, Mountain Chorus Frogs (and likely P. collinsorum as well) have a blue-­mode phototactic response, indicating they have true color vision (Hailman and Jaeger, 1974). They are monotonically photopositive, meaning that given a choice, individuals ­will seek out optimal illumination (Jaeger and Hailman, 1973). Presumably, ­these characteristics assist in terrestrial movements and in locating prey.

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314  Hylidae

CALLING ACTIVITY AND MATE SE­L ECTION

Mountain Chorus Frogs emerge from dormancy in late winter (December–­March), depending on location and temperature. Males arrive at breeding sites 2–8 days before females, and immediately begin calling (Green, 1938; Barbour and Walters, 1941). The call of P. brachyphona has been compared with that of P. triseriata, but at a more rapid rate and at a higher pitch. Barbour (1971) describes it as a rasping “reke-­rake.” The dominant frequency is 1,050–3,000 cps (mean 2,290 cps) (Thompson and Martof, 1957; Forester et al., 2003). Call length and the number of calls per minute are negatively correlated with temperature in P. brachyphona, but the number of pulses per second is positively correlated with temperature (Forester et al., 2003). The midpoint dominant frequency of the call is negatively correlated with the male’s body mass. The mean duration of the call of P. collinsorum is approximately 220–460 ms (range 400–508), which is much shorter than Pseudacris of the Nigrita clade (mean >600 milliseconds) (Thompson and Martof, 1957; Forester et al., 2003). Notes are repeated 50–70 times a minute, and they may be sustained over several minutes. From 24 to 26 notes (or pulses) comprise each call. Breaks last 15–20 sec between calls. ­There are distinct differences in the calls of ­these species. The pulse rate of P. brachyphona is 84.2 pulses/s, and for P. collinsorum it is 88.5 pulses/s. The mean dominant frequency for P. brachyphona is 2,456.3 Hz, whereas for P. collinsorum it is 2,716.2 Hz (Ospina et al., 2020). As such, the call of P. collinsorum is faster and higher pitched than that of its northern ­sister species. ­There is no significant difference in pulse number (ca. 25) between ­these species. Most calling occurs from March to July (Wright and Wright, 1949). However, P. collinsorum have been heard calling in early December in Alabama (Mount, 1975) and late February in the Hiawassee River floodplain of southwestern North Carolina (Schwartz, 1955), and P. brachyphona in January in West ­Virginia (Green and Pauley, 1987). At this latter time, the air temperature was 15ºC and the ­water temperature was 10ºC. Calling begins when ­water temperatures reach 7.5ºC and corresponding air temperatures are at least 5ºC (Barbour and Walters, 1941). In Mary­land, calling by P. brachyphona took place over a 26 day period in 1996 at temperatures of 5.4–18ºC (Forester et al., 2003). Mean chorus attendance by males was 13 days. Calling occurs both diurnally and at night. Males often call from hidden locations ­under dead leaves and debris along the shoreline of ditches and shallow temporary wetlands, from short grass clumps, or while floating on algal mats or other vegetation. Individuals may call from ­water, where only the head may be vis­i­ble, or from land. Males often float in the ­water with their back legs outstretched

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Eggs of Pseudacris brachyphona. Photo: J. Michael Butler

while calling, as is common in other species of Pseudacris, and Barbour and Walters (1941) reported that males ­were distributed throughout a wetland rather than being confined to areas near the shore. Barbour (1953) recorded a solitary male calling from the forest floor, 183 m from the nearest potential breeding site. While calling, males tend to face away from the pond if calling in ­water, or ­toward the pool if they are calling from along the shore. The diffuse nature of the call makes them difficult to locate when they are concealed by vegetation. According to Green (1938), however, males make no attempt to conceal their calling sites, and they space themselves in such a manner that ­there may be 3 or 4 males ­every 60 cm. The male makes no attempt at amplexus ­until actually contacted by the female. Females approach and back into calling males between the male’s forelegs (Green, 1952). The male then clasps the female, and the female swims to deeper ­water for oviposition. Amplexus is axillary, and the female usually rests with legs outstretched throughout amplexus. Oviposition is initiated as the female flexes her back. Amplexed males strug­gle vigorously to dislodge an inappropriate suitor. BREEDING SITES

Both quiet and slow-­moving temporary ­water may be used for breeding; permanent ­water is avoided. Breeding occurs in shallow, flooded pools, grassy pastures, roadside ditches, and in the vicinity of small springs. Mountain Chorus Frogs also have been observed calling from slow flowing ­waters adjacent to a culvert (Schwartz, 1955), from road ruts (Barbour, 1971; Adam and Lacki, 1993; Barry et al., 2008), from swampy areas, along small woodland streams, and from farm ponds (Barbour, 1957). Martof and Humphries (1955) recorded P. collinsorum breeding in open fields, although most accounts indicate that P. brachyphona prefers wooded areas. Males sometimes move between

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PSEUDACRIS BRACHYPHONA AND PSEUDACRIS COLLINSORUM 315

nearby breeding sites within a breeding season, but males also commonly breed in the same location from 1 year to the next (Green, 1952). Pseudacris brachyphona may sometimes occupy constructed ponds with shallow ­water (Drayer and Richter, 2016). REPRODUCTION

Breeding occurs from late winter to midsummer, depending on latitude and elevation, although Barbour (1953) recorded egg-­bearing females as late as 14 August. Breeding commences shortly ­after the initiation of calling. In North Carolina, P. collinsorum choruses ranged from approximately 30 to 35 calling males (Schwartz, 1955), and Green (1938) noted that 15–20 male P. brachyphona may call from a shallow pool only 1.5 m in dia­meter. Males remain at breeding sites during the entire breeding season, whereas females only visit sites on a single night to mate and oviposit their eggs. As a result, reports of sex ratios are highly male biased (6:1 in West ­Virginia, Green, 1938; 4.4 males per female in eastern Kentucky, Barbour, 1953). The extended breeding-­site residency also suggests that males mate more than once during a breeding season (Green, 1952). Eggs are deposited in small, soft, gelatinous masses of 3–50 eggs, with most masses containing about 14–34 eggs (Green, 1938; Barbour and Walters, 1941; Forester et al., 2003). The eggs are attached to vegetation, pool debris, or submerged grasses. Masses do not float, and eggs are often found on the pool substrate (Brown, 1956; Mitchell and Pauley, 2005). Females deposit from several hundred to more than 1,000 eggs during a breeding season. Forester et al. (2003) counted clutches of 90, 108, and 118 eggs. In West ­Virginia, Green (1938) recorded total egg counts of 318, 383, 406, and 1,479. In Kentucky, the total number of eggs was 983–1,202 (mean 1,092) (Barbour and Walters, 1941). Oviposition occurs over a period of several hours. Eggs hatch in 3–5 days at laboratory temperatures (18–22ºC), but at 7–10 days in the field (2–13ºC) (Barbour and Walters, 1941). The newly emerged larvae are 4.5–5 mm in total length. Although specific data are not available except for Brown (1956), reproduction is likely similar in P. collinsorum.

Tadpole of Pseudacris brachyphona. Photo: David Dennis

Adult Pseudacris brachyphona. Photo: J. Michael Butler

LARVAL ECOL­O GY

Larval development of P. brachyphona occurs over a period of about 45–60 days (Green, 1938; Barbour and Walters, 1941; Walker, 1946; Green, 1952). Newly metamorphosed frogs are 8 mm SUL, and frogs 11–13 mm ­were found in Ohio from mid-­June to mid-­August. POPULATION BIOLOGY

Sexual maturity of P. brachyphona occurs at 22 mm SUL for males and 28 mm SUL for females. Based on a sample of

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Adult Pseudacris collinsorum. Photo: Emily Lemmon

1,189 marked individuals followed over a 6 yr period, the population of Mountain Chorus Frogs studied by Green (1952) in West ­Virginia was composed of 71% 1 yr old frogs, 20% 2 yr old frogs, 6.8% 3 yr old frogs, and 1.8% 4 yr old

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316  Hylidae

frogs. Green (1952) recorded 2 Mountain Chorus Frogs 5 years ­after they ­were initially marked, but suggested that the mean life-­span was only 1.4 yrs ­after metamorphosis. ­These results imply that maturity is reached the first breeding season ­after metamorphosis, but this has not been verified. DIET

Prey of ­these species consists of small, ground-­dwelling invertebrates. Green (1952) reported that beetles (45%), spiders (25%), and true bugs (Hemiptera) (13%) formed the bulk of the prey items found in 42 individuals. Other prey included ants, leaf hoppers, flies, centipedes, earthworms, and larval lepidoptera (butterflies and moths). PREDATION AND DEFENSE

According to E.C. Lemmon (in Jensen et al., 2008), predators of larvae and adults include dragonfly larvae, aquatic beetles, fishing spiders, fish, salamander larvae (Ambystoma talpoideum), Eastern Red-­spotted Newts (Notophthalmus viridescens), Eastern Garter Snakes (Thamnophis sirtalis), ­water snakes (Nerodia sp.), and birds. Predation on adults by American Bullfrogs (L. catesbeianus) was recorded by Barbour (1957). When disturbed while calling, males cease calling or dive to the bottom of the breeding pool and attempt to conceal themselves ­under debris and grass. In terrestrial situations, they are reported to jump vigorously, resembling juvenile Wood Frogs. COMMUNITY ECOL­O GY

As noted above, t­ here is a sharp difference in microhabitat preference between P. brachyphona/P. collinsorum and the often-­sympatric P. feriarum. The Mountain Chorus Frog prefers wooded areas, whereas the Upland Chorus Frog is found only in open habitats. Although ­these frogs are often in close proximity to one another, they only rarely breed in the same wetlands or ditches (Wilson, 1945; Walker, 1946; Martof and Humphries, 1955; Barbour, 1957). Even then, they may breed at dif­fer­ent times, with the Mountain Chorus Frog breeding ­later than the Upland Chorus Frog (Walker, 1932). The Spring Peeper, Wood Frog, and American Toad are often found in association with t­ hese species. DISEASES, PARASITES, AND MALFORMATIONS

Larval Mountain Chorus Frogs are more sensitive to ranavirus than FV3 but experience a greater ­percent mortality from FV3 than ranavirus infection (Hoverman et al., 2011).

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Breeding habitat of Pseudacris brachyphona, Powell County, Kentucky. Photo: J. Michael Butler

SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Green (1938) considered P. brachyphona “fairly common throughout the central and southern part of West ­Virginia,” and Barbour and Walters (1941) described it as “one of the most abundant spring frogs to be found in northeastern Kentucky.” However, ­there is some evidence that the species has declined since the 1930s, primarily due to the loss of habitat (e.g., Mitchell and Reay, 1999; Davis and Menze, 2000; Weir et al., 2014). McClure (1996) found that none of Green’s dissertation sites had populations of P. brachyphona in the early 1990s and that nearby populations ­were small, scattered, threatened by habitat loss, and skewed ­toward larger individuals. Still, Weir et al. (2009) found no significant trends in occupancy for West ­Virginia populations followed over a 7 yr period. Hulse et al. (2001) stated that “all of the reports in the state [of Pennsylvania] are historical. No specimens have been reported in the past 20 yrs or so.” Only a single population in Savage River State Forest remained in western Mary­land as of 1999 (Forester et al., 2003). Mitchell and Pauley (2005) stated that P. collinsorum (as P. brachyphona) no longer occurred in North Carolina where Schwartz (1955) found them, but this species was rediscovered in the state in 2001 in Cherokee County (Floyd and Kilpatrick, 2002). A range-­ wide assessment of the status of both species clearly needs to be conducted. Pseudacris brachyphona is considered Endangered in Mary­land.

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Pseudacris brimleyi 317

Pseudacris brimleyi Brandt and Walker, 1933 Brimley’s Chorus Frog ETYMOLOGY

brimleyi: a patronym honoring Clement S. Brimley (1863– 1946), a North Carolina zoologist who wrote extensively on southeastern amphibians and reptiles (Cooper, 1979). NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pseudacris) brimleyi Synonyms: Hyla brimleyi IDENTIFICATION

Adults. This is a long-­legged chorus frog with 3 dark stripes on the dorsum bordered by a dark dorsolateral line ­running from the snout through the eye to the groin. The ground color is yellowish to reddish shades of brown; some individuals are dark brown. The lateral 2 dorsal stripes are often well defined, but the median stripe may be lighter; in other frogs all the dorsal stripes are light. The skin is smooth. A light line is pre­sent on the upper jaw and extends to the shoulder. ­There is no dark triangle between the eyes. The tympanum is distinct and smaller than the eye. ­There are longitudinal rather than transverse markings on the legs, and ­there is a dark line along the outer edge of the tibia. ­There is no webbing on the front digits, and the rear digits are only weakly webbed. The toe disks are only slightly enlarged. The venter is distinctly yellow with small dark spots on the chest; the amount of spotting is variable. Males have a dark vocal sac. Females are larger than males. Wright and Wright (1949) give a male range of 24–28 mm SUL and a female range of 27–35 mm SUL. Larvae. The ground color of larval P. brimleyi is dark with a greenish tinge, often with scattered gold, yellow, or brassy flecks over the body and dorsal tail musculature. Bodies are deep with a broadly rounded head. The chin and throat are darkly pigmented, with flecks or blotches. The tail musculature is sharply bicolored or striped, with a light dorsal stripe that extends forward through the eye to near the snout. The dark tail muscle pigmentation and stripe pattern may be red orange. Tail fins are sparsely speckled. Venters are heavi­ly spotted. Tadpoles reach about 30 mm TL. Eggs. The eggs are dark brown to black dorsally and white ventrally. ­There is a single jelly envelope surrounding the egg which averages 7.5 mm in dia­meter (range 6.75–8.64 mm); the mean vitellus dia­meter is 1.45 mm (range 1.3–1.71 mm) (Gosner and Black, 1958). Eggs are deposited in loose clumps.

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Distribution of Pseudacris brimleyi DISTRIBUTION

Brimley’s Chorus Frog occurs from eastern ­Virginia south to eastern Georgia on the Atlantic Coastal Plain. The rec­ord for northern Georgia (Brandt and Walker, 1933) is in error. The species has been recorded from Roanoke Island, North Carolina (Gaul and Mitchell, 2007). Impor­tant distributional references include Georgia (Jensen et al., 2008), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007), and ­Virginia (Tobey, 1985; Mitchell and Reay, 1999). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

A number of researchers have suggested hypotheses concerning the evolutionary relationship between P. brimleyi and other trilling chorus frogs (Hedges, 1986; Cocroft, 1994; Da Silva, 1997), although the results often ­were not congruent. The species is clearly allied with other Pseudacris based on its albumins (Maxson and Wilson, 1975). Pseudacris brimleyi is a member of the Trilling Chorus Frog clade and is most closely related to P. brachyphona and P. collinsorum of the Appalachian Mountains (Moriarty and Cannatella, 2004; Barrow et al., 2014; Banker et al., 2020). ­These species diverged during the Miocene about 4.6 mya, thus the Appalachian orogeny had no impact on speciation since it occurred much ­earlier (Lemmon et al., 2007a). Instead, competition between an ancestral P. brachyphona–­brimleyi species and an

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318  Hylidae

expanding ancestral P. feriarum–­kalmi–­triseriata species may have bisected the range leading to P. brachyphona in the mountains and P. brimleyi on the coast. The P. brachyphona–­ P. brimleyi clade is s­ ister group to the Nigrita clade. Successful hybridization in the laboratory has been reported with P. feriarum, P. ornata, P. nigrita, and P. brachyphona (Mecham, 1965). ADULT HABITAT

Brimley’s Chorus Frog is a species of low swampy woodlands along the Atlantic Coastal Plain.

Tadpole of Pseudacris brimleyi. Photo: Steve Bennett

TERRESTRIAL ECOL­O GY

­ ittle is known of the terrestrial ecol­ogy of this species. L Presumably they disperse to fields and swampy woodlands surrounding breeding sites. They forage terrestrially, hiding ­under woody debris, leaf litter, and downed logs by day and emerge at night to feed. Brimley’s Chorus Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination (Hailman and Jaeger, 1974). Brimley’s Chorus Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males call from the base of grass clumps and adjacent shallow ­water. The call is described as a short, rasping trill of 15–22 pulses (“kr-­r-­r-­a-­k”) lasting about 0.25 sec (Jensen et al., 2008); this is uttered in a series with short intervals in between. Males compete for calling positions and ­will drive off intruders. Calling may precede egg deposition by 1 to 2 weeks (Mitchell, 1986). Amplexus is supra-­axillary. BREEDING SITES

Breeding occurs in heavi­ly wooded, shallow (15–20 cm) grassy temporary pools in wet forest and floodplains. Gosner and Black (1958) recorded breeding in a flooded field adjacent to a flooded pine woodland, at the edge of red maple swamps, and in scrub thickets and roadside ditches. Breeding sites may be located at some distance from the nearest woodlands. REPRODUCTION

Breeding occurs in late winter to spring at air temperatures >4.5°C. Calling, amplexus, and egg deposition have been observed from February to April in North Carolina (Brandt and Walker, 1933; Gaul and Mitchell, 2007) and ­Virginia (Werler and McCallion, 1951; Mitchell, 1986). Gosner and

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Black (1958) counted egg clutches of 264 and 290 in 2 North Carolina females. Mitchell and Pague (2014) reported a mean of 245 (range 170–324) in ­Virginia. The eggs are deposited in multiple clumps on vegetation and woody debris just below the ­water’s surface. Eggs hatch in 4.5–5.5 days at 18–20°C, although Jensen et al. (2008) gave a figure of 1–2 weeks. LARVAL ECOL­O GY

The larval period is about 35–60 days (Gosner and Black, 1958). Newly metamorphosed froglets are 9–11 mm SUL. DIET

Nothing appears to be published on the feeding habits of P. brimleyi. They likely feed on small invertebrates, especially insects. PREDATION AND DEFENSE

The ground coloration makes this species cryptic and hard to see around breeding ponds and in woodland terrestrial habitats. Adults are preyed upon by Eastern Garter Snakes (Thamnophis sirtalis), watersnakes (Nerodia sp.), and other vertebrates. Larvae are eaten by mole salamanders (Ambystoma sp.), newts (Notophthalmus viridescens), fishing spiders, dragonfly larvae, predaceous beetles, fish, and birds (Jensen et al., 2008). POPULATION BIOLOGY

Sexual maturity is prob­ably attained by the first spring ­after metamorphosis, but nothing ­else is known about the species’ demography and population characteristics. COMMUNITY ECOL­O GY

The Savannah River Site (SRS) is one of the best-­studied locations in the United States in terms of its amphibian community. For >50 yrs, no rec­ords of P. brimleyi ­were made at this site despite intensive long-­term sampling. In 2007, however, Luhring (2008) recorded Brimley’s Chorus Frogs from the southern part of SRS. ­Either the frogs ­were absent for de­cades and had recolonized the site, or they had just

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Pseudacris clarkii 319

Adult Pseudacris brimleyi. Photo: Todd Pierson

been overlooked ­because of their small, scattered populations. Luhring (2008) suggested that the rediscovery of P. brimleyi illustrated “hidden biodiversity” that can be overlooked during even intensive faunal surveys. DISEASES, PARASITES, AND MALFORMATIONS

Breeding habitat of Pseudacris brimleyi, Hampton County, South Carolina. Photo: C.K. Dodd, Jr.

and R. ranae; and the acanthocephalan Centrorhynchus sp. (Brandt, 1936). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Parasites include the protozoans Nyctotherus cordiformis, Octomitus intestinalis, Opalina chorophili, O. hylaxena, O. oblanceolata, O. obtrigonoidea, O. pickeringii, O. virguloidea, Trichomonas augusta, Trypanosoma rotatorium, and an unidentified flagellate (Brandt, 1936). Other parasites include the trematodes Brachycoelium hospitale and Diplodiscus temperatus; the nematodes Agamascaris odontocephala, Agamonema sp., Cosmocercoides dukae, Ozwaldocruzia pipiens, Physaloptera sp., Rhabdias sp.,

No information is available.

Pseudacris clarkii (Baird, 1854) Spotted Chorus Frog

Information on P. clarkii may be listed in older publications as simply P. nigrita or P. triseriata (see Lord and Davis, 1956, for discussion).

ETYMOLOGY

clarkii: a patronym honoring Lt. John Henry Clark (ca. 1830–1885), a zoologist with the US-­Mexican Boundary Survey. Clark made extensive collections, including many new species. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pseudacris) clarkii Synonyms: Chorophilus triseriatus clarkii, Helocaetes clarkii, Heloecetes clarkii, Holocoetes clarkii, Hyla clarkii, Pseudacris nigrita clarkii, Pseudacris triseriata clarkii

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STATUS AND CONSERVATION

Brimley’s Chorus Frog appears to have declined on the southeastern Coastal Plain, especially in areas of urban expansion. The species is treated as of Special Concern in Georgia, where ­there are few recent rec­ords. In North Carolina, it is considered as a “species in need of monitoring.” An assessment of the species’ status is overdue.

IDENTIFICATION

Adults. Pseudacris clarkii is an attractive frog with green dorsal markings surrounded by thin black or dark gray borders on a light gray or greenish background. The green markings may appear as rows of spots and even coalesce to give the appearance of stripes or bars; the borders of the stripes are quite irregular. Some frogs may have a dark or green triangle between the eyes. ­There is a distinct dark or greenish band beginning at the snout and continuing through the eye ­toward the groin. A light line is pre­sent on the upper jaw. The limbs have dark green bands or spots. Venters are white. Males have dark brown throats. Both sexes can have spots or stripes, despite Wright and Wright’s

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320  Hylidae

(1933) speculation concerning sexual dimorphism in this character. The mean adult size in Texas was 29.1 mm SUL (Lord and Davis, 1956), but Collins et al. (2010) give a normal adult size of 19–28 mm SUL in Kansas. Smith (1934) mea­sured 4 individuals from 29.7 to 31 mm SUL in Kansas. Wright and Wright (1949) reported males at 20–29 mm SUL and females at 25–31 mm SUL. Larvae. The ground color of a mature tadpole is grayish olive. The tail musculature is darkly pigmented above the lateral line, and the tail fins are splotched with pigment, the dorsal fin more so than the ventral fin. The snout is rounded in lateral view. Maximum size is 30–31 mm TL. Bragg (1943b), and Eaton and Imagawa (1948) illustrated larval mouthparts, embryos, and larvae, and described developmental stages. Eggs. Eggs are pale brown, brownish gray, or dark gray dorsally and light gray, white, or ivory yellow ventrally. The eggs have a mean dia­meter of 2.3 mm (range 2.1–2.7) (Grubb, 1972), although Bragg (1943b) gave the mean dia­meter as 1.04 mm (range 0.99–1.3 mm), and Eaton and Imagawa (1948) gave a mean of 1.28 mm (range 1–1.66 mm); it seems likely that ­these latter mea­sure­ments did not include 1 or more of the jelly envelopes, whereas Grubb’s observations did. Wright and Wright (1949) reported the vitellus as 0.65–0.9 mm in dia­meter, with 2 surrounding jelly envelopes. The outer envelope is loose and 2.2–2.4 mm in dia­meter, whereas the inner envelope is 1.4–1.8 mm in dia­meter (Bragg, 1943b). DISTRIBUTION

Spotted Chorus Frogs occur from south central Kansas south to the Texas Gulf Coast and slightly into adjacent México. They occur throughout much of the Texas Panhandle and central Texas south into Tamaulipas, México, with an isolated population in Quay County, New Mexico (Kissner and Griffis-­Kyle, 2012). An isolated population was reported from central Montana (Black, 1970) but was based on misidentified P. maculata (see Corn, 1980b; Maxwell et al., 2003). Impor­ tant distributional references include Kansas (Collins et al., 2010), Oklahoma (Sievert and Sievert, 2006), and Texas (Burt, 1936; Brown, 1950; Dixon, 2000, 2013; Tipton et al., 2012). FOSSIL REC­O RD

Fossils of P. clarkii are known from Pleistocene locations in Nebraska and Texas (Holman, 1969, 2003). Differences in the scapula, radio-­ulna, sacral condyles, and ilium separate this species from other Pseudacris.

Distribution of Pseudacris clarkii

North Amer­i­ca (Moriarty and Cannatella, 2004; Barrow et al., 2014; Banker et al., 2020) rather than P. nigrita as indicated by Hedges (1986), Cocroft (1994), and Da Silva (1997). This group of frogs has the lowest ge­ne­tic variation among members of the genus Pseudacris (Lemmon et al., 2007a). Lemmon et al. (2007a) attributed this to aridification of the ­Great Plains. The species is clearly allied with other Pseudacris based on its albumins (Maxson and Wilson, 1975). Burt (1936) suggested that intergradation occurred with P. fouquettei (as P. triseriata) in southern Kansas and eastern Texas, but this is disputed by Smith (1934) and Bragg (1943a). Laboratory hybridization between P. streckeri and P. clarkii produces tadpoles that successfully complete metamorphosis (Mecham, 1957). Crosses between P. clarkii and the Dryophytes versicolor complex, P. fouquettei (possibly listed as P. triseriata by Moore, 1955), P. ornata or P. feriarum (listed as P. nigrita by Mecham, 1957) may produce larvae and metamorphs of uncertain viability. Crosses with Acris blanchardi, Dryophytes cinereus, D. arenicolor, or Hyliola regilla were not successful (Moore, 1955; Littlejohn, 1961a; Pierce, 1975). ADULT HABITAT

SYSTEMATICS AND GEOGRAPHIC VARIATION

Pseudacris clarkii is a member of the Trilling Chorus Frog clade and is most closely related to P. maculata of central

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The Spotted Chorus Frog is a species of the short-­grass and mixed prairies and prairie islands in wooded savanna. Along the eastern border of its range, P. clarkii may be found along

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Pseudacris clarkii 321

woodland borders and islands of prairie in the oak-­hickory savanna, but this is not a woodland or floodplain species (but see Bragg, 1941c). TERRESTRIAL ECOL­O GY

Most activity occurs during the spring and summer ­after rains. In Kansas, activity occurs from March to September (Collins et al., 2010). Spotted Chorus Frogs have been found ­under rocks and debris near breeding ponds and may use tunnels and burrows of other animals during the nonbreeding season. They migrate from terrestrial sites to breeding ponds, perhaps using olfactory cues. Grubb (1973) noted that reproductively active male P. clarkii ­were capable of detecting odors in ­water from ponds in which they had previously bred as opposed to ­water from “foreign” ponds or distilled ­water. This ability persisted in ­trials weeks ­after breeding and suggests the possibility that breeding adults might return to the ponds from which they metamorphosed, or at least return to ponds at which they had previously bred. Pseudacris clarkii is occasionally found around the entrances of caves (Black, 1973a). Most nonbreeding activity occurs at night, when frogs forage in pastures and fields (Bragg, 1943b). Activity tends to cease in the hot, dry, late summer. Spotted Chorus Frogs are photopositive in their phototactic response, suggesting they can use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination (Hailman and Jaeger, 1974). Spotted Chorus Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Pseudacris clarkii is an opportunistic breeder that can form breeding choruses at just about any time except during midsummer (Bragg, 1943b). Most calling occurs in the winter and spring. For example, Blair (1961b) recorded calling in Texas from February to May but noted a few instances of calling from late August to early October depending upon the year. Wiest (1982) heard calls from January to June in Texas. Lindsay (1958) also recorded amplexus in October. Bragg (1950a) reported calling in Oklahoma from March to September. Farther to the north in Kansas, breeding occurs from March to August with a peak from April to June (Collins et al., 2010). Choruses form rapidly with peak calling occurring soon ­after heavy rainfall, and chorusing continues both day and night. Choruses are heard at air temperatures of 8–27°C (Blair,

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1961b). Wiest (1982) noted calling at 3–23.2°C over a 60 day period in Texas. Males call from the center of dense grass clumps, which makes them difficult to locate. Calls are made rapidly and in succession. The dominant frequency is 2,850 cps with a trill rate of 75/sec and a duration of 0.21 sec (Michaud, 1962). In choice tests, female P. clarkii ­favor the calls of conspecific males over t­ hose of P. fouquettei (Michaud, 1962). Amplexus is axillary. BREEDING SITES

Breeding occurs in open shallow clear temporary ponds and pools of the arid ­Great Plains, such as ­those that form in meadows, on playas, or in pastures and roadside ditches (Bragg, 1943b). Such pools have grassy vegetation onto which frogs hold while calling. The species also uses buffalo wallows and pools in agricultural regions, as long as tadpoles have enough time to complete metamorphosis. Other wetland types include mesquite ponds, shallow ­water lily ponds, and occasionally ponds in floodplains (Bragg, 1941c). Ponds should have relatively tall (36 cm) and dense (213 stems/m2) vegetative cover. Pseudacris clarkii prefers breeding sites with extensive vegetative cover (52%), low aluminum concentrations, a more neutral pH (7.2 vs. 7.7), and deeper ­water (mean 22.5 cm), at least in the playas of west Texas (Anderson et al., 1999); adjacent land use, oxygen, conductivity, temperature, nitrate, and phosphate did not affect occupancy. Pseudacris clarkii does not occur in ponds with fish, as its eggs are readily palatable (Grubb, 1972), but it is occasionally found at permanent ponds. REPRODUCTION

Egg masses (6–37 eggs/mass) are deposited on upright vegetation, such as dead weed stems, sedges, and grasses just below the ­water’s surface. Bragg (1943b) provided a total clutch size of 916 eggs from a single female, whereas Wright and Wright (1949) recorded a clutch of 154 eggs (14–37 eggs/mass) from a single female. Blair (1961b) suggested that females may produce more than 1 set of eggs per year ­because of the prolonged and opportunistic breeding season. Eggs hatch in 2–10 days. At hatching, the gray larvae are 3.8–4.7 mm TL (Bragg, 1943b).

Tadpole of Pseudacris clarkii. Photo: Laurie Vitt

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322  Hylidae

Adult Pseudacris clarkii. Photo: C.K. Dodd, Jr.

Breeding habitat of Pseudacris clarkii, Travis County, Texas. Photo: Gary Nafis

LARVAL ECOL­O GY

between them appear to involve call discrimination and female preference. In experimental ­trials, female P. clarkii are able to discriminate and choose the calls of male conspecifics over ­those of P. streckeri males (Littlejohn, 1961b). Lord and Davis (1956) noted that P. fouquettei (as Pseudacris nigrita triseriata) nearly always bred at the same pools as P. clarkii; they never observed interspecific amplexus, however.

Nothing is known about larval ecol­ogy. ­After metamorphosis, young may remain in the vicinity of the breeding pool for 3–4 days before gradually dispersing (Bragg, 1943b). Dispersal by recent metamorphs is diurnal. DIET

No published information is available, but Spotted Chorus Frogs likely opportunistically consume a variety of small invertebrates. PREDATION AND DEFENSE

Upon disturbance, males cease calling and dive underwater, where they hide in the substrate and among submerged vegetation. The call has a ventriloquist effect that makes it difficult to locate a calling male. The eggs of this species are readily palatable to mosquitofish (Gambusia) (Grubb, 1972). Postmetamorphs likely are eaten by a wide variety of vertebrate predators. The Garter Snake Thamnophis marcianus has been reported to eat this species (Ford, 2020).

DISEASES, PARASITES, AND MALFORMATIONS

The myxozoan parasite Myxidium serotinum is known from P. clarkii, as are the protozoans Opalina sp., Hexamita intestinalis, Tritrichomonas augusta, and Nyctotherus cordiformis, the cestode Cylindrotaenia americana, the nematodes Gyrinicola batrachiensis and Cosmocercoides variabilis, and the mite Hannemania sp. (McAllister, 1991; Pierce et al., 2018). Bd has been reported from Oklahoma (Watters et al., 2016, 2019). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

POPULATION BIOLOGY

No information is available. COMMUNITY ECOL­O GY

Strecker’s and Spotted Chorus Frogs often are found at the same breeding sites, as their habitat preferences and breeding seasons overlap considerably. The main isolating mechanisms

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Spotted Chorus Frog populations are relatively tolerant of agriculture and rangeland as long as vegetative cover is maintained and hydroperiods of breeding ponds are sufficient for larval development (Anderson et al., 1999). Nothing is known concerning status and population trends, although populations undoubtedly have been lost, especially in rapidly urbanizing areas (e.g., Bragg, 1952).

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Pseudacris crucifer 323

Pseudacris crucifer (Wied-­Neuwied, 1838) Spring Peeper Rainette crucifère ETYMOLOGY

crucifer: from the Latin cruces, meaning ‘cross,’ and—­ifer, meaning ‘­bearer.’ The name refers to the X on the dorsum of this small frog. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Limnaoedus) crucifer Synonyms: Acris pickeringii, Hyla crucifer, Hyla crucifer bartramiana, Hyla crucifera, Hyla pickeringii, Hyliola pickeringii, Hylodes pickeringii, Parapseudacris crucifer IDENTIFICATION

Adults. Adult Spring Peepers are light tan to dark brown with a distinctive, dark-­colored X on the back. In a few individuals, the arms of the X may approach one another but not actually come into contact. The frogs may appear darker during the day than at night. A dark V-­shaped line connects the eyes. The belly is light and normally unmarked, although a few animals may have a slightly spotted venter. The legs are banded above but light under­neath, and the underside of the rear legs is light to lemon yellow on the femur. The toe tips are only faintly expanded, and ­there is no webbing between the toes. Males have a black vocal pouch that is evident throughout the year, although it is more pronounced in the breeding season. Males also have darkly pigmented testes. Juveniles are colored like adults, but the pattern may not be as evident, and the ground color is usually light tan. Erythristic peepers are reported from vari­ous localities in the Canadian Maritimes (McAlpine and Gilhen, 2018). Adult males are slightly smaller than adult females. In Florida, males are 23–30 mm SUL (mean 27.7 mm) and females 29–34 mm (mean 31.3 mm) (Owen, 1996); in New York, males are 18–30 mm (mean 24.4 mm) and females 23–33 mm (mean 28.3 mm) (Oplinger, 1963); and in Ohio, the male mean was 28.6 mm SUL and the female mean was 30.1 mm SUL (Gatz, 1981b). Other size data include: males >23.0 mm SUL and females 26.2–29 mm SUL in North Carolina (Alexander, 1966); males 21–34 mm SUL (mean 27.9 mm) in Maine (­Sullivan and Hinshaw, 1990); males 25–33 mm SUL (mean 29.9 mm) in Connecticut (Flores, 1978); males average 24–25 mm SUL and females 28 mm SUL in Rhode Island (Paton in Raithel, 2019); males 21–30 mm SUL (mean 24.6 mm) and females 23–35 mm

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SUL (mean 28 mm) in Connecticut (Klemens, 1993); males 25–32 mm SUL (mean 27.8 mm) in Nova Scotia (Bleakney, 1952); males 20–33 mm and females 27–37 mm in Nova Scotia (Gilhen, 1984); males 27–35 mm (mean 31.4 mm) and females 34–36 mm (mean 35 mm) on Prince Edward Island (Cook, 1967); and males 21–27 mm SUL (means 24.5–24.6 mm) and females 24–31 mm SUL (means 26.3–28.2 mm) in Pennsylvania (Meshaka et al., 2012c; Meshaka and Wingert, 2016). Body length and mass are highly correlated for both males and females (Owen, 1996). Larvae. Tadpoles of this species are small and deep bodied with a medium-­sized tail. The tail musculature is mottled, but the fins are clear or have blotches. Gosner and Black (1957b) noted 2 distinct tail morphologies: 1 narrow and thin and the other with a very wide tail fin. ­There are no dots on the grayish to light brown body. When viewed from above, the snout is square ­shaped. The larval mouthparts have 2 rows of marginal papillae, and the second anterior tooth row is longer than the first anterior tooth row. Larvae in the South are 8–10% larger than northern larvae (Gosner and Rossman, 1960). Mean length at hatching is 4.46 mm in Florida and 4.21 mm in New Jersey (range 4–4.25 mm; Gosner and Black, 1957b). At transformation, larvae are 33–39 mm in length (Harper, 1939a; Wright and Wright, 1949; Gosner and Black, 1957a; Gosner and Rossman, 1960). Larvae from northern and southern populations have the same coloration, however. Detailed descriptions of tadpoles are in Wright (1929) and Gosner and Black (1957b). Descriptions and illustrations of larval mouthparts are in Hinckley (1882) and Dodd (2004). Logier (1942) reported “intense inky black” tadpoles from the Sault St. Marie region of Ontario. Eggs. The eggs have a mean dia­meter of 1.5 mm (1.4–2 mm) in the North and 2.56 mm (2.45–2.8 mm) in the South (Gosner and Rossman, 1960), and a vitellus of 0.9–1.13 mm. They are deposited singly or in small bunches attached to submerged vegetation near the bottom. Females oviposit both large and small eggs in a single clutch. As many as 1,500 eggs may be oviposited by a single female during the breeding season (Wright, 1914; Livezey and Wright, 1947; Gilhen, 1984), usually in ­water 0.25–0.5 m in depth. The eggs are black or brown dorsally, white or cream ventrally, and have 2 gelatinous envelopes. Most authors report that eggs hatch in 5–14 days, depending on temperature, although MacCulloch (2002) states that hatching requires up to 3 weeks in Ontario. DISTRIBUTION

Spring Peepers are widely distributed in eastern North Amer­i­ca. They occur from the Churchill River Valley in Labrador and the Canadian Maritimes across southern

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324  Hylidae

Distribution of Pseudacris crucifer

Québec, around James Bay, and westward across Ontario to eastern and central Manitoba. ­There is a single report from far northwestern Ontario near Sachigo Lake (Weller and Green, 1997). Observations from Labrador ­were questioned (Maunder, 1983) but have been confirmed (Bergman, 1999; Rashleigh and Crowell, 2018). The western extent of the range includes northern and eastern Minnesota and central Iowa, most of Missouri (but not the northwest tip), to eastern Oklahoma and Texas. The species barely enters Kansas. Pseudacris crucifer occurs throughout the eastern and southeastern United States, and southward on the Florida peninsula to Orange and Sumter counties (Stevenson and Crowe, 1992; Owen, 1996). ­There may be occasional gaps in its distribution; for example, it appears to be absent from the Nashville Basin in Tennessee (Niemiller and Reynolds, 2011) and from western Illinois. Elevations range from sea level to 1,650 m in the ­Great Smoky Mountains (Mathews and Echternacht, 1984). Spring Peepers are found on islands, including Ile d’Orleans in Québec (Fortin et al., 2004a), Prince Edward Island (Cook, 1967), Pelee Island in Lake Erie (Langlois, 1964; Hecnar et al., 2002), the Apostle Islands in Lake Superior (Hecnar et al., 2002; Bowen and Beever, 2010), Isle Royale (Ruthven, 1912), the Georgian Bay islands in Lake Huron (Hecnar et al., 2002), Walpole Island in Lake St. Clair (Woodliffe, 1989), Martha’s Vineyard and Nantucket (Lazell, 1976), islands in Narragansett Bay (Raithel, 2019), Long Island, New York (Overton, 1914), Staten Island, New York,

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but declining (Nicholls et al., 2017), Three Mile Island in the Susquehanna River, Pennsylvania (Meshaka and Wingert, 2016), Kent Island, Mary­land (Grogan and Bystrak, 1973b), islands at the tip of the Delmarva Peninsula (Mitchell, 2012), and the Pleistocene barrier islands of Georgia (Shoop and Ruckdeschel, 2006). Reports of introductions to Cuba are apparently not accurate (Estrada and Ruibal, 1999). Impor­tant distributional references include: Alabama (Mount, 1975), Arkansas (Trauth et al., 2004), Canada (Bleakney, 1954, 1958a), Connecticut (Klemens, 1993; Klemens et al., 2021), Florida (Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), ­Great Lakes Region (Harding and Mifsud, 2017), Illinois (Smith, 1961; Mierzwa, 1998), Indiana (Brodman and Kilmurry, 1998; Minton, 2001; Brodman, 2003), Iowa (Vandewalle et al., 1996), Kansas (Rundquist, 1978; Collins, 1993; Collins et al., 2010), Labrador (Bergman, 1999; Rashleigh and Crowell, 2018), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Maine (Hunter et al., 1999), Mary­land (Cunningham and Nazdrowicz, 2018), Mas­sa­chu­setts (Lazell, 1976), Michigan (Ruthven, 1912), Minnesota (Oldfield and Moriarty, 1994; Moriarty and Hall, 2014), Missouri (Johnson, 2000; Daniel and Edmond, 2006), New Brunswick (Gorham, 1970), New Hampshire (Oliver and Bailey, 1939; Taylor, 1993), New York (Gibbs et al., 2007), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007), Nova Scotia (Gilhen, 1984), Ohio (Walker, 1946; Pfingsten, 1998; Davis and Menze, 2000), Ontario (Logier, 1928; Schueler, 1973; MacCulloch, 2002); Pennsylvania (Hulse et al., 2001), Prince Edward Island (Cook, 1967), Québec (McCoy and Durden, 1965; Bider and Matte, 1996; Desroches and Rodrigue, 2004), Rhode Island (Raithel, 2019), South Carolina (Dodd and Barichivich, 2017; Fields, 2019), Tennessee (Redmond and Scott, 1996; Butterfield et al., 2009; Niemiller and Reynolds, 2011), Texas (Smith and Sanders, 1952; Dixon, 2000, 2013; Tipton et al., 2012), Vermont (Andrews, 2001, 2019), ­Virginia (Tobey, 1985; Mitchell and Reay, 1999), West ­Virginia (Green and Pauley, 1987), and Wisconsin (Suzuki, 1951; Vogt, 1981; Mossman et al., 1998). FOSSIL REC­O RD

This species is known from Pleistocene (Irvingtonian and Rancholabrean) fossil deposits in Georgia, Mary­land, Pennsylvania, Tennessee, ­Virginia, and West ­Virginia (Holman, 2003). Most of the fossil sites represent cave deposits. SYSTEMATICS AND GEOGRAPHIC VARIATION

Two subspecies have been recognized historically: the Northern Spring Peeper (Pseudacris crucifer crucifer) and the Southern Spring Peeper (Pseudacris crucifer bartramiana).

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Pseudacris crucifer 325

Populations of the Spring Peeper in southern Georgia and Florida differ phenotypically from northern populations by the presence of ventral spotting, possession of dark spots instead of a stripe on the upper jaw, having a richer coloration, and by having broader dorsal stripes (Harper, 1939a). However, ­there is no molecular evidence to support the recognition of ­these subspecies as distinct evolutionary lineages (Austin et al., 2002; Moriarty and Cannatella, 2004; Cairns et al., 2021). Gilhen (1984) also noted that populations in the Canadian Maritimes have a more distorted X mark dorsally than populations farther south, and that the X may be fragmented or connected to other dark markings. Pseudacris prob­ably evolved from a Dryophytes-­like ancestor in the Tertiary—­that is, during the Miocene to early Pliocene (Hedges, 1986; Cairns et al., 2021). Moriarty and Cannatella (2004) suggested that ­there are 4 evolutionary lineages in Pseudacris. Pseudacris crucifer was more closely related to Pseudacris ocularis in the Crucifer clade than it is to the Trilling Chorus Frog clade (most eastern Pseudacris), the Fat Chorus Frog clade (e.g., P. streckeri), or the West Coast clade (e.g., Hyliola regilla) (Barrow et al., 2014; Banker et al., 2020). Interpretations of the number of ge­ne­tic lineages within this species have evolved as more data have become available. Austin et al. (2002, 2004a) suggested ­there ­were 3 major phyloge­ne­tic clades (a complex and diverse eastern, a southwestern, and a western) reflecting the complex effects Pleistocene climatic fluctuation had on distribution, particularly as frogs dispersed from refugia in the Ozark Central Highlands and from areas within the Southern Appalachians. ­These authors noted that diversification from other Pseudacris possibly originated in the Pliocene, but that phyloge­ne­tic diversity was amplified by isolation in refugia through the Pleistocene glacial cycles. More recent analyses comparing male advertisement calls and mitochondrial and nuclear DNA markers suggest extensive geographic and topological mitonuclear discordance involving 3 nuclear lineages (Western, Northern, Southern) containing 6 structured mtDNA lineages (East, Interior, West, Southeast, Southwest, Texas) (Cairns et al., 2021). Male advertisement calls are incongruent with the ge­ne­tic structure of this species (Cairns et al., 2021). ­These results suggest that allopatry was impor­tant in the origins of ­these phylogeographic patterns, but that subsequent range expansions, hybridization, and introgression maintained its status as a single species rather than resulting in radiation into multiple species. Some of the highest levels of haplotype diversity occur in northern populations, where rapidly expanding lineages have come into contact and introgression has occurred (Austin et al., 2002; Austin et al., 2004a). This is particularly evident in southwestern Ontario, where 2 major clades come into

Dodd_Canada_int_5pgs_B1&B2.indd 325

contact. Stewart et al. (2016) found evidence of pre-­mating isolating mechanisms and divergence in a range of traits between the 2 (Interior and Eastern) lineages in the area of contact. Specifically, ­there ­were differences in morphology (Eastern frogs are shorter and heavier than Interior frogs), male call attributes, and female preference for male calls from their natal lineages. Hybrids between the lineages ­were morphologically distinct, but hybrid males had calls intermediate to ­those of the 2 lineages. Female hybrids, however, preferred the calls of Eastern males. Behavioral differences also ­were evident even among calling males, with Eastern males almost always actively calling, while hybrid males assumed a satellite position (Stewart, 2013). Despite ­these suggestions of pre-­mating isolation between the 2 lineages and post-­zygotic complications (more abnormalities occur in hybrid tadpoles), isolation is incomplete. ­These results provide insights into the pro­cess of early-­stage speciation and divergence in a phylogeographic context. Spring Peepers usually have been assigned to ­either the genera Pseudacris or Hyla (now Dryophytes) over the last several de­cades. Hardy and Burroughs (1986) compared lit­er­a­ture data on osteology, external morphology, internal anatomy, biochemistry, and life history and concluded that Spring Peepers ­were intermediate between Pseudacris and Hyla. They proposed a new genus, Parapseudacris, to accommodate this species. This arrangement has not received subsequent support. Hedges (1986) first suggested that Spring Peepers should be placed in Pseudacris based on allozyme data, but this transfer was disputed by Cocroft (1994), who analyzed a series of morphological characters. ­After adding additional morphological characters to the Cocroft (1994) dataset, Da Silva (1997) concluded that Spring Peepers belonged in Pseudacris, an alignment that also coincides with chromosomal banding patterns (Wiley, 1982). The expanded morphological data and the more recent phyloge­ne­tic analyses of Moriarty and Cannatella (2004) confirm the placement of Spring Peepers within Pseudacris. Spring Peepers can be hybridized artificially in the laboratory with a number of other Pseudacris. Crosses of male P. feriarum, P. brachyphona, P. kalmi, P. triseriata, or P. streckeri with female P. crucifer produce a few tadpoles, but most are not ­viable, and developmental abnormalities are common (Blair, 1941b; Gosner, 1956; Mecham, 1957, 1965). Likewise, hybridization with P. nigrita does not result in larvae that metamorphose (Moore, 1955). Crosses between male P. crucifer and female P. ornata, P. feriarum, P. kalmi, or P. brachyphona also may produce a few tadpoles. Very few tadpoles are produced in crosses between male P. crucifer and female P. streckeri, and a high proportion of developmental abnormalities prevent successful metamorphosis

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326  Hylidae

(Mecham, 1957). Crosses between male P. crucifer and female P. triseriata result in about 10% of the eggs producing larvae, but survival to metamorphosis is extremely rare, again ­because of severe developmental abnormalities (Blair, 1941b). Mecham (1965) noted that sexual development in hybrids is abnormal, and that hybrids that survive do not appear to be fertile. Gosner (1956) found no evidence of hybridization between P. crucifer and P. kalmi in a large sample from a natu­ral population. ADULT HABITAT

The Spring Peeper is a frog of eastern forested habitats (Knutson et al., 2000; Price et al., 2004), and the extent of forests at vari­ous scales (to 3,000 m) is a good predictor of the presence of peepers at individual breeding sites. Likewise, the extent of open habitats is inversely correlated with Spring Peeper occupancy. They are found in a wide variety of terrestrial communities, including xeric hammock, headwater wetlands, upland hardwood forest, mixed-­hardwood and pine forest, mesophytic forest, urban wetlands, ­Great Lakes pine barrens, and forested ravines (Carr, 1940a; Marshall and Buell, 1955; DeGraaf and Rudis, 1990; Enge et al., 1996; Enge, 1998a, 1998b; Enge and Wood, 1998; Varhegyi et al., 1998; Evrard and Hoffman, 2000; Alix et al., 2014b). Peepers are pre­sent but tend to be less common in tamarack (Larix laricina) and coniferous forest than in hardwood or mixed-­hardwood forest. In other­wise developed land, they occur in riparian forest along streams (Burbrink et al., 1998; Barrett et al., 2016). Occupancy is often quite high in south central Michigan and is related to wetland proximity, but it is difficult to predict which habitat variables are most impor­tant to the species (Roloff et al., 2011). At the southern end of its range, P. crucifer prefers mesic hardwood hammocks with some degree of topographic relief; the lack of topographic relief south of the Orlando area, coupled with their cool-­weather breeding cycle, may limit their distribution on the peninsula. Likewise, their close association with forested habitats limits their distribution into the prairie and more arid regions to the west. Spring Peepers have occasionally been found at the entrances to and passages within caves (Franz, 1967; Black, 1973a; Niemiller et al., 2016; Zigler et al., 2020; Camp and Jensen, 2021). TERRESTRIAL ECOL­O GY

Away from breeding wetlands, Spring Peepers are not commonly observed. They are found in terrestrial leaf litter and among surface debris, where they forage for small invertebrates. McAlister (1963) noted an arboreal aggregation of Spring Peepers in September, when numerous frogs ­were feeding on small arthropods in bushes along a road.

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Carr (1940a) also noted arboreal retreat sites ­under bark and in knotholes, and they are occasionally observed in pitcher plants (Sarracenia purpurea) (Russell, 2008). They appear to establish small home ranges (dia­meter of 1.7– 5.4 m) around downed logs, woody debris, stumps, and surface vegetation (Delzell, 1958). Home ranges overlap with one another, but away from the breeding site individuals do not interact. Overwintering sites may be as far away as 300 m from the summer home range. Activity occurs throughout the warm season, even in Florida where I have occasionally observed them hopping among litter in a mesic hammock in August (also noted by Meshaka, 2009, in Pennsylvania). In ­Virginia, they are active year-­round (Gibson and Sattler, 2020). As noted below, Spring Peepers are heard calling during periodic warm weather in autumn and early winter, even in the far North (e.g., Raithel, 2019). This suggests that they routinely remain active late in the season, depending upon weather conditions. Most activity away from breeding sites is devoted to feeding and growth during the warm activity season. In nature, feeding occurs in a bimodal pattern—­that is, in early morning and late after­noon. Oplinger (1967) noted that peepers often did not feed ­after 30 min of concentrated feeding, even though prey was still available. Presumably, ­these individuals ­were temporarily satiated. Most individuals do not have food in their stomachs first ­thing in the morning, suggesting that feeding does not occur at night. By midmorning, most individuals have fed. The fact that Spring Peepers have color vision (Hailman and Jaeger, 1974) also suggests that most feeding activity occurs during daylight hours. Although often portrayed as a cold-­favoring species, P. crucifer can tolerate warm temperatures. The CTmax of P. crucifer is 34.8–39.4°C (John-­Alder et al., 1988 [New York]; Katzenberger et al., 2018 [Pennsylvania]), although this species seeks refuge during very hot days in summer. In Minnesota, Brattstrom (1963) recorded body temperatures 3.5–6°C warmer than ambient air and soil temperatures and suggested that the frogs ­were purposely absorbing solar radiation through their exposed position in the vegetation. Since peepers feed during the early morning hours, increasing temperature through basking could facilitate digestion. Spring Peepers appear to prefer higher temperatures when in the vicinity of conspecifics than when alone, although the significance of this preference is not known (Gatten and Hill, 1984). Thermal analyses suggest that microclimate temperatures ­will be more impor­tant than mean air temperatures in this species’ response to climate change. The presence of Spring Peepers is positively correlated with forested habitats within 100–1,000 m of breeding sites

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(Herrmann et al., 2005; Eigenbrod et al., 2008), depending on location and method used to identify habitat variables, suggesting that they spend much of their nonbreeding activity in terrestrial habitats in rather close proximity to wetlands. Owen (1996) also hypothesized that they spent most of their time in terrestrial habitats adjacent to the forested wetlands in Florida in which they bred. Density seems to increase with the percentage of forest cover, with greatest densities at >80% forest cover. On a localized scale (20 days ­after the first individual calls are heard. Chorusing occurs at a rather constant level for several months thereafter (Owen, 1996; Todd et al., 2003) ­until the end of the breeding season, when it tapers off with warmer weather. The optimum temperature for calling is 10–20°C, but calls can be heard from 30°C (Steelman and Dorcas, 2010). Hanna et al. (2014) noted that temperature was inversely related to call duration and positively correlated with call rate. Calling sequences are highly variable in terms of the number of calls per sequence; Gerhardt (1973) recorded a male whose sequence consisted of 66 consecutive calls. Call duration, dominant frequency, and the number of calls per minute are similar throughout the species’ geographic range, ­after adjusting for the effects of temperature. For example, calls in South Carolina lasted a mean of 0.18 sec, with a mean number of 49 calls/min (range 30–70). The midpoint dominant frequency at 15°C was 2,600 Hz (range 2,200– 2,900). ­These values are close to ­those reported for P. crucifer in Maine (mean 2,809 cps, 0.21 sec duration, call rate 53.9 calls/min; ­Sullivan and Hinshaw, 1990), Mary­land

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(2,625–3,549 cps, mean 3,061 cps; Forester and Czarnowsky, 1985), Missouri (2,800–3,360 cps, 0.09–0.25 sec duration; Doherty and Gerhardt, 1984), New York (2,588– 3,212 cps, mean 2,895 cps; Wilczynski et al., 1984), Texas (2,400–2,600 cps, mean 2,467 cps, 0.14 sec duration; Blair, 1958a), and North Carolina (3,200 cps, 0.13 sec duration; Martof, 1961). The dominant call frequency and call rate are positively correlated with temperature, but the duration of the call is negatively correlated with temperature (­Sullivan and Hinshaw, 1990). Dominant frequency is negatively correlated with male body size (Lykens and Forester, 1987). Call characteristics are specific to individuals; that is, a male with certain call characteristics 1 night ­will have the same characteristics on subsequent nights. Females have definite call preferences (calls of 0.15–0.30 sec duration and 2,875 cps), but the female’s auditory perceptions are only roughly attuned to the temporal and spectral properties of the male’s call (Doherty and Gerhardt, 1984). In another series of experiments, females had no real preferences for the frequency of the male’s call if no background noise was evident, but in situations mimicking natu­ral calling background noise, they chose callers with high frequencies (Schwartz and Gerhardt, 1998). This makes the male’s call audible to females in a variety of environmental conditions, particularly at a variety of temperatures and levels of background noise. However, males can barely detect their own calls (Wilczynski et al., 1984). In terms of hearing, the extent of the male’s call is roughly 1.1–3.8 m, whereas females have an auditory space of 2.2–11.2 m. Calling from trees or brush, even at only 50 cm, increases the male’s auditory space to 1.8–11.6 m and that of females from 4.6 to 69.4 m (Brenowitz et al., 1984). Thus, males space themselves generally 80% ­after only 3 sampling nights during Ontario road transect surveys (de Solla et al., 2005), 79–86% in Québec (Bonin et al., 1997a; Lepage et al., 1997), and 76% in Wisconsin (Mossman et al., 1998). During call surveys in Illinois over a 4 yr period (1986– 1989), populations appeared to be increasing (Florey and

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Breeding habitat of Pseudacris crucifer, Noxubee County, Mississippi. Photo: C.K. Dodd, Jr.

Mullin, 2005). Interestingly, Spring Peeper occupancy of breeding ponds does not seem to be affected by nearby road traffic density (Eigenbrod et al., 2008). That does not mean, however, that significant mortality does not occur on roads. Over a 3 day period in March, Carpenter and Delzell (1951) recorded 1,080 Spring Peepers from a road approximately 10 m from a breeding pond. They noted that 75% of the amphibians they observed (10 species) ­were found dead. Mortality of ­these small frogs is prob­ably quite high on roads, but their squashed delicate bodies prob­ably do not remain long and are thus vastly ­under counted in road-­effect studies. The area of immediate adverse impact (the “road-­ zone” effect) extends 200–300 m from the pavement (Eigenbrod et al., 2009). Spring Peepers respond to noise by producing shorter calls. When exposed to high-­frequency noise levels, they

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lower the frequency of their calls immediately ­after the noise ceases. However, the call rate does not change. Hanna et al. (2014) concluded that noise affects the structure of the advertisement call of this species, but that Spring Peepers have mechanisms for altering signal structure in response to anthropogenic noise. This response might be particularly impor­tant in breeding populations near roads. Spring Peepers prefer hardwood sites to cutover hardwoods, pines, and burned sites (McLeod, 1995; McLeod and Gates, 1998). They tend to avoid recently mined sites, and their numbers may be severely reduced in agricultural areas, especially if forested habitats are not in close proximity (Knutson et al., 2004; Anderson and Arruda, 2006). However, old mine sites may be recolonized through time as succession occurs; Myers and Klimstra (1963) recorded P. crucifer as common on sites where coal mining had ceased 19–29 yrs previously. In Ontario, Spring Peeper abundance was negatively associated with mining activities as a result of the alteration of vegetative structure (Sasaki et al., 2015). Peepers dis­appear from clearcut sites but may return if suitable breeding sites remain nearby (Clawson et al., 1997). However, it is difficult to evaluate Clawson et al.’s (1997) results and gauge the extent of the effects of clearcutting since numbers ­were not provided, and the study was conducted for only 6 months postharvest; presence is not equivalent to having no effects or minimal effects. In another study in New Brunswick, conversion of natu­ral forests to conifer plantations had detrimental impacts on P. crucifer. Breeding sites in converted areas tended to have short hydroperiods, which resulted in poor recruitment (Waldick et al., 1999). Spring Peepers are somewhat resistant to habitat fragmentation, occurring in a wide variety of forest fragmentation gradients (Gibbs, 1998b; Kolozsvary and Swihart, 1999). They may breed in wetlands within

power-­line rights-­of-­way, as long as ­there are forest patches immediately nearby (Fortin et al., 2004b). ­There is evidence that Spring Peepers are adversely affected by ­cattle using their breeding ponds. Larvae from such sites are smaller than comparable sites where ­cattle do not have access. Cattle-­access ponds have high turbidity, which may inhibit tadpole development and survival (Schmutzer et al., 2008). Repatriation efforts ­were carried out for P. crucifer at the Gateway National Recreation Area, New York, from 1980 to 1990. A total of 68 adults and 12,700 larvae ­were used in the repatriation. According to Cook (1989, 2008), this repatriation has been successful at 3 of 4 re­introduction sites. Spring Peepers normally colonize restored and newly created ponds (Briggler, 1998; Nyberg and Lerner, 2000; Pechmann et al., 2001; Stevens et al., 2002; Touré and Middendorf, 2002; Brodman et al., 2006; Palis, 2007; Shulse et al., 2010; Denton and Richter, 2013; Terrell et al., 2014a; Walls et al., 2014a; Mitchell, 2016; Stiles et al., 2017a; Baecher et al., 2018), but not in all circumstances, at least over the time monitored (Kline, 1998; Lehtinen and Galatowitsch, 2001). In some cases, they may actually be more abundant in constructed wetlands than in natu­ral wetlands (Drayer and Richter, 2016). Numbers at recent mine treatment and reclamation sites may be low, although ­these sites may harbor calling males (Lacki et al., 1992; Anderson and Arruda, 2006). In South Carolina, a series of 3 created ponds produced 1,500 juveniles over a 7 yr period (Pechmann et al., 2001). Playing conspecific chorus sounds does not increase the likelihood of colonizing artificial ponds (Buxton et al., 2018). While it is difficult to imagine commercial trade in Spring Peepers, Enge (2005a) reported that 2,361 P. crucifer ­were collected for the pet trade from 1990 to 1994 in Florida.

Pseudacris feriarum (Baird, 1854) Upland Chorus Frog

Synonyms: Helocaetes feriarum, Chorophilus feriarum, Chorophilus nigritus feriarum, Hyla feriarum, Pseudacris nigrita triseriata (in part), Pseudacris triseriata feriarum ­There is considerable taxonomic confusion in the lit­er­a­ture on chorus frogs. In the past, authors may have assigned a name to a chorus frog population ­under study that has no resemblance to the current nomenclature based on molecular analy­sis. For example, Smith and Smith (1952) compared vari­ous morphological characteristics of “Pseudacris nigrita feriarum” with “P. n. triseriata.” Upon looking at their collecting localities, it is clear that what we now know as P. feriarum, P. fouquettei, P. triseriata, and P. kalmi ­were all included ­under their name P. n. feriarum, and that P. fouquettei, P. triseriata, and P. maculata ­were included ­under

ETYMOLOGY

feriarum: Beltz indicates that the name is derived from the Latin feriarum meaning ‘holidays’ or ‘leisure.’ However, I suggest Baird was more likely referring to the Latin ferrum, meaning ‘of iron,’ ‘sword.’ The name would thus refer to the parallel, iron-­colored stripes down the back of the frog. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pseudacris) feriarum

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their name P. n. triseriata. In another example, Platz (1988a, 1989) analyzed calling characteristics of several species of striped chorus frogs from throughout their range, using the then current subspecific designations. He noted significant differences in many call characteristics, and suggested a revision of the subspecies. Based on ge­ne­tic evidence (Lemmon et al., 2007a), Platz’ species now are known to have included P. maculata, P. fouquettei, P. feriarum (possibly in western Kentucky), and P. kalmi. Always check the location where individuals originated to verify correct taxonomic allocation. The name P. triseriata has often been used for this species in the published lit­er­a­ture, and a number of authors have discussed its confusing nomenclatural history (e.g., Brown, 1956). In the following sections, I have based accounts only on studies with precise locations in order to ensure proper taxonomic allocation. IDENTIFICATION

Adults. This is a small, ground-­dwelling frog that is tan, medium to dark brown, or gray, with 3 dark parallel bands down its back. Postmetamorphs have a dark triangle between the eyes, but this trait is often lacking in Coastal Plain populations from South Carolina. The upper lip has a white line, ­there may be a dark band extending through the eye and continuing to just above the forelimb, and the belly is white to cream in contrast to the darker dorsal color; the belly may also have small brown flecks. ­There may be some dark coloration on the chest. The toes have almost imperceptible expanded pads, and webbing between toes is virtually lacking. Some individuals may completely lack dorsal markings. During the breeding season, males have a brown to yellowish coloration on the throat where the vocal sac is located. Males are slightly smaller than females, with South Carolina Coastal Plain populations having the largest frogs. Adult males are 23–31 mm SUL, whereas in North Carolina females are 25–33 mm SUL (Alexander, 1966). Mitchell (1986) gives maximum sizes of 31 mm SUL for males and 32 mm SUL for females in ­Virginia, and Brown (1956) recorded a size range of 22–31 mm SUL (mean 28 mm) in Alabama. In West ­Virginia, males reach 24.9 mm SUL, whereas females reach 28.7 mm SUL (Sias, 2006). Pennsylvania males average 24.1 mm SUL (range 20–27 mm) and females average 25.1 mm (23–27 mm) (Schwartz, 1957). South Carolina Coastal Plain P. feriarum males are 25– 35 mm SUL and females are 33–39 mm SUL; males from the Piedmont are smaller (21–29 mm SUL), as are females (22–31 mm) (Schwartz, 1957). An albino adult was reported in ­Virginia (Ackroyd and Hoffman, 1946; Hensley, 1959). Larvae. The tadpoles are small and olive, bark brown, or nearly black, and they have gold flecking uniformly across

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the dorsum; they have a whitish or silver-­to-­bronze coloration on the belly. The tail is medium in length. Tail fins may have small dark flecks, and the dorsal tail fin is slightly higher than the ventral tail fin. The snout is rounded when viewed from the side. Tadpoles reach 34–36 mm TL prior to metamorphosis. The mouthparts are figured by Dodd (2004). Eggs. The eggs are white to yellow on the vegetal pole, with darkly pigmented brown hemi­spheres. They are contained in a loose irregular jelly mass about 25 mm in dia­meter attached to submerged stationary objects such as leaves, twigs, and branches. Egg masses are usually located within 25 cm of the ­water’s surface. Egg counts in the lit­er­a­ture vary considerably. Alexander (1966) recorded 4–159 eggs per mass, whereas Mitchell (1986) gave a figure of 600 as maximum clutch size. In ­Virginia, Mitchell and Pague (2014) counted a mean of 441 eggs (range 197–835), and Gibson and Sattler (2020) reported a single clutch of 476 eggs. Brown (1956) counted 10–40 eggs per mass. A full breeding season complement of eggs ­will comprise many clusters, with a pos­si­ble output of about 1,500 eggs during a breeding season. Eggs contain only a singular gelatinous envelope 5–7.8 mm in dia­meter; the vitellus is 0.9–1.3 mm (Livezey and Wright, 1947; Brown, 1956). DISTRIBUTION

Upland Chorus Frogs are found in the Piedmont and in the Ridge and Valley Province and foothills from central Pennsylvania southward in an arc to northwest Mississippi,

Distribution of Pseudacris feriarum

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western Tennessee, and central Kentucky. Although generally found in the Piedmont, populations in Georgia follow the Savannah and Altamaha rivers into the Atlantic Coastal Plain in the east (Stevenson and Chandler, 2017), down the Chattahoochee and Flint river drainages into the Gulf Coastal Plain, and along the lower reaches of the Apalachicola River in Florida. In North Carolina, they follow the Cape Fear River southeast ­toward the Coastal Plain. They barely enter southern Illinois, but not in the area indicated by Cagle (1942). They occur west of the Mississippi River in extreme northeastern Arkansas and the Missouri boot heel. A rec­ord from Lafayette County, Arkansas (Black and Dellinger, 1938), is not valid. ­These frogs occur in east Tennessee and southwestern ­Virginia in the Tennessee River Valley and Smoky Mountain foothills, but appear to be sparse or absent from the Cumberland Plateau and Allegheny Mountains. They follow the Apalachicola and Yellow rivers south into the Florida Panhandle (Mays, 2016), thus verifying the observations by Carr (1940a) and Neill (1949b). A disjunct population of P. feriarum occurs in Berkeley, Dorchester, and Charleston counties, South Carolina (Schwartz, 1957; Lemmon et al., 2007b). ­Because of previous taxonomic changes, the best reference on distribution is the systematic study by Lemmon et al. (2007b). Other useful references include: Alabama (Mount, 1975), Florida (Bartlett and Bartlett, 1999; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Illinois (Phillips et al., 1999), Mary­land (Harris, 1975; Cunningham and Nazdrowicz, 2018), Mid-­Atlantic (Beane et al., 2010), Missouri (Johnson, 2000), North Carolina (Dorcas et al., 2007), Pennsylvania (Hulse et al., 2001), South Carolina (Schwartz, 1957; Fields, 2019), Tennessee (Redmond and Scott, 1996; Niemiller and Reynolds, 2011), ­Virginia (Tobey, 1985; Mitchell and Reay, 1999), and West ­Virginia (Sias, 2006). FOSSIL REC­O RD

No fossils have been specifically attributed to this species or to any member of the striped trilling frog species complex within the known range of P. feriarum. SYSTEMATICS AND GEOGRAPHIC VARIATION

Pseudacris feriarum is a member of the Trilling Chorus Frog clade of the genus Pseudacris, a group that includes P. brachyphona, P. brimleyi, P. clarkii, P. collinsorum, P. fouquettei, P. kalmi, P. maculata, P. nigrita, and P. triseriata (Moriarty and Cannatella, 2004; Barrow et al., 2014; Banker et al., 2020). ­These species are distinguished by their advertisement call structures, color patterns, ge­ne­tics, the presence of a cuboidal intercalary cartilage (Paukstis and

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Brown, 1987), and similar albumin phylogeny (Maxson and Wilson, 1975). Upland Chorus Frogs are the ­sister taxon of Western Chorus Frogs. ­These 2 species plus P. kalmi are the ­sister taxa of P. nigrita and P. fouquettei. Initially, Lemmon et al. (2007b) suggested that ­there ­were 2 geo­graph­i­cally separate lineages within P. feriarum, an inland and a coastal lineage roughly separated by the Altamaha River in Georgia. Further analyses, however, have identified 5 monophyletic lineages resulting from 2 to 5 parallel invasions and contact with P. nigrita that are correlated with 5 separate southeastern river systems: Escambia, Apalachicola, Altamaha, Edisto/ Santee, and James/Anna. Natu­ral hybridization occurs with P. brachyphona in northeast Mississippi and south central Kentucky (Lemmon et al., 2007b). Fouquette (1975) found evidence of call-­ based character displacement and no evidence of a hybrid zone at a contact zone between P. nigrita and P. feriarum along the Apalachicola River, despite low levels of hybridization (Crenshaw and Blair, 1959). This result is not surprising since ­these are nonsister taxa (Lemmon et al., 2007b). In contrast, Gartside (1980) found evidence of high levels of hybridization between P. nigrita and P. fouquettei along the Pearl River. Since ­these are closely related taxa, the level of hybridization at the contact zone between them is more easily explained than it would be without the phyloge­ne­tic interpretation of Lemmon et al. (2007b). Note that the microsatellite-­based estimates of hybrid frequency in Lemmon and Juenger (2017) are incorrect (see Banker et al., 2020). In the laboratory, P. feriarum readily hybridizes with P. nigrita, P. ornata, P. brimleyi, and P. brachyphona, and it does not appear to make much difference ­whether the P. feriarum is male or female (Mecham, 1965). However, crosses with P. crucifer are much less successful, but more so when the P. feriarum is female. This species can be hybridized with P. clarkii, and some larvae ­will metamorphose. Crosses with Acris crepitans (= A. blanchardi) ­were unsuc­ ying prior to hatching (Littlejohn, 1961a). cessful, usually d ADULT HABITAT

During the nonbreeding season, Upland Chorus Frogs are found in nearby swamp forest, moist forests, or river bottomland forest. They may be pre­sent in riparian corridors in other­wise partially developed habitats (Barrett et al., 2016). They tend not to disperse very far from breeding sites when ­these sites are in proximity to nonbreeding forested habitats. However, they ­will cross an extensive amount of open area to reach breeding sites in grassy marshes and temporary wetlands surrounded by grasslands. They inhabit the leaf litter, and shelter ­under surface debris, tree bark on the ground, or in small animal burrows.

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TERRESTRIAL ECOL­O GY

Upland Chorus Frogs spend most of the year in swamp or moist forests, where they forage in the leaf litter. They are usually well hidden in thick vegetation and are rarely encountered outside the breeding season, leading authors to speak of their “disappearance.” Despite this, they are likely active throughout the warmer parts of the year and may be active year-­round in the southern portion of their range. For example, Dodd (2004) found occasional individuals in a drying wetland from July to September in Tennessee, and Alexander (1966) found food in stomachs all year in North Carolina. Individuals forage on moist shaded forest floors or semifossorially, and they hide ­under surface debris. The species is capable of extensive terrestrial movements, and Ferguson (1963) found that displaced Upland Chorus Frogs ­were capable of correct orientation from the release site, moving 422 m to the site of original capture. They have been reported from cave entrances (Niemiller et al., 2016). Upland Chorus Frogs use sun-­compass (i.e., celestial or Y-­axis) orientation to direct their movements to breeding sites, as well as auditory cues (Ferguson, 1963, 1966a, 1966b). Indeed, auditory cues may be more impor­tant in finding breeding ponds than celestial cues since most movement occurs at night or on overcast days. Migrating frogs are capable of locating a breeding site by moving ­toward calling conspecifics. As they move ­toward the site, they ­will occasionally utter a call. However, if a chorus is interrupted, they ­will call more frequently, as if waiting for a response. Martof (1962a) noted their ability to discriminate odors from dif­fer­ent habitats, suggesting a third potential orientation mechanism, but this has not been studied since his observations. Ferguson’s (1966b) experiments on celestial orientation eliminated odor as a potential cue. Like P. nigrita, Upland Chorus Frogs prob­ably have a blue-­mode phototactic response, indicating they have true color vision (Hailman and Jaeger, 1974). CALLING ACTIVITY AND MATE SE­L ECTION

The call of the male Upland Chorus Frog sounds like ­running a fin­ger along the tines of a pocket comb. General characteristics of the call include a mean duration of 0.8 sec (range 0.5–1.25), a mean interval of 1.2 sec between calls, notes averaging 17 in number (range 15–23), and a dominant frequency of 2,800 cps (range 1,100–3,100) (Thompson and Martof, 1957; Jensen et al., 2008). Notes tend to be closer together at the beginning of the call compared with the end of the call. Martof and Thompson (1964) produced a series of “artificial” calls by altering call interval, intensity, duration, frequency, and uniformity and number of notes. They found that the number and spacing of notes and

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patterns of spacing and frequency ­were not necessary by themselves to induce proper choices by Upland Chorus Frog females. Calls needed to be at least 0.5 sec in duration with short intervening pauses for correct discrimination, but no single characteristic of the call was effective in eliciting discriminatory be­hav­ior. Mecham (1961) has discussed differences between the call of this species and that of P. nigrita. On the Coastal Plain in areas of overlap between the species, the call is fast and shorter (43 pulses/sec, duration of 0.62 sec) than in areas of nonoverlap (22 pulses/sec, duration of 0.75 sec). Coupled with changes in the call of P. nigrita in the zone of overlap, ­these differences may help maintain premating isolating mechanisms. Females exert strong se­lection in terms of mate choice, preferring conspecific males over heterospecific males, especially in areas of contact with other members of the Trilling Chorus Frog clade. Signal diversification occurs in ­these areas of sympatry, especially in terms of pulse rate and the number of pulses, with P. feriarum males increasing their calling effort when compared to areas where sympatry does not occur. In areas of sympatry, females have evolved a greater within-­species discriminatory ability as a consequence of between-­species discrimination. Such directional se­lection is metabolically costly to the males ­because of the increased energy expenditure needed to increase the number and rate of pulses. As a result, males may have to reduce the amount of time spent calling or adopt other be­hav­ior that ­will increase their fitness, despite the cost. Lemmon (2009) discusses ­these options in detail. Males that are amplexed by another male give off a harsh and penetrating warning call that ­causes the conspecific male to let go. Males ­will strug­gle to escape the grasp of a conspecific, often appearing to wrestle the intruder. When males approach one another, they become motionless at about 30 cm apart, appearing to watch one another. ­After no longer than about 6 min, 1 of the males ­will turn away from the other male. Male–­male interactions have been described by Martof (1958) ­under laboratory conditions. Calling occurs from late winter to early spring (e.g., Steen et al., 2013), and Upland Chorus Frogs are among the first frogs heard calling each year. Occasional calls may be heard into the summer or autumn, well past the breeding season. For example, Burt (1933) and Gibson and Sattler (2010) recorded an individual calling in June, Murphy (1963) heard calls in late autumn, and Mount (1975) noted calling in summer during cool rains. The timing of calling is variable and dependent on environmental conditions. Peak breeding occurs in December and January in South Carolina (Schwartz, 1957). In ­Virginia, calling occurs from January to April, but activity begins in December and rare calls have been heard

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even in October (Gibson and Sattler, 2020). Scattered calls ­were heard by Alexander (1966) from October to May in North Carolina, but full breeding choruses occurred usually from January to March, and rarely in December. In Mississippi, calls are heard from late November through mid-­ March (Landreth and Ferguson, 1966a), whereas calling occurs from January to April in Tennessee (Burkett, 1991). Snow and ice may remain on the ground and in ponds when breeding begins. Chorusing usually occurs for several days prior to the onset of breeding. Calling occurs throughout the day, with peaks around 12:30, 18:00, and 24:00 hrs; the least calling occurs around 08:00 hrs (Todd et al., 2003; Steelman and Dorcas, 2010). The intensity of calling does not occur synchronously even between closely situated breeding sites; calling intensity may be high at 1 location yet greatly decreased at the next, and calling may extend longer in the season at 1 location than another. Temperature has a direct influence on calling be­hav­ior. Calling is initiated at ca. 10ºC air temperature and at 6ºC ­water temperature, but it is infrequent at ­these cold temperatures. Strong choruses are heard when ­water temperatures reach 6.5ºC, even when air temperatures remain as cold as 5ºC. As temperatures warm to 7–10ºC, chorusing occurs both day and night and continues as long as temperatures remain ca. 10–25ºC. Calls are heard from ca. 5 to 6.0 (Turner and Fowler, 1981), and in restored or mitigation wetlands (Palis, 2007) and retention ponds (Birx-­Raybuck et al., 2010).

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Pseudacris fouquettei Lemmon, Lemmon, Collins, and Cannatella, 2008 Cajun Chorus Frog ETYMOLOGY

fouquettei: The specific epithet fouquettei is a patronym honoring Martin J. (Jack) Fouquette (1930–2014) of Arizona State University, who conducted extensive studies on chorus frogs in the 1960s and 1970s. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pseudacris) fouquettei This morphologically cryptic species has been referred to in the lit­er­a­ture as Pseudacris triseriata, P. nigrita, P. feriarum, or even P. clarkii, depending upon the date of publication and location within its range. For example, Platz (1988a, 1989) analyzed calling characteristics of several species of striped chorus frogs from throughout their range using the then-­current subspecific designations. He noted significant differences in many call characteristics and suggested a revision of the subspecies. Based on ge­ne­tic evidence (Lemmon et al., 2007a), Platz’ species now are known to have included P. maculata, P. fouquettei (his P. feriarum), P. feriarum, and P. kalmi. Among the authors that have discussed the confusing nomenclature of frogs now referred to as P. fouquettei is Bragg (1943a), who recognized early on the distinctness of P. fouquettei (as P. triseriata) from P. clarkii. Always check the location where individuals originated to verify correct taxonomic allocation.

Oklahoma averaged 25.8 mm SUL (range 22–28 mm) for males and 28.3 mm SUL (range 24–31 mm) for females (Goldberg, 2021a). Lemmon et al. (2008) provided a series of photo­graphs showing variation in color and stripe pattern in comparison with some of the other trilling chorus frogs. Males have a yellowish-­orange or dark gray coloration in the area of the subgular vocal sac. Larvae. Like other members of the Trilling Chorus Frog clade, the tadpoles are small and dark brown or gray to black. Hatching occurs at 6–6.5 mm TL. The snout is rounded in lateral view, and ­there is a light line posterior to the eye. Tail fins may be pigmented via fine mottling. The dorsal tail musculature is darkly pigmented, whereas the ventral portion of the tail musculature is not pigmented. The tail is about twice as long as the body. The maximum tadpole size is 36–39 mm TL (Siekmann, 1949). Siekmann (1949) described larvae in greater detail, including tooth formulas and variation in oral morphology. Eggs. Eggs are gray brown to deep brown, with white to cream on the vegetal pole. Bragg (1948) noted that eggs in turbid ­waters ­were brown, whereas eggs in clear ­water ­were much darker. Siekmann (1949) noted light-­tan-­colored eggs in Louisiana. Eggs are deposited in a loose, irregular, oblong cluster of about 25 mm in dia­meter, and a full breeding season complement of eggs ­will comprise many clusters of 8–300 eggs each. Females produce approximately 1,500 eggs during a breeding season, perhaps during 2 separate periods of amplexus. Eggs contain only a singular gelatinous envelope mea­sur­ing 3–6.1 mm in dia­meter (mean 4.6 mm); the vitellus is 1.2–1.5 mm (Livezey, 1952). Egg mea­sure­ ments in Smith (1934) and Livezey and Wright (1947) refer to other members of the Trilling Chorus Frog clade or confuse species.

IDENTIFICATION

Adults. The Cajun Chorus Frog is a small, slender, tan to medium brown member of the Trilling Chorus Frog clade (Barrow et al., 2014). Other than by ge­ne­tic data (Lemmon et al., 2008) and distribution, it can be identified by its subacuminate snout, a pattern of 3 medium brown to dark brown longitudinal dorsal stripes or rows of spots on a pale tan to gray background, and the presence of a white iridescent labial stripe. The head is slightly narrower than the body and the tympanum is distinct. The arms are long and robust, and the legs are slender and of moderate length. The toe tips are only slightly wider than the digits. Dorsally, the skin is weakly granular, but ventrally it is noticeably granular. The belly is cream colored with scattered dark flecks, and the throat is yellowish brown. Lit­er­a­ture rec­ords suggest males (maximum 30 mm SUL) are slightly larger than females (maximum 27 mm SUL), with adults ranging from 22 to 30 mm SUL. However, museum specimens from

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DISTRIBUTION

The Cajun Chorus Frog is found from central Oklahoma near the border with Kansas southward through central Texas. It occurs throughout the states of Arkansas and Louisiana, and in southwest Mississippi. It does not occur farther east than the Pascagoula–­Leaf River system in southern Mississippi. The species barely enters southern Missouri. Impor­tant distributional references include Lemmon et al. (2008) as well as: Arkansas (Trauth et al., 2004), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Oklahoma (Sievert and Sievert, 2006), and Texas (Smith and Sanders, 1952; Hardy, 1995; Dixon, 2000, 2013). FOSSIL REC­O RD

No fossils have been specifically attributed to this species. However, fossil P. triseriata have been described from Rancholabrean sites in Oklahoma (Holman, 2003). The

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Pseudacris fouquettei 347

­There is a considerable amount of variation in the patterning of stripes and spots on the dorsum of this species. Lemmon et al. (2008) noted strong three-­stripe patterns, a three-­stripe pattern with dark spots bounding the stripes, a broken three-­stripe pattern, and even patterns with no stripes at all, except for markings on the legs; vari­ous numbers of transverse bars (2–15) are pre­sent on the legs. Bragg (1948) noted that ­there was equal repre­sen­ta­tion between striped and spotted dorsal patterns in eastern Oklahoma. Although most frogs have a cream-­colored venter, some have gray pigment. ADULT HABITAT

Distribution of Pseudacris fouquettei

major lineages within the genus Pseudacris ­were established prior to the age of the fossil deposits. Therefore, ­these fossils are within the range of P. fouquettei as currently understood (Lemmon et al., 2007a). SYSTEMATICS AND GEOGRAPHIC VARIATION

Pseudacris fouquettei is a member of the morphologically conservative Trilling Chorus Frog clade of the genus Pseudacris, a group that includes P. brachyphona, P. brimleyi, P. collinsorum, P. clarkii, P. feriarum, P. kalmi, P. maculata, P. nigrita, and P. triseriata (Moriarty and Cannatella, 2004; Banker et al., 2020). ­These species are distinguished by their advertisement call structures, color patterns, the presence of a cuboidal intercalary cartilage (Paukstis and Brown, 1987), and ge­ne­tics. Cajun Chorus Frogs are the ­sister taxon of Southern Chorus Frogs. Together, ­these 2 species are the ­sister taxa of a group that includes P. triseriata, P. kalmi, and P. feriarum. Natu­ral hybridization occurs with P. nigrita in southeast Mississippi along the Pearl River floodplain (Gartside, 1980; Lemmon et al., 2007b). Smith and Smith’s (1952) intergrades between P. n. feriarum and P. n. triseriata are, in part, based on contact zones between P. fouquettei and P. maculata. Burt (1936) recorded intergradation between “P. triseriata” and P. clarkii in southern Kansas on the Oklahoma border and perhaps in east Texas; ­these instances would involve P. fouquettei and P. maculata in the former, but it is unclear ­whether intergrades actually occur in the latter case. U ­ nder laboratory conditions, P. fouquettei readily hybridizes with P. clarkii (Lindsay, 1958, but see Lord and Davis, 1956).

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Pseudacris fouquettei generally is a frog of the moist forests, woodlands, and savannas within its range, as opposed to P. clarkii, which occurs primarily in prairies and grasslands. However, Bragg (1943a) found P. fouquettei in tall-­grass prairie and mixed-­grass prairie in Oklahoma. The species requires open-­canopied breeding sites in proximity to woodlands. TERRESTRIAL ECOL­O GY

­ ittle is known of the terrestrial ecol­ogy of this species. It is L likely a surface-­dwelling frog that forages in the moist leaf litter during the nonbreeding season, although a semifossorial existence cannot be excluded. For example, Black (1973a) found a Cajun Chorus Frog in a limestone crack near a cave entrance. ­These frogs are rarely encountered outside the breeding season. Like P. nigrita, Cajun Chorus Frogs prob­ably have a blue-­mode phototactic response, indicating they have true color vision (Hailman and Jaeger, 1974). CALLING ACTIVITY AND MATE SE­L ECTION

Males precede females to the breeding sites by about a day or so, and ­there is usually a slight time lag from when males are first heard calling ­until the first eggs are deposited. Males call from the ­water within 5–20 cm of the shoreline and are usually spaced around a breeding site, although a few may call from the bank. They generally face ­toward the pond or wetland while calling from very shallow ­water. Males sit with about three-­quarters of their body out of the ­water, with the rear legs folded and floating at right ­angles to the body. The front limbs are used to maintain balance on the substrate or vegetation. This posture essentially thrusts the male upward as he emits his call, and it is maintained throughout the calling bout over a period of hours. If the frog relaxes his posture, he slips down into the ­water with his back legs under­neath his body. This altered posture allows him to jump should a predator approach, or to approach a rival. Wiest (1982, as P. triseriata) noted calling at -2.5–22°C over an 84 day period in Texas. In northeast

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348  Hylidae

Texas, calling extends from January to March (Car­ter et al., 2018, as P. triseriata). The Cajun Chorus Frog has a trilling call somewhat similar to ­running a fin­ger over the tines of a comb. The species has a slower mean call rate (0.34 calls/sec, range 0.14–0.43 calls/ sec) than allopatric P. feriarum and P. maculata, a higher mean call-­duty cycle (0.36, range 0.26–0.44) than P. nigrita, a long mean call length (1,115.4 ms, range 867–1,554 ms), and a mean pulse number (13.1, range 9.9–15.7) intermediate between P. maculata, P. feriarum, and P. nigrita. The mean dominant frequency is 3,138 Hz (range 2,846–3,900 Hz) with a duration of 0.74–0.85 sec (Michaud, 1962, 1964; Lemmon et al., 2008). As for frogs in general, temperature affects vari­ous par­ameters of Cajun Chorus Frog calls. For example, the duration of the call and the interval between calls are inversely proportional to temperature. The trill rate increases with temperature, but the number of trills remains constant regardless of temperature. Even in areas of sympatry with other members of the Trilling Chorus Frog clade, the call of P. fouquettei is distinctive and does not deviate from its basic characteristics, suggesting no evidence of character displacement (Michaud, 1964). In choice tests, female P. fouquettei ­favor the calls of conspecific males over ­those of P. clarkii (Michaud, 1962). Males stop calling ­after amplexus, but ­there is 1 report of a male calling ­under laboratory conditions while still in amplexus (McCallum and Trauth, 2009). BREEDING SITES

Cajun Chorus Frogs breed in shallow temporary pools and wetlands, similar to ­those of other members of the Trilling Chorus Frog clade. They may be found in vari­ous habitat types, from forested areas to open fields and grassy swales, and even in shallow roadside ditches or cultivated fields. Livezey (1952) also recorded them breeding in deep semipermanent ponds, and McKnight and Ligon (2016) reported calling from beaver-­impounded lakes. Breeding sites are characterized by having an open canopy that allows sunlight to warm the shallow ­water, especially since ambient temperatures during the breeding season often are quite cold, especially at night. The wetland substrate may be covered with grasses or vegetation, or rotting detritus may be pre­sent. Most breeding ponds are small, although Livezey (1952) mea­sured 1 that was 10 m by 25 m. REPRODUCTION

Breeding normally occurs from December to May, depending on location and environmental conditions, although the initiation of the reproductive cycle begins in November, at least in Oklahoma (Goldberg, 2021a). In that regard, Livezey (1952) also suggested that breeding activity begins as early as

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November in east Texas, and Dundee and Rossman (1989) reported calling in late October in Louisiana. Goldberg (2021a) noted mature females in March, April, May, June, and November. Most reproduction occurs early in the breeding season rather than throughout the winter and spring, and it may occur on only a few days. Initially, and as the season progresses, occasional egg masses are found, but they are usually few. Goldberg (2021a) also noted that some females had ovaries in spawning condition in addition to postovulatory follicles, also suggesting that spawning occurs more than once in a breeding season in some individuals. Breeding patterns are variable and weather dependent, and the breeding phenology 1 year may be quite dif­fer­ent from the following year. Breeding frogs prefer ambient temperatures from 4 to 24°C (with most calling occurring on warmer nights, McKnight and Ligon, 2016), and rainfall usually triggers calling and mating. As noted by Livezey (1952), reproduction often occurs as temperatures rise and rain falls following a previous sudden drop in temperature. Liner (1954) noted an instance in Louisiana where snow and ice ­were on the ground as Cajun Chorus Frogs ­were breeding on 7 February, and Livezey (1952) observed intense breeding as temperatures ­rose following an 18 cm snowfall. Most egg deposition occurs before minimum temperatures are about 10°C. Amplexus occurs in ­water. A female ­will approach the vicinity of a calling male, at which point he ­will swim ­toward her and grasp her in an axillary position. Females control the movement of the mated pair. She may remain in the vicinity of the male’s calling site, or she may move elsewhere within the wetland. Females deposit their eggs in shallow temporary wetlands and pools, and the exact oviposition site is chosen by the female (Livezey, 1952). Prior to deposition, the female ­will grasp a twig with her front limbs, with the hind limbs ­free and somewhat off to the side. Egg deposition is preceded by abdominal contractions, which alternate from side to side. Abdominal contractions in conjunction with the female arching her back eject the eggs slightly upward ­toward the male’s cloacal opening. Eggs are emitted in short strings, with a mass being composed of eggs emitted over at least 3 abdominal contraction sequences. The eggs are attached to slanting debris such as twigs, sticks, or branches, and usually not to vegetation such as sedges or grasses. As the eggs are extruded, the female climbs upward along the stem she’s holding in order to allow for more eggs to be deposited. As eggs are extruded, the male squeezes the female and hunches his body ­toward hers, assuming a concave position, moving his vent ­toward hers. As eggs are extruded, they are fertilized. More than 1 egg mass may be attached to a branch. The egg mass is placed 2.5–10 cm below the ­water’s surface. Egg masses may be clumped in distribution, or they may be scattered throughout

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Pseudacris fouquettei 349

the wetland depending on ­water depth and the spacing of available branches. Livezey (1952) provides a comprehensive description of mating be­hav­ior and egg deposition. Pseudacris fouquettei females may deposit up to 9 egg masses during a single night on a variety of attachment substrates. This pro­cess takes up to about 3 hrs (Livezey, 1952); a considerable time (12–60 min) may pass between successive ovipositions. Indeed, the number of egg masses produced by a female may be dependent on the number of attachment sites. When few attachment sites are available, ­there ­will be fewer and larger egg masses; when attachment sites are plentiful, egg masses are smaller and more scattered (Bragg, 1948; Livezey, 1952). Egg masses range from a few to dozens, depending on the size of the breeding site. Livezey (1952) found from 3 to 58 egg masses per site, even in wetlands as small as 80 days, depending

Nearly all aspects of the population biology of this species are unknown. Goldberg (2021a) noted the smallest males in reproductive condition ­were 22 mm SUL, whereas the smallest mature females ­were 28 mm SUL.

DIET

­ ittle or no food is consumed by juveniles ­until the tail has L been completely resorbed. Livezey (1952) recorded copepods, springtails, beetles, and ­water mites from juveniles, and beetles, fly larvae, lepidopteran larvae, and true bugs from the stomachs of adults. Adults consume shed skin. It seems likely that Cajun Chorus Frogs eat any small invertebrate they can capture. PREDATION AND DEFENSE

Observations of predation on this species are nearly non­ex­is­tent. Dundee and Rossman (1989) reported that a Garter Snake (Thamnophis proximus) regurgitated a Cajun Chorus Frog. The species prob­ably relies on its cryptic coloration and jumping ability, especially in cold weather, to avoid predators. A calling male also is very difficult to locate, due to the ventriloquist-­like nature of the call. Adams et al. (2011) found that larvae are unpalatable to some species, although the effectiveness of larval unpalatability is variable. Schiwitz et al. (2020) found no effect on larval activity in the presence of predator chemical cues or conspecific alarm cues.

COMMUNITY ECOL­O GY

Tadpole of Pseudacris fouquettei. Photo: Stan Trauth

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Cajun Chorus Frogs occur in the same pools with P. clarkii in central Pontotoc County, Oklahoma, in a narrow band where ­these species come into contact. According to Bragg (1943a), the transition between ­these species is rather abrupt with ­little geographic overlap. A similar area of contact once occurred just east of Dallas, Texas, but this area has been highly disturbed in recent years. The exact nature of interspecific interaction is unknown, but differences in

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350  Hylidae

Breeding habitat of Pseudacris fouquettei. Photo: Chris Brown

Adult Pseudacris fouquettei, amplexus. Photo: James Beck

female call preference and male mating calls prob­ably serve as effective isolating mechanisms. DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus (Batrachochytrium dendrobatidis) has been found in individuals from Louisiana (Rothermel et al., 2008), Oklahoma (Watters et al., 2016, 2019; Marhanka et al., 2017), and Texas (Saenz et al., 2010). Ranavirus is reported from P. fouquettei in Oklahoma (Davis et al., 2019). Parasites include the protists Opalina sp. and Nyctotherus cordiformis, the cnidarian Cystodiscus melleni, the trematodes Brachycoelium salamandrae, Glypthelmins quieta, Megalodiscus temperatus, Mesocoelium sp., M. monas, and Renifer metacercaria, the cestode Cylindrotaenia americana, and the nematodes Cosmocercoides variabilis, Physaloptera sp., Oswaldocruzia leidyi, and O. pipiens (McAllister et al., 2013, 2015a). SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. Briggler (1998) reports calling from a pond with a pH of 6.0. It is likely that this species has an acid tolerance similar to P. kalmi, that is, about 4.0.

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Plant extracts. In mesocosm experiments, larval P. ­fouquettei had lower survivorship and ­were smaller when raised on Chinese tallow (Triadica sebifera) leaf litter than larvae raised on litter from native tree species (Cotten et al., 2012). STATUS AND CONSERVATION

The Cajun Chorus Frog tolerates silviculture relatively well, as long as forest cover is retained during the nonbreeding part of the year. This species prefers breeding in open-­ canopied temporary wetlands of a type that form in clearcut and selectively cut areas. As such, they are frequently found in clearcuts, sometimes even far from the nearest forest cover. In such areas, they prefer flooded, brushy areas with sparse overstory (Fox et al., 2004). As trees are cut, the ­water ­table tends to rise, providing numerous flooded pools, especially for a few seasons ­after the cut. Intensive forest management may actually benefit this species by providing open breeding sites in a mosaic of habitats. Pseudacris fouquettei rarely has been observed calling from permanent constructed ponds in forested habitats (Briggler, 1998). It is unlikely that such ponds would be a significant asset to maintenance of Cajun Chorus Frog populations ­unless hydroperiod could be varied and fish excluded. However, this species readily colonized temporary, open-­canopied, fishless, artificial ponds created by military activity (Ecrement and Richter, 2017), and occupied created ephemeral wetlands in tall-­grass prairie in northwest Arkansas (Baecher et al., 2018).

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Pseudacris illinoensis 351

Pseudacris illinoensis Smith, 1951 Illinois Chorus Frog ETYMOLOGY

illinoensis: referring to the State of Illinois. NOMENCLATURE

Fouquette and Dubois (2014): Pseudacris (Pycnacris) streckeri illinoensis Synonyms: Hyla streckeri illinoensis, Pseudacris streckeri illinoensis, Pseudacris streckeri illinoisensis, Pseudacris triseriata illinoensis IDENTIFICATION

Adults. The Illinois Chorus Frog is a stout frog with a variable ground color of gray, tan, or brown, with dark brown or black marks on the dorsum. ­There is a distinct V-­shaped pattern between the eyes, a dark stripe from the snout to the shoulder, and a dark spot below each eye. Most individuals have a dark, inverted Y-­shaped mark on 1 or both shoulders. The forearms are enlarged and used for burrowing, and the front digits lack a terminal disk. ­There is no yellow groin coloration as ­there is in P. streckeri. Venters are white. ­There is virtually no webbing on the hind feet. Males in Arkansas had a mean SUL of 36.9 mm, whereas Illinois males averaged 35 mm SUL (Trauth et al., 2007). In another Arkansas study, males averaged 38 mm SUL (range 35–41), whereas females averaged 39 mm SUL (range 37–40) (Butterfield, 1988). Tucker (1995) reported an adult size of 32–43 mm SUL in Illinois. The maximum size is 48 mm SUL. Larvae. The brownish-­gray tadpoles are heavy bodied with a high tail fin containing many small, scattered markings. The total length is 38 mm. Johnson (2000) provided a figure of the tadpole and Smith (1951) an illustration of tadpole mouthparts. Eggs. The eggs are brownish gray dorsally and white ventrally. ­There is a single jelly envelope surrounding the egg. Eggs are 1.9–2.6 mm (mean 2.3 mm) in dia­meter (Butterfield, 1988). DISTRIBUTION

The Illinois Chorus Frog is known from 3 scattered populations in west central Illinois, southwest Illinois, and the junction of southern Illinois, southeastern Missouri, and northeastern Arkansas (Clay County). Impor­tant distributional references include Arkansas (S.E. Trauth et al., 2004; J.B. Trauth et al., 2006), Illinois (Smith, 1961; Holman et al., 1964; Brown and Brown, 1973; Axtell and Haskell, 1977;

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Distribution of Pseudacris illinoensis

Brown and Rose, 1988; Brandon and Ballard, 1998; Phillips et al., 1999), and Missouri (Smith, 1955; Johnson, 2000; Daniel and Edmond, 2006). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Pseudacris illinoensis was described as a subspecies of P. streckeri by Smith (1951). Differences between the subspecies ­were said to include a lack of bright coloration in the groin in P. illinoensis, a more uniform distribution of pigment, a reduction in the dark lateral stripe, a more general pallid color, and a smaller fifth row of labial teeth in the larvae (Smith, 1951). Except for the lack of groin coloration, variation in ­these other characters make them less useful in separating the species. Trauth et al. (2007) examined a number of morphological characters and identified differences among P. illinoensis and P. streckeri populations, but none ­were strong enough to support status as a separate species. Based on mtDNA, however, Moriarty and Cannatella (2004) and Pyron and Wiens (2011) concluded that specific status was warranted. The species is a member of the Fat Chorus Frog clade and is most closely related to P. streckeri of the south central United States (Hedges, 1986; Moriarty and Cannatella, 2004; Barrow et al., 2014; Banker et al., 2020). The species is likely a remnant of ancestral P. streckeri, that extended

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352  Hylidae

The Illinois Chorus Frog is found on river floodplains that contain sands or sandy soils deposited ­either by ­water or wind. It is a remnant of sand prairie habitat that once extended into Illinois but has largely been eliminated by agriculture. In areas where the sand prairies no longer exist, P. illinoensis may survive in agricultural fields. It does not occur on substrates that inhibit burrowing, such as sod (Brown et al., 1972). Many localities are associated with old glacial meltwater and river terraces that ­were deposited during the Pleistocene (Brown and Rose, 1988).

ground. The call is a short, high-­pitched, bird-­like whistle that is repeated rapidly. Calls are heard from March to April (33 days) in Illinois (Brown and Rose, 1988) and from February to April in Arkansas (Butterfield, 1988). Within a region, calling may occur on vari­ous nights so that calls may be heard in 1 area but not in an adjacent area. However, males call virtually ­every night in large choruses. Calling may be stimulated by heavy rainfall. Calling occurs at night early and ­later in the season (productive at 18:30–00:50 at air temperatures of 6–15°C) (Brandon and Ballard, 1998), but during peak breeding in early April, males call during the day. Males likely remain at or near a breeding site throughout the spring, with females returning to their burrows soon ­after oviposition.

TERRESTRIAL ECOL­O GY

BREEDING SITES

This species is largely fossorial, digging into deep sands where it spends most of the year, except when breeding. Burrows are made in open sandy habitats, and frogs remain 15–20 cm below the surface (Axtell and Haskell, 1977). The burrows are dug using synchronized movements of the forelimbs rather than the hind limbs; as a consequence, the forelimbs are stout and well muscled (Brown et al., 1972). It takes a frog 85–142 sec to complete a burrow. During the winter, Illinois Chorus Frogs must burrow beneath the frost line (>25 cm) ­because they are intolerant of freezing (Packard et al., 1998). Feeding likely occurs under­ground within burrows, an ability facilitated by the frog using its front limbs to catch and hold prey (Brown, 1978; McCallum and Trauth, 2001b). Traces of the burrows may be pre­sent on the surface of the substrate. Dispersing individuals tend to occupy old-­field habitats. Tucker (1998) found marked individuals a mean of 0.52 km (range 0–0.9 km) from their breeding pond, with females tending to disperse slightly—­but not significantly—­farther than males. Breeding individuals may not return to the same breeding site from 1 year to the next. Movement is by short toad-­like hops, quite unlike the jumps of other hylid frogs. Illinois Chorus Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), but with a slight departure from monotonicity at low frequencies. The blue-­mode response would help them orient ­toward areas of increasing illumination (Hailman and Jaeger, 1974). Illinois Chorus Frogs likely have true color vision.

Most of the natu­ral habitat of this species has been modified by agriculture for a long time. Breeding occurs in remnant shallow pools (600 m (Spencer, 1964a). Most frogs remain within 20– 50 m of their breeding site, but they have been reported to move 183–244 m over cutover upland habitat to reach adjacent wetland sites (Spencer, 1964a, c). Movements may ­favor a par­tic­u­lar direction, and a maximum distance between recaptures has been recorded as ca. 580 m for a movement rate >61 m/day (Spencer, 1964c). During the nonbreeding season, Boreal Chorus Frogs take refuge in rodent burrows, such as ­those of the pocket gopher Thomomys or the prairie dog, Cynomys, ­under surface debris (rocks, human-­created refuse), in thick grass clumps, and within the vegetation of wetlands. Population estimates are unknown, but in northern Alberta, Roberts and Lewin (1979) estimated ­there ­were 2.3 P. maculata per 1,000 m2. The end of the activity season is determined by the arrival of cold weather, which is latitude and elevation dependent. When weather permits, some frogs may be active at any time of the year. Specific rec­ords for autumnal activity include late September in Wisconsin (where individuals ­were observed in a streambed; Edgren, 1944), Kansas (where activity was observed in an agricultural field on a sunny day; Busby et al., 2005), and northern Alberta (Harper, 1931). Activity occurs through September at high elevations in Colorado (Spencer, 1964a) and October in Yellowstone (Koch and Peterson, 1995), but extends into November on the prairies (Hammerson, 1999; Geluso and Harner, 2013b). Tucker et al. (1995) observed this species active in Illinois in late November ­after being displaced by floodwaters. In Kansas, Taggart (in Collins, 1993) found an adult active on 23 December. According to Harper (1931), Boreal Chorus Frogs ­were easily observed and captured ­after the breeding season along the shores of wetlands and in marshes in northern Alberta, although most reports indicate they are uncommonly observed ­after the breeding season. The Boreal Chorus Frog occurs at much higher elevations (to 3,720 m; Spencer, 1971) and latitudes (65° N) than nearly all other North American anurans. As such, it must

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362  Hylidae

have a method for surviving cold temperatures for long periods. Overwintering occurs in animal burrows, ant mounds, and root channels below decaying stumps. Moving under­ground below the frost line may be an option in parts of the species’ range, but the presence of permafrost may limit vertical movement. Boreal Chorus Frogs survive freezing by producing a cryoprotectant in the form of glucose, which is made available via liver glycogen. The cryoprotectant allows extracellular ­water to freeze while preserving intracellular function. ­After about 2 hrs at freezing temperatures, glucose levels in the liver increase substantially (Edwards et al., 2000) as a result of a generalized stress response, the heart continues to beat, and the glucose is circulated throughout the body. The core body organs contain the most glucose and are the most protected. The use of glucose in conjunction with changes in plasma osmolality allow Boreal Chorus Frogs to survive temperatures of -3°C to -2°C for a considerable period (Hunka, 1974; MacArthur and Dandy, 1982), although survivorship is dependent on the level of supercooling prior to freezing (Swanson et al., 1996). Reserves of glycogen are ­limited, however, so repeated cycles of freezing and thawing could limit the ability of the frog to produce the cryoprotectant. However, Jenkins (2000) found no significant differences in glycogen levels in frogs exposed to 2 or 3 freeze-­thaw cycles, although her sample size was small. Survivorship may be low in some particularly cold winters, especially if liver glycogen levels are low. Laboratory studies have demonstrated that glycogen levels vary among frogs acclimated to cold and that survivorship is low when glycogen levels are low (Edwards et al., 2000; Jenkins, 2000). To what extent ­these levels vary in nature and by size class is undetermined. Although the Boreal Chorus Frog inhabits an environment that may be rather cool during the peak activity season, the CTmax for specimens acclimated at 5°C is 37.1°C, and at 20°C, it is 38.7°C (Miller and Packard, 1977). As elevation increases, the CTmax decreases so that frogs from higher elevations have a lower CTmax than frogs from low elevations, regardless of acclimation temperature. Clarke (1958) recorded a wide range of temperatures (4.4–32°C) at which this species is active. CALLING ACTIVITY AND MATE SE­L ECTION

The phenology of calling by Boreal Chorus Frogs depends on environmental conditions, elevation, vegetation, and latitude. Chorus Frogs have been reported calling during snowfall when air temperatures ­were -1ºC to 0.7ºC (Degenhardt et al., 1996; Bezy et al., 2004), but most calling occurs at >5°C (Brinley Buckley et al., 2021). Indeed, ice is not uncommon on breeding sites at the start of the breeding

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season. Other impor­tant predictors of calling activity are hydropattern and weekly precipitation. Calling begins in March in Arizona (Bezy et al., 2004) and Nebraska (and from then ­until May; Brinley Buckley et al., 2021). In Colorado, it occurs from April to mid-­June (Vertucci and Corn, 1996), when patchy ice still covers the breeding site, although most calling occurs from May to June (Corn and Muths, 2002). In southern Ontario, calling occurs from early to mid-­April (Piersol, 1913). In northwestern Ontario, calling begins in late April and continues through June (Cook, 1964a), whereas calls are heard May–­July in northern Manitoba (Harper, 1963), and into July in North Dakota (Bowers et al., 1998) and South Dakota (Peterson, 1974). Calls have been recorded at the southern end of James Bay, Québec, from late May to mid-­June (Ouellet et al., 2009). Occasional calls have been reported in August and mid-­September in northern Alberta (Harper, 1931), although breeding had long ceased by this time, and in October in South Dakota (Blais et al., 2015). The mean duration of the call of P. maculata from Churchill, Manitoba, is 578 ms (range 572–584 ms). The call has 18 notes and a dominant frequency of 3,069 cps (range 2,750–3,300) (Thompson and Martof, 1957). Platz (1989) found the number of pulses to vary from 14.3 to 17.4 in a transect from the northern to southern ­Great Plains; dominant frequencies varied clinally, from 2.91 (south) to 3.58 (north) kHz. Call duration (0.59–1.16 sec) and pulse rate (12.7–23.7 pulses per sec) also varied geo­graph­i­cally (Platz, 1989). The calls of this species also vary with elevation, particularly the dominant frequency (decreases with elevation; 3,500 to 20 m from a chorus, depending on the location of the recorder in relation to the chorus. Aural surveys conducted where frogs are distant from a listening site likely underestimate occupancy. The species is considered Threatened by COSEWIC.

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Smilisca baudinii  393

Smilisca baudinii (Duméril and Bibron, 1841) Mexican Treefrog ETYMOLOGY

baudinii: a patronym honoring Nicolas Thomas Baudin (1754–1803), a French commander in México who donated the type specimen to the Museum National d’Histoire Naturelle in Paris. NOMENCLATURE

Synonyms: Hyla baudinii, Hyla beltrani, Hyla manisorum, Hyla muricolor, Hyla pansosana, Hyla vanvlietii, Hyla vociferans, Smilisca daudinii, Smilisca daulinia IDENTIFICATION

Adults and juveniles. This is an attractive, pale green to tan treefrog with dark, irregular dorsal spots and a network of black-­on-­yellow lateral coloration. Heads are wide and flat, and snouts are rounded and short. ­There is a dark marking posterior from the eye that becomes a black vertical shoulder bar; a distinctive light yellow or green spot is pre­sent below the eye. A black interocular bar and a distinct creamy-­white anal stripe usually are pre­sent. The throat is greenish or yellow except in breeding males when it is gray. Limbs have 3–4 dark transverse bands. The rear of the thigh has greenish-­yellow to purplish-­russet reticulations. Toes have moderately large disks and are three-­fourths webbed. The vocal sac is paired and subgular. Venters are white. Males are smaller than females. Males are 47.3–75.9 mm SUL (mean 58.7) from throughout its range; females reach a maximum size of 90 mm SUL from Sinaloa (Duellman, 2001). Juveniles are dull olive green dorsally and white ventrally. ­There are faint brown transverse bars on the limbs. A distinctive white suborbital spot is pre­sent. Larvae. Tadpoles are dark brown with a pale crescent mark posteriorly and a body that is wider than deep. Snouts are rounded. Eyes are widely separated and located dorsolaterally. The caudal musculature is pale tan with a dark brown longitudinal streak. Prominent tail fin markings are absent, as are any indications of dorsal ­saddles; small brown flecks may be pre­sent, however. Tails are long in mature larvae, and tail fins are deepest at one-­third down the length of the tail. Venters are transparent, with brown flecks anteriorly below the eye. Larvae reach 35–37.5 mm TL. Tadpoles are described in detail by Duellman (2001). Eggs. Eggs have a dia­meter of ca. 1.3 mm and are encased in a single membrane of 1.5 mm in dia­meter (Duellman, 2001).

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Distribution of Smilisca baudinii

DISTRIBUTION

Smilisca baudinii occur from south Texas through Costa Rica. In the United States, this species occurs only along the lower Rio Grande Valley, with additional reports from 2 nearby counties to the north (Bexar, Rufugio), which prob­ably represent accidental individual introductions that never resulted in the establishment of breeding populations (Lott, 2016). Impor­tant distributional references include Dixon (2000, 2013), Duellman (2001), and Lott (2016). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Smilisca baudinii is a member of a tropical group of hylid frogs, the ­Middle American/Holarctic clade (Faivovich et al., 2005). The genus Smilisca reaches its northernmost distribution in Texas and southern Arizona. Duellman (2001) provided characteristics defining the genus, which includes at least 9 species. ­There is substantial morphological and color variation among S. baudinii populations over its considerable range, particularly with regard to maximum size and larval coloration. ­These have been discussed by Duellman (2001). ADULT HABITAT

The habitat in Texas is described by Wright and Wright (1938) as “on ­houses, in meadows, overflow lands, on trees of forests, in palm groves.” Throughout its range, it is a

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394  Hylidae

species of xeric and subhumid habitats that have a prolonged dry season. TERRESTRIAL ECOL­O GY

Smilisca baudinii occupies tree cavities and refugia ­under debris during prolonged dry periods. It does not tolerate freezing temperatures. Mexican Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination (Hailman and Jaeger, 1974). Mexican Treefrogs likely have true color vision. Eggs of Smilisca baudinii. Photo: Cecil Schwalbe CALLING ACTIVITY AND MATE SE­L ECTION

The advertisement call is described as a series of short explosive notes of “wonk-­wonk-­wonk.” Two to 15 notes comprise a call group, with call groups spaced from 15 sec to several minutes apart. The duration of the notes is 0.09–0.13 sec with a fundamental frequency of 135–190 cps. The lowest major frequency is 175–495 cps, whereas the highest major frequency is 2,400–2,725 cps (Duellman, 2001). Males usually call from the ground near ­water but also occasionally from bushes and trees. Males call in duets, with each chorus made up of several pairs of calling males. Successive choruses are initiated by the same duet. Males have an encounter call that is directed ­toward other males that come within 30–50 cm of the primary calling male (in Wells, 1977a). Males ­will engage encroaching intruders aggressively. BREEDING SITES

Breeding occurs in overflow pools, wet grassy meadows, resacas (former channels of the Rio Grande), or streams. Duellman (2001) noted that it breeds in nearly any body of ­water. REPRODUCTION

Most breeding rec­ords in Texas are from June to July, but the breeding season may be longer than currently understood. In México, it extends from June to October. Eggs are oviposited in a surface film, with each deposition having several hundred eggs. Total reproductive output was reported as 2,620–3,320, but the sample size was very small (Duellman, 2001). Hatchlings are 5.1–5.4 mm TL.

Tadpole of Smilisca baudinii. Photo: Seth Patterson

tadpoles grow to 37 mm TL (Rorabaugh and Lemos Espinal, 2016). Newly metamorphosed froglets are 12–15.5 mm SUL. DIET

Tadpoles feed on suspended inorganic and organic particles. Postmetamorphs prob­ably eat a wide variety of invertebrates and occasional small vertebrates. PREDATION AND DEFENSE

Both males and females have a high-­pitched distress call. Predators have not been reported, but likely include snakes, birds, and mammals. POPULATION BIOLOGY

LARVAL ECOL­O GY

Nothing is known of the larval ecol­ogy in Texas. Metamorphosis occurs 14–20 days ­after fertilization, and

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Throughout most of its range, this is an extremely abundant species with sometimes thousands of frogs calling at a single site (Duellman, 2001). Nothing has been reported on its

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Smilisca fodiens 395

Adult Smilisca baudinii, bronze phase. Photo: Cecil Schwalbe

Adult Smilisca baudinii, green phase. Photo: Cecil Schwalbe

demography or life history in Texas. Elsewhere within its range, sexual maturity is reached at 47 mm SUL in males and 56 mm SUL in females.

Adult Smilisca baudinii, patterned. Photo: Seth Patterson

Breeding habitat of Smilisca baudinii in south Texas. Photo: Mike Forstner SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

DISEASES, PARASITES, AND MALFORMATIONS

No information is available.

Smilisca fodiens (Boulenger, 1882) Lowland Burrowing Treefrog ETYMOLOGY

fodiens: from the Latin fodio meaning ‘to dig’ or ‘dig up,’ in reference to the spade-­like inner metatarsal tubercles, which are an adaptation for digging.

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This species is considered Threatened by Texas. T ­ here is virtually nothing known of the species’ biology in Texas.

NOMENCLATURE

Stebbins (2003): Pternohyla fodiens Synonyms: Hyla fodiens, Pternohyla fodiens Most of what is known of the biology of Smilisca fodiens is based on observations from much farther south in México. Duellman (2001) and Sredl (2005b) review the general biology of this species from throughout its range.

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396  Hylidae

IDENTIFICATION

Adults. Smilisca fodiens is a large, yellow-­brown frog with reddish-­brown blotches surrounded by cream or yellow coloration. Snouts are conspicuously rounded, and ­there is a distinctive skin fold along the back of the head. The skin on the top of the head is noticeably hard and firmly attached to the skull (termed cranial co-­ossification), identifying it as a member of the tropical casque-­headed treefrogs. Juveniles are pale green with scattered brown flecks or spots dorsally and a dark stripe between the eyes and the nostrils; dorsal patterns are less prominent than they are in adults. Venters are creamy white. Males have a dark patch on ­either side of the throat that is absent on females. In Arizona, adults averaged 55.1 mm SUL (range 45–70 mm) (Goldberg et al., 1999b). The maximum size of males in México is 62.6 mm SUL, whereas females reach 63.7 mm SUL (Duellman, 2001). Larvae. Tadpoles are dull tan with olive-­brown mottling and dusty white venters. The body of the tadpole is as wide as it is deep and has a bluntly rounded snout. The eyes are small and laterally placed. The caudal musculature is slender, and the dorsal tail fin does not extend to the body. Both are lightly pigmented. Larvae reach a maximum size of 45– 50 mm TL. Descriptions and illustrations of the larvae and mouthparts are in Duellman (2001). Eggs. The eggs have not been described. DISTRIBUTION

The Lowland Burrowing Frog occurs from Pinal County, Arizona, southward into Jalisco and Michoacán, México. Murphy (2019) notes that it is found to 1,500 m in eleva-

tion. Impor­tant distributional references include: Chrapliwy and Williams (1957), ­Sullivan et al. (1996b), Enderson and Bezy (2000), Duellman (2001), Brennan and Holycross (2006), Rorabaugh and Lemos Espinal (2016), Murphy (2019), and Holycross et al. (2021). FOSSIL REC­O RD

Pleistocene fossils of S. fodiens are known from Sonora, México (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

Originally described within the genus Pternohyla (Boulenger, 1882), this species was relegated to the genus Smilisca by Faivovich et al. (2005). Smilisca fodiens is a member of the Tribe Hylinae within the ­Middle American/Holarctic clade of New World hylids. ADULT HABITAT

In Arizona, Smilisca fodiens is associated with mesquite-­lined desert washes. Surrounding vegetation is desert scrub or semidesert grassland. Farther south, this is a species of arid tropical scrub forests. It occurs from sea level in México to at least 2,300 m. TERRESTRIAL ECOL­O GY

Smilisca fodiens is largely fossorial in the winter and dry months, emerging only ­after heavy summer monsoon thunderstorms to breed and forage. In that regard, the head (with the skin attached to the skull) can be used to plug the burrow and thus retard ­water loss and protect the species from terrestrial predators. ­Whether the frog digs its own burrows is unknown but seems likely based on hind foot morphology. The frog can also shed its skin in layers and form a transparent cocoon in order to create a protective barrier, insulating it from ­water loss (Ruibal and Hillman, 1981). Brennan and Holycross (2006) stated that ­these frogs are ­adept at climbing mesquite trees but are mostly terrestrial. ­After metamorphosis, juveniles disperse by day and by night. Lowland Burrowing Treefrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Lowland Burrowing Treefrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Distribution of Smilisca fodiens

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Smilisca fodiens is an explosive breeder, with males calling in large numbers following summer thunderstorms. Males call from in or immediately adjacent to pools, from mud flats, in sparse vegetation, or even from ­under rocks or grass clumps

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Smilisca fodiens 397

Tadpole of Smilisca fodiens. Photo: Ronn Altig

Calling male Smilisca fodiens. Photo: Brent Sigafus

(Duellman, 2001). The advertisement call is described as a loud, raspy “wonk-­wonk-­wonk” repeated at a high rate. It is uttered at 81–115 notes/min with a duration of 0.21– 0.28 sec and a pulse rate of 118–125 pulses/sec; the dominant frequency is 2,200–2,278 cps (Duellman, 2001). Males also have a territorial call that sounds like the advertisement call of Pseudacris maculata—­that is, a fin­ger ­running over comb teeth (­Sullivan et al., 1996b). Calling occurs within only a short period of time ­after rainfall ends. For example, ­Sullivan et al. (1996b) noted that they never heard calling males more than 36 hrs ­after rain had ceased. Males have an encounter call made in response to other males calling within 60–90 cm; this call also is made in response to recordings of advertisement calls from other S. fodiens males (in Wells, 1977a). ­These results suggest that males establish a calling territory that they ­will defend against intruders.

Adult Smilisca fodiens. Photo: Rob Schell

to float within the ­water column tail down. When disturbed, they swim away or float to the bottom of the pool. In Sonora, the larval period may only be 30 days, with larvae growing to 45–50 mm TL (Rorabaugh and Lemos Espinal, 2016). Metamorphosis occurs at 18–24 mm SUL. DIET

Breeding occurs in rain-­formed temporary pools in desert washes, along roads, and in ­cattle tanks (­Sullivan et al., 1996b; Sredl, 2005b). Vegetation in the vicinity of breeding pools may be dense.

Specific data are not available for specimens from Arizona, but the frog likely eats a wide variety of invertebrates. In México, the diet consists of beetles, grasshoppers and their relatives, and other invertebrates (Rorabaugh and Lemos Espinal, 2016).

REPRODUCTION

PREDATION AND DEFENSE

Breeding occurs opportunistically from June to September depending upon rainfall. Eggs are oviposited singly or in small clusters in jelly envelopes scattered on the pool bottom. Eggs hatch rapidly, within a day or so (Rorabaugh and Lemos Espinal, 2016). Clutch size and other aspects of reproduction are unknown.

The hard head is used to seal burrows and thus protects the species from both predators and desiccation. The frog also has an unken reflex whereby it flexes its head downward and elevates its limbs causing it to rest directly on the belly. Nothing is known of its predators.

BREEDING SITES

POPULATION BIOLOGY LARVAL ECOL­O GY

Larvae are ca. 10 mm TL upon hatching. Larvae may feed at the surface of the ­water, but the diet is unknown. They tend

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Juveniles lack the cranial co-­ossification, which suggests they have a dif­fer­ent terrestrial life history from adults. Nothing is known about the demography of adults.

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398  Hylidae

DISEASES, PARASITES, AND MALFORMATIONS

Smilisca fodiens is parasitized by the cestode Distoichometra bufonis and the nematodes Aplectana itzocanensis, Cosmocercella haberi, Rhabdias americanus, Physaloptera sp., and Skrjabinoptera sp. (Goldberg et al., 1999b). Bd has been detected on this species along the Mexican-­US border (Sigafus et al., 2014). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

No information is available. Breeding habitat of Smilisca fodiens. Photo: Cecil Schwalbe

F ­ amily Leptodactylidae

Leptodactylus fragilis (Brocchi, 1877) Mexican White-­lipped Frog ETYMOLOGY

fragilis: from the Latin frag meaning ‘brittle.’ The meaning of the name in relation to the frog is unknown. NOMENCLATURE

Fouquette and Dubois (2014): Leptodactylus (Leptodactylus) fragilis Synonyms: Cystignathus caliginosus, Cystignathus fragilis, Cystignathus labialis, Leptodactylus albilabris, Leptodactylus albilebris, Leptodactylus caliginosus, Leptodactylus gracilis, Leptodactylus labialis [in part], Leptodactylus melanotus, Leptodactylus mystaceus ­There is some discussion in the lit­er­a­ture as to which specific epithet, fragilis or labialis, has priority for this species. I follow the nomenclature recommended by Heyer (2002), Heyer et al. (2006), and Crother (2017) in using fragilis. Heyer (2002) and Heyer et al. (2006) discussed the nomenclatural history in detail and provided a synonymy.

snout. Tails are long, ending in an obtuse point. The tail musculature is cream and heavi­ly pigmented with brown. Venters are pale brown. The tail fins are translucent, do not extend onto the body, and are heavi­ly marked with brown, especially on the dorsal fin. Mulaik (1937) described and figured the larval mouthparts. Heyer et al. (2006) described the larva in detail. Eggs. Eggs are light yellow and lack gelatinous envelopes. The vitellus is about 1.5 mm in dia­meter (Mulaik, 1937). DISTRIBUTION

This species occurs from extreme southern Texas in the Rio Grande Valley (Cameron, Hidalgo, Starr, and Zapata counties) to central Colombia and northern Venezuela. Impor­tant distributional references include Dixon (2000, 2013), Heyer et al. (2006), and Adams (2015). FOSSIL REC­O RD

Fossil Mexican White-­lipped Frogs are reported from Pleistocene deposits in Tamaulipas, México (Holman, 2003; Heyer et al., 2006).

IDENTIFICATION

Adults. This is a very streamlined frog with a pointed snout. The ground coloration is olive yellow, olive, brown, or reddish brown, with irregular, dark dorsal spots. ­These spots may or may not be circled in white. ­There is a distinctive white or cream stripe on the upper jaw and a dark stripe from the nostril to the tympanum. Dorsolateral and lateral folds are pre­sent. ­There is no webbing on the long fin­gers and toes, and ­there are no terminal disks on the digits. ­There is a distinct disk on the belly. Sexual dimorphism is not pre­sent. Males are 27–43 mm SUL and females are 30– 44 mm SUL (Heyer et al., 2006). Larvae. At hatching, tadpoles are nearly invisible except for their yolk sac. ­After several hours, they begin to attain a light brown coloration. Mature tadpoles are dark brown with an elongate snout. Nostrils are nearer the eyes than the

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Distribution of Leptodactylus fragilis

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400  Leptodactylidae

SYSTEMATICS AND GEOGRAPHIC VARIATION

Leptodactylus fragilis is a member of the Fuscus group of the genus Leptodactylus, a diverse assemblage of >23 species that inhabit the Neotropics (Heyer, 1978).

where males ­were calling from thick bunches of grass. Male frogs of the genus Leptodactylus typically excavate shallow cavities located near ponds or pools for egg deposition. REPRODUCTION

ADULT HABITAT

The habitat of this species in Texas was described as “moist meadows, irrigated cane fields, drains and gutters in towns, beneath stones, logs, in sandy banks and fields; near streams and marshy places” (Wright and Wright, 1938). TERRESTRIAL ECOLOGY

Nothing is known concerning the life history of L. fragilis at the northern end of its range. In Costa Rica, the species actively hunts insects at night, and hides ­under logs and rocks or in piles of debris by day. In terms of vision, Mexican White-­lipped Frogs are unclassifiable with regard to spectral response, but the response is similar to the “blue-­ mode response” but shifted to green (Hailman and Jaeger, 1974). Mexican White-­lipped Frogs may have color vision.

Eggs are deposited terrestrially in a frothy mass in small excavations at the base of vegetation near ­water (Livezey and Wright, 1947). Males construct ­these shallow depressions into which females deposit their eggs. The female whips up a foam mass from her body secretions; the foam ­will protect the tadpoles even during dry weather. However, the normal sequence is to have the depression flooded, which releases the tadpoles into ­water. The clutch size is 80 eggs or greater, with Mulaik (1937) reporting 1 clutch of 86 eggs. Hatching occurs within 40 hrs of deposition, when larvae are 6.6 mm TL. ­After 24 hrs, they are ca. 8.1 mm TL. LARVAL ECOLOGY

Larvae are free-­living benthic feeders that live in nonflowing ­water. The larval period is ca. 30–35 days (Mulaik, 1937). Transforming juveniles average 16.1 mm SUL.

CALLING ACTIVITY AND MATE SE­L ECTION

Calling occurs from concealed positions at the base of grass hummocks. Males construct shallow cavities that ­will be used for egg deposition and call from ­these cavities. Mulaik (1937) described the call as resembling the “plunk-­plunk” from a drop of ­water falling into a cave pool, although Heyer et al. (2006) described it as “a rising harsh whistle.” Calls are given at the rate of 120–150 calls/min with a call duration of 0.16–0.2 sec. The call begins at about 600– 750 Hz and has a fast rise in intensity during the first third of the call, a weak increase in the next third, and another weak increase followed by a sharp drop in intensity at the end. The call rises to 1,000–1,200 Hz. The dominant fundamental frequency is 740–780 Hz in Texas (Fouquette, 1960; Heyer et al., 2006). Males call in the late after­noon and night during the rainy season. Adams (2015) found them calling ­after heavy rains in mid-­May and ­after a tropical depression in September. Males ­will also call away from the nest depression. The female responds to the male’s call and moves ­toward him. As she moves next to him, he leads her to the shallow depression. During this movement, the female makes what appears to be a reciprocal call, consisting of a rapid series of short notes or a trill that is barely audible (Bernal and Ron, 2004).

DIET

Nothing is reported on the diet of this species in Texas. Presumably it eats a wide variety of small invertebrates. PREDATION AND DEFENSE

Leptodactylus fragilis is a superb jumper, giving rise to the common name rocket frog. Nothing is known concerning its predators, but they likely include snakes, birds, and mammals. POPULATION BIOLOGY

No information is available.

BREEDING SITES

Mulaik (1937) found a clutch in a shallow depression 4 cm in dia­meter and 3 cm deep at the base of a grass hummock 30 cm from the nearest ­water. Adams (2015) found them breeding in a flooded roadside ditch in Starr County, Texas,

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Adult Leptodactylus fragilis. Starr County, Texas. Photo: Clint Guadiana

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Leptodactylus fragilis 401

DISEASES, PARASITES, AND MALFORMATIONS

No information is available from the United States. The following nematodes have been reported from Costa Rica: Cosmocerca podicipinus, Schrankiana formosula (Goldberg et al., 2013). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

For a long time, t­ here was a question as to ­whether this species still occurred in Texas. However, it was recorded from Starr County in 2002 and 2005, and was said to be not

Dodd_Canada_int_5pgs_B3.indd 401

uncommon (Linam, 2009); additional observations—­with photo­graphs—­were available from 2014 to 2015 in Starr and Zapata counties (Adams, 2015). In 18 yrs of field research, Fred Zaidan (University of Texas Rio Grande Valley, personal communication), has never seen this frog in the region. Apparently, it is difficult to detect in the field, although recent sightings suggest observers should concentrate survey efforts ­after very heavy rainfall events. Dixon (2000, 2013) stated that the use of organophosphates could have caused its extirpation. Although this is not the case, populations could have been affected adversely. This species is very common from México to South Amer­i­ca, but it is considered Threatened in Texas.

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F ­ amily Microhylidae

Gastrophryne carolinensis (Holbrook, 1835) Eastern Narrow-­mouthed Frog ETYMOLOGY

carolinensis: Latinized noun referring to Carolina. The species was described from individuals found near Charleston, South Carolina. NOMENCLATURE

Synonyms: Engystoma carolinense, Engystoma rugosum, Gastrophryne rugosum, Microhyla areolata, Microhyla carolinensis, Stenocephalus carolinensis Although commonly called a toad, this species is in the ­family Microhylidae, not the ­family Bufonidae, the true toads. Hecht and Matalas (1946) and Nelson (1972) reviewed the nomenclatural history of Gastrophryne. IDENTIFICATION

Adults. The Eastern Narrow-­mouthed Frog is a small, rotund, smooth-­skinned, pointy-­snouted frog. The dorsum is brownish red to blue black and may be highly mottled with coppery or silvery pigmentation. A reddish (or chestnut) color may give the appearance of dorsolateral bands; the reddish pigmentation also may be pre­sent on the front legs. Meshaka and Layne (2015) suggested ­there are 3 color morphs: carolinensis -­dark dorsum blotched or with indistinct dorsolateral stripes, venter mottled; “Key West” -­ dorsal pattern of 2 distinct, light tan dorsolateral stripes bordered by distinct dark margin on tan background; olivacea-­like -­virtually without pattern with reduced ventral coloring. ­These patterns may vary in frequency geo­graph­i­ cally, that is, although considerable pattern variation exists within populations, one of the patterns may be dominant within a par­tic­u­lar geographic region. The head is flattened and small. ­There is a transverse dermal groove ­behind the eyes that may be covered by a fold of skin. This fold becomes more pronounced when the frog is irritated. The belly is medium to dark gray; brown

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splotches or light mottling may be pre­sent. ­There is no tympanum nor is ­there webbing between the toes. The rear feet have small spades that are used for digging into friable soils. Sexual dimorphism is apparent. Males have a dark throat patch that never ­really dis­appears, although it may intensify in coloration during the breeding season. Most males also develop conical chin tubercles (1–28) along the ventral surface of the lower jaw (Mittleman, 1950), at least in some populations, and the area of the subgular throat patch appears wrinkled. ­These latter 2 characteristics are evident only during the breeding season. A few males may not have a dark throat or chin tubercles, however, and some breeding females may have tubercles in the perianal region (Nelson, 1972). Adults are 22–39 mm SUL. Albino adults from Louisiana ­were reported by Gordon (1955). Males are generally smaller than females in SUL and body mass. ­There are lit­er­a­ture rec­ords for males and females from Texas (means 27.4 mm SUL, 29.5 mm SUL), Florida (means 23.4 mm SUL, 25.9 mm SUL) (Blair, 1955a), and Georgia (20–65 mm SUL, mean 31.9 mm) (O’Hare and Madden, 2018). Loftus-­Hills and Littlejohn (1992) recorded males with a mean SUL of 28.6 mm (range 29.9–32 mm) also from Texas, and Meshaka and Woolfenden (1999) recorded a mean of 25.7 mm SUL for males (range 19.8– 29.3 mm) and 26.4 mm SUL for females (range 20–33 mm) from south Florida. Also in south Florida, Duellman and Schwartz (1958) recorded a mean of 26.2 mm SUL for males (range 18.8–30.5 mm) and 28.3 mm SUL (range 22.4– 32.5 mm) for females. In north central Florida, males ranged from 22 to 33 mm SUL and females from 21 to 35 mm SUL (Dodd, 1995). In Missouri, the largest males normally are 32.2 mm SUL, whereas the largest females normally are 34.3 mm SUL (Anderson, 1954). However, ­there is 1 rec­ord of an individual 39 mm SUL (in Johnson, 2000). In Arkansas, the mean size for males was 27.6 mm SUL and for females 29.6 mm SUL; a few individuals of both sexes reached 36.5 mm SUL (Trauth et al., 1999). In Alabama, adults ­were 22.9–33.6 mm SUL (Brown, 1956), whereas ­Virginia males averaged 28.3 mm SUL (range 25.5–32.2 mm)

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and females 30.1 mm SUL (range 24.4–34.2 mm) (Meshaka et al., 2015a). ­There is considerable overlap among age classes in terms of SUL. Larvae. Tadpoles are small and jet black, with distinct, white to pink lateral stripes on ­either side of the posterior portion of the body extending onto the tail musculature. Bellies are deep and light in coloration. Viewed from the side, the head comes to a fairly sharp point. The jaws do not have keratinized teeth, and no oral disk is pre­sent. Tails are 1–1.5 times the length of the body and have a pointed tip. The maximum TL is 38 mm, although normally the TL is 15–30 mm. Albino tadpoles ­were reported from Louisiana (Anderson, 1951). Descriptions of the tadpole are in Wright (1929, 1932), Orton (1946), and Siekmann (1949). Eggs. Individual eggs are small and dark brown or black dorsally and white, yellowish, or brownish ventrally. The mean dia­meter of the envelope is 3.35 mm (range 2.8– 4.0 mm); the vitellus is 0.7–1.3 mm (Wright, 1932; Livezey and Wright, 1947; Trauth et al., 1999; Meshaka et al., 2015a). The eggs are deposited in a small surface film with a mosaic-­like structure; the egg mass is round or squarish, with 10–150 eggs deposited in each mass. The egg jelly is truncate (i.e., flattened or appearing to be cut off). If disturbed, the surface film breaks up into individual eggs. Total annual clutch size varies considerably (see Reproduction), and females likely deposit partial clutches rather than a full complement with each oviposition. DISTRIBUTION

The Eastern Narrow-­mouthed Frog occurs from the Delmarva Peninsula (White and White, 2007) and southeastern

­ irginia throughout much of the Piedmont and Southeastern V and Gulf Coastal Plains to the southern tip of Florida and the Florida Keys. In the mid-­South, populations are found in Tennessee, Kentucky, and southwestern ­Virginia along tributaries of the Tennessee River. The species barely enters Illinois. The range extends across the Mississippi River through southern Missouri and into eastern Oklahoma and Texas. A population also occurs in extreme southeastern Kansas. Eastern Narrow-­mouthed Frogs have been introduced into the Bahamas (­Grand Bahama, New Providence) and to ­Grand Cayman Island (Lever, 2003). The species naturally occurs on islands, including islands at the tip of the Delmarva Peninsula (Mitchell, 2012), Cape Hatteras, Shackleford Banks, Smith, Roanoke, and Bodie islands in North Carolina (Lewis, 1946; Engels, 1952; ­Brothers, 1965; Gibbons and Coker, 1978; Gaul and Mitchell, 2007; Parlin et al., 2019), Kiawah Island in South Carolina (Gibbons and Coker, 1978; Hanson and McElroy, 2015), Isle of Hope, Blackbeard, Ossabaw, St. Catherines, Tybee, Sapelo, ­Little Cumberland, and Cumberland islands in Georgia (Martof, 1963; Laerm et al., 2000; Shoop and Ruckdeschel, 2006; O’Hare and Madden, 2018), Big Pine Key, Cudjoe Key, Egmont Key, Key Largo, Key West, Lignumvitae Key, ­Little Torch, Matecumbe Key, Stock Island, Sugarloaf Key, Summerland Key, St. George Island, and St. Vincent Island in Florida (Duellman and Schwartz, 1958; Lazell, 1989; Franz et al., 1992; Irwin et al., 2001), Dauphin Island, Alabama (Jackson and Jackson, 1970), and Padre Island, Texas (https://­www​.nps​ .­gov​/­pais​/­learn​/­nature​/­amphibians​.­htm). Impor­tant distributional references include: range-­wide (Nelson, 1972), Alabama (Brown, 1956; Mount, 1975), Arkansas (Black and Dellinger, 1938; Trauth et al., 2004), Florida (Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Illinois (Smith, 1961; Phillips et al., 1999), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Kansas (Collins, 1993; Collins et al., 2010), Kentucky (Barbour, 1971), Mary­land (Cooper, 1953; Harris, 1975; Cunningham and Nazdrowicz, 2018), Missouri (Johnson, 2000; Daniel and Edmond, 2006), North Carolina (Meyers and Pike, 2006; Dorcas et al., 2007), Oklahoma (Sievert and Sievert, 2006), South Carolina (Dodd and Barichivich, 2017; Fields, 2019), Tennessee (Redmond and Scott, 1996; Niemiller and Reynolds, 2011), Texas (Hardy, 1995; Dixon, 2000, 2013; Tipton et al., 2012), and ­Virginia (Fowler and Hoffman, 1951; Tobey, 1985; Mitchell and Reay, 1999). FOSSIL REC­O RD

Distribution of Gastrophryne carolinensis

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Miocene fossils referred to this species are known from Hemingfordian deposits in Florida (Auffenberg, 1956).

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Pliocene fossils are reported from Florida (Meylan, 2005), and Pleistocene Rancholabrean fossils are known from Florida and Georgia (Holman, 2003). The ilium of this species is identifiable by its well-­developed, triangular dorsal prominence. SYSTEMATICS AND GEOGRAPHIC VARIATION

Frogs of the genus Gastrophryne are derived phyloge­ne­ tically from the South American clade of microhylid frogs (Meijden et al., 2007). ­There are 4 species within this genus, and 3 enter the United States. Gastrophryne is most closely related to the genus Hypopachus (Nelson, 1972). Natu­ral hybridization with G. olivacea has been reported in a narrow contact area in eastern Texas and Oklahoma (Hecht and Matalas, 1946), although the larger G. carolinensis males are reluctant to clasp the much smaller G. olivacea females. Strecker’s (1909) description of Microhyla areolata appears to represent ­these hybrid individuals. Pattern phenotypes intermediate between Gastrophryne olivacea and G. carolinensis are known from a number of locations even where the species are not sympatric. Therefore, pattern variation is not based on hybridization alone (contrary to Hecht and Matalas, 1946), and care must be taken when identifying “hybrids” solely on morphological grounds (Nelson, 1972). Most pattern variation relates to the extent of mottling on the belly, sides, and throat. Laboratory crosses with G. olivacea may result in tadpoles (Blair, 1950). The diploid chromosome number is 22. ­There appear to be differences in the size of adult G. ­carolinensis in varying parts of its range. Such variation becomes impor­tant when examining demographic traits such as size and age at maturity. For example, Blair (1955a) provided mea­sure­ment data for males from 9 locations in Florida, Oklahoma, and Texas. He noted that ­there was a geographic trend in size, with smaller individuals in the East and larger individuals in the West. In addition, ­there appears to be variation in size by habitats. Nelson (1972) showed that males and females from lowland habitats ­were smaller than males from upland habitats. This size variation was consistent throughout the range of the species. Variation in size is noted in age at sexual maturity among populations (see Population Biology). The basis for the variation could be ge­ne­tic or could result from differences in environmental conditions. In addition to size, morphological variation among populations is evident in the presence or absence of tubercles in males and in dorsal pattern. In some populations of G. carolinensis, individuals lack chin tubercles (e.g., in Illinois; Smith, 1961), whereas in other populations, a small percentage of individuals have them (e.g., 10% in Alabama; Mount, 1975). Florida Keys populations of G. carolinensis also have dorsal patterns that differ from mainland popula-

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tions. Most Keys frogs have 2 prominent, light tan dorsolateral stripes bordered by a distinct dark margin on a tan background. ­Others have a G. olivacea-­like color pattern. Frogs with a typical G. carolinensis-­like pattern are found in the Keys, but they make up less than 25% of the population (Duellman and Schwartz, 1958). Similar pattern variation is evident in other G. carolinensis populations, but not with the same high frequencies of aty­pi­cal patterns as in the Keys (also see Meshaka et al., 2015a). ADULT HABITAT

Gastrophryne carolinensis inhabits a wide range of environments, from flooded swamps, steephead ravines, and open pastures and fields to mesophytic hammocks, and upland dry pine, mixed deciduous, or hardwood forests (Carr, 1940a; Anderson et al., 1952; Harima, 1969; Bennett et al., 1980; Enge et al., 1996; Enge, 1998a, 1998b; Enge and Wood, 1998; Baxley and Qualls, 2009). Dif­fer­ent habitat types are occupied in dif­fer­ent parts of the range, depending on availability. In Missouri, for example, Anderson (1954) identified 4 major habitats: cypress-­gum swamps, bottomland hardwoods, live oak ridges, and pine-­oak woodlands. In uplands, this species is frequently associated with sandy, friable soils. Gastrophryne carolinensis inhabits coastal marshes, caves and sinkhole depressions, springs, and river bluffs, all of which serve as refugia from extreme weather conditions. They may be found along small streams (Metts et al., 2001), and ­there are reports of this species in close proximity to beaches (Viosca, 1923) or brackish habitats (Noble and Hassler, 1936; Werler and McCallion, 1951; Hardy, 1953; Neill, 1958). In the West, G. carolinensis tends to occupy lowland riverine forests, extending its distribution into the prairie region. The Eastern Narrow-­mouthed Frog requires constant moisture, as frogs ­will quickly desiccate when exposed to very dry conditions or when left in traps without a moisture source. They generally ­favor habitats with dense leaf litter and surface debris covering organic soils. If retreats with high humidity (animal burrows, crevices, root channels) are available, however, they ­will occupy dry, sandy areas. They often are found ­under surface debris, rocks, boards, logs, decaying vegetation (such as mats of ­water hyacinth), and in decaying stumps and rotten wood that remains moist or saturated. I have found them in palm stumps in mixed palm-­hardwood hammocks on dry offshore islands in west central Florida. They even occur in Florida muskrat (Neofiber alleni) ­houses in flooded wet prairies and in Florida woodrat (Neotoma floridana) middens (Lee, 1968; Smith and Franz, 1994). TERRESTRIAL AND AQUATIC ECOLOGY

­ hese are mostly semifossorial or fossorial frogs, although T Anderson (1954) found an individual 2.4 m off the ground

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in a dead stump, and arboreal be­hav­ior has been reported on bushes and trees in Louisiana (to 2.5 m; Godwin et al., 2018). The Eastern Narrow-­mouthed Frog is capable of making short burrows ­under the surface of the ground, or they may occupy the burrows of other animals (e.g., crayfish, Gopher Tortoises, mammals) or stump holes (Murphy et al., 2021). Individuals ­will sit with only the top portion of the head vis­i­ble at the surface, waiting for prey, normally ants, to pass by. Gastrophryne occurs in just about ­every terrestrial situation available during the nonbreeding season, from short-­grass fields to dense meadows, and from cypress-­gum swamps to upland forests and stands of palmetto flatwoods (e.g., Dodd, 1992; Chandler et al., 2015a; Erwin et al., 2016). They occur in anthropogenic habitats such as golf courses, lawns, gardens, urban lots, suburban developments, and even in agricultural fields. By day, the frogs remain concealed in retreats. Most activity occurs at dusk or during the early eve­ning, and surface activity is especially common during and immediately ­after precipitation. ­After about 22:00 hrs, activity is reduced, and it ceases by about 24:00 hrs. Activity occurs year-­round in Florida (Dodd, 1995), with usually only brief periods of inactivity due to cold weather. Seasonal activity occurs from May through September or October in more northern locations (Gentry, 1955; Dodd, 1995; Gibson and Sattler, 2020). Eastern Narrow-­mouthed Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Eastern Narrow-­mouthed Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males call throughout the breeding season when temperatures exceed 19.5°C; ­water and air temperatures can approach 36°C and 33°C, respectively, when calling (Dundee and Rossman, 1989). Calling dates vary among sites, with frogs initiating and ending calling at dif­fer­ent dates even among nearby sites (McKnight and Ligon, 2016). In south Florida, calling may extend from March to October, whereas in North Carolina and South Carolina it occurs from June to August (Gaul and Mitchell, 2007; Todd et al., 2011a). Rec­ords are available for ­Virginia from May to August (Mitchell, 1986; Meshaka et al., 2015a; Gibson and Sattler, 2020), eastern Tennessee from June to September (Dodd, 2004), Arkansas from April to August (Trauth et al., 2004), Oklahoma from May to July (Bragg, 1950a), and Texas from April to September, where the timing of the first calling date has shifted to ­earlier in the year (Car­ter et al., 2018).

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Both males and females move back and forth between retreat sites and breeding sites on a daily basis. Immigration and emigration patterns at breeding ponds may change annually and by sex and life stage (Dodd and Cade, 1998). Movements are nonrandom, depending on the type and availability of surrounding habitats. Gastrophryne carolinensis does not appear to use movement corridors between nearby terrestrial refugia and aquatic breeding sites, although juveniles use temporary wetlands as stopping points as they disperse from other wetlands. Terrestrial habitats may be occupied at considerable distances from the nearest wetland. In north central Florida, for example, Dodd (1996) captured G. carolinensis 42–914 m (mean 420 m) from the nearest pos­si­ble breeding site while trapping for snakes in dry sandhills habitats. Most terrestrial movements occur at dusk. Males arrive at breeding ponds about 10–20 min prior to females and immediately begin calling from very shallow positions in the ­water. Calling occurs in bouts, with periods of silence in between bouts. Choruses usually consist of 2 or more males calling in tandem, although not all frogs call in each bout, and dif­fer­ent combinations and numbers of frogs may call in successive choruses. Certain individuals may call far more often than ­others. Single males may call outside the bouts, but ­these individuals call less often than ­those in larger choruses. Anderson (1954) provided examples of male chorus calling patterns. Males use vegetation, branches, detritus, bark, logs, or nearly any other object in the ­water for cover. The vegetation serves to conceal the male as well as to offer him support and balance while calling. When calling, males raise their body in a vertical or near-­vertical position and use their forelimbs for stability as they call. The substrate at the calling site may consist of mud, clay, sand, or vegetative debris. Indeed Carr (1940a) noted a unique calling be­hav­ior in this species whereby males buried their bodies in soft sand in very shallow ­water and called with only their snouts protruding. The call has been compared to the bleating of a sheep, but some authors have likened it to a buzz with up to 40 consecutive harmonics; it is difficult to describe verbally. The mean call duration ranges from 0.04 to 2.4 sec at a mean frequency of 2.4–3.9 kHz, depending on temperature (Blair, 1955b; Nelson, 1972, 1973), and Anderson (1954) noted a duration of about 1.4 sec at 25°C in Missouri. Frequency tends to increase with temperature. The harmonic interval is 160–250 Hz. Nelson (1972) provided data on call characteristics from throughout the species’ range. When a full chorus sings, the sound can be quite intense and has been described as a “din.” During the eve­ning, males tend to remain in the same place and do not change their calling station even over many hours. ­After mating, they may leave their station. Calling

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males tolerate one another, even in close proximity, and calling site territoriality is not evident. Anderson (1954) recorded nearest males calling from within 25 mm of one another to 6.1 m away from one another. Females approach males and swim with their bodies floating and their rear legs widely extended. At the approach of the female, the male increases his calling rate. The cues for triggering amplexus are unknown, but males often remain in position calling even as females approach in very close proximity. Soft grunts and croaks are often heard just prior to amplexus. A male ­will approach a female from ­behind and swim into an amplexing position (­either semi pectoral or axillary), inching his way slowly up the female’s back. Males are able to remain clasped to females by using their arms and by the adherence of special dermal secretions; essentially, they become glued to one another. Glands on the male’s venter in the region of the sternum secrete an adhesive substance (Conaway and Metter, 1967), and the “breeding secretion” allows rotund males to remain in amplexus despite their short ­little arms. Amplexus may be necessary to trigger ovulation, and oviposition occurs 1.5–2 hrs ­after the initiation of amplexus. Males may try to amplex other males or unreceptive females, resulting in inguinal amplexus. In successful amplexus with a receptive female, the male firmly holds the female with his thumbs, and his palms are turned out and upward. The male’s back ­will ­ride just above ­water, the thighs are flexed, and the feet are held laterally and anteriorly. The female’s body is held at an a­ ngle of 45° from the ­water’s surface, her thighs are extended, and her feet also are held laterally and slightly anteriorly. An amplexed male and female ­will initially swim together, but the male soon stops and holds his legs tightly anterior. The female ­will swim in short, jerky motions with the male firmly clasping her. Shortly before oviposition, the female ­will stop moving and raise her hind limbs and the rear part of her body ­toward the ­water’s surface. This results in a backward flexed posture with the rear limbs at right ­angles and even with the ­water’s surface. She moves backward a ­little, then extends her legs rigidly in a ventrolateral position. At this time, the vent ­will be above the ­water surface. The male then slides forward, bringing his vent into close proximity to the female’s vent. The eggs are fertilized as they burst from the female’s vent in groups of about 30 eggs per ejaculation. ­After oviposition is completed, the male may remain clasped for a short time period before releasing the female. Anderson (1954) described mating be­hav­ior in detail.

depression wetlands, sinkhole ponds, cypress savannas, seasonally inundated wetlands in pastures, small to large permanent and temporary ponds, large semipermanent lakes, cypress/gum ponds, beaver ponds, and a variety of anthropogenic wetlands (Cash, 1994; Dodd, 1995; Babbitt and Tanner, 2000; Metts et al., 2001; Surdick, 2005; Babbitt et al., 2006; Lichtenberg et al., 2006; Liner et al., 2008). They are frequently found in borrow pits and roadside ditches, and they can even breed in wheel ruts. Wetlands may contain deep ­water but breeding only takes place in relatively shallow w ­ ater (21 mm) has some remaining spermatozoa year-­round. Anderson (1954) described 3 stages of oocyte development, the last of which results in pigmented oocytes. Oocytes grow ­little in the fall and early winter, and pigmented oocytes first appear in January. Oocyte growth occurs steadily ­under normal conditions, but it can be delayed due to cold weather. Growth ­will then accelerate to make up for the delay. A single female may contain oocytes destined for 2 or 3 breeding seasons, and thus eggs may be at very dif­fer­ent sizes and stages of development. Egg counts vary considerably. In Missouri, Anderson (1954) counted 152–1,089 eggs per female (overall mean 510); the means ­were 378 for 2-­yr-­old females (23–24 mm SUL), 466 for 3-­yr-­old females (24.5–26.9 mm SUL), and 681 for 4-­yr-­old females (>27 mm). In Arkansas, counts ranged between 186 and 1,459 (mean 673); neither clutch size nor ovum size increased with female body size (Trauth

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et al., 1999). In ­Virginia, clutch size averaged 825 (range 278–1,618) (Meshaka et al., 2015a) and 820 (range 208– 1,614) (Mitchell and Pague, 2014). Wright and Wright (1949) reported clutches of up to 850 eggs, and Deckert (1914) noted that 100–150 eggs ­were oviposited at a time. The wide variation in reports of clutch size could be due to oviposition of partial clutches or could reflect regional and individual variation. Early development was described by Ryder (1891). Clutch size was not correlated with female SUL in ­Virginia (Meshaka et al., 2015a). LARVAL ECOLOGY

Hatching occurs 1.5–3 days ­after oviposition (Ryder, 1891; Wright, 1932). Tadpoles are filter feeders (on zoo-­and phytoplankton) that do not possess rasping mouthparts. Larvae grow rapidly in order to achieve metamorphosis prior to pond drying; the larval period is 23–67 days (Wright, 1932). Recently transformed metamorphs are 10 mm SUL in Missouri (Anderson, 1954), 11 mm SUL in Florida (Dodd, 1995), and 7–12 mm SUL in Georgia (Wright, 1932). Many tadpoles may metamorphose from a single wetland. Degregorio et al. (2014) recorded 270 metamorphs from a single pond over a 1 yr period in South Carolina. Tadpoles of G. carolinensis are capable of acclimating to vari­ous thermal regimes, even to temperatures of 43–44°C (Cupp, 1980). Newly hatched larvae are capable of tolerating higher temperatures than later-­stage larvae. As they approach metamorphosis, larvae can only tolerate temperatures to 39–40°C. Larvae are sensitive to the presence of predator cues in the ­water. In experimental ­trials, larvae decreased activity levels when chemical cues of dragonfly nymphs ­were pre­sent, but not when directly exposed to the threat of predation (Schiwitz et al., 2020). Presumably, they swim rapidly to escape rather than become unresponsive when directly threatened. DIET AND FEEDING

The Eastern Narrow-­mouthed Frog is an ant specialist. Deyrup et al. (2013) reported 43 species of ants consumed by a Florida population of G. carolinensis, where 77% of the ant

Tadpole of Gastrophryne carolinensis. Photo: Stan Trauth

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species ­were in the genera Pheidole or Nylanderia. In ­Virginia, Ivanov and Gibson (2020a) noted 3 species of ants in a single individual. Individuals ­will often sit next to or even within an ant-­hill (Wood, 1948) and gorge themselves on ants leaving the mound. Other small invertebrates also may be consumed, such as mites, collembolans, isopods, spiders, snails, beetles, and termites (Anderson, 1954; Brown, 1974). In Arkansas, Brown (1974) found a mean of 35–45 prey items per gut in G. carolinensis, depending upon sampling location. PREDATION AND DEFENSE

This species is more of a walker than a hopper, and terrestrial escape attempts are awkward and cover only short distances. However, they can be surprisingly agile in short bursts and can quickly burrow into leaf litter or loose soil to escape. Individuals often ­will remain immobile, crouch, or try to walk away when faced by a predator (Marchisin and Anderson, 1978). When floating in the ­water, a disturbed animal or an amplexed pair ­will dive to escape. Gastrophryne have a noxious skin secretion that makes them less prone to predation than other small frogs (Garton and Mushinsky, 1979). The secretion also protects them from attack by the ants on which they feed. It ­causes a burning or irritating sensation to delicate tissues such as the eyes and mucous membranes of the mouth and throat, and it entangles attacking ants. Anderson (1954) noted that other frogs (Dryophytes cinereus, D. squirellus) placed in close proximity to Eastern Narrow-­mouthed Frogs could die from prolonged exposure. Secretions are produced by granular glands located throughout the skin; ­these glands first appear late in development, coinciding with the eruption of the forelimbs. Anderson (1954) reported that G. carolinensis makes a faint clicking or chirp when roughly handled. ­Whether ­these sounds function in a defensive or alarm context is unknown, although it seems unlikely that such a relatively soft sound would deter a predator. ­There are a few reports of predation by snakes (Thamnophis sp., T. sauritus, Agkistrodon contortrix) and unidentified mammals (Wright, 1932; Anderson, 1942; Luhring and Ross, 2012). Adams et al. (2011) found that larvae are unpalatable to some species, although the effectiveness of larval unpalatability is variable. Aquatic invertebrates are likely predators on larvae, but older larvae are distasteful or noxious to many predators. However, larvae are readily eaten by some native fish and crayfish (Baber and Babbitt, 2003; Adams et al., 2011). Individuals are also attacked by the mosquito Uranotaenia lowii (Blosser and Lounibos, 2012). POPULATION BIOLOGY

Sexual maturity is reached at 21–26.7 mm SUL, depending upon location, in the second spring (first spring following

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Adult Gastrophryne carolinensis. Photo: Todd Pierson

metamorphosis), although some females may not deposit eggs ­until their third spring (second spring following metamorphosis). Secondary sex characteristics in males (black chins, mature spermatozoa, development of gonads) occur by 21–22.9 mm SUL in Missouri (Anderson, 1954) and 21.5 mm SUL in Arkansas (Trauth et al., 1999); pigmented eggs are first observed in females by 23 mm SUL in Missouri and 26.7 mm SUL in Arkansas. Based on recapture data in north central Florida, 2-­yr-­old and 3-­yr-­old G. carolinensis are a minimum of 27 mm SUL; a few G. carolinensis have been found 4 yrs ­after having been marked as adults and ­were >32 mm SUL (Dodd, 1995). Sex ratios appear to be male biased both at breeding ponds and in many terrestrial samples (Anderson et al., 1952; Anderson, 1954; Dodd, 1995; Trauth et al., 1999; but see Meshaka and Woolfenden, 1999). For example, Trauth et al. (1999) found a sex ratio of 1.52 males per female in Arkansas, and Dodd (1995) found 1.3 males per female over a 5 yr period of continuous sampling (1,154 males, 929 females). Perceptions of skewed sex ratios may result from biased sampling, differential movement between the sexes, or differences in the amount of time each sex spends at a breeding site. However, males may indeed outnumber females as a result of differences in survivorship or fertilization rates. Whereas most females frequent breeding ponds ­every season, reproduction is not successful ­every year. Indeed, metamorphs are produced only when sufficient hydroperiod is maintained to complete larval development, and ­there may be many years when few or no metamorphs are produced at any par­tic­u­lar breeding site or when no breeding occurs at all (Brown, 1956; Dodd, 1995; Semlitsch et al., 1996; Daszak et al., 2005). As a result, ­there is likely considerable fluctuation in recruitment and survivorship among G. carolinensis populations, making them vulnerable to drought and climate change. Salice (2012) has shown that

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catastrophic reproductive failure would be a strong driver of extinction risk ­under increasing aridity due to climate change. As the probability of insufficient hydroperiods increases, a population becomes more demographically vulnerable, especially if other stressors are pre­sent. ­There have been no mark-­recapture attempts to document population sizes of G. carolinensis at any breeding site. Populations often appear large, but ­there may be considerable variation annually and among sites (e.g., Gibbons and Bennett, 1974; Greenberg and Tanner, 2005b). Dodd (1992) recorded 4,573 dif­fer­ent G. carolinensis using a very small sandhill depression marsh in north central Florida over a 5 yr period, despite a severe drought that ­limited reproduction to only 1 yr. Greenberg and Tanner (2005b) captured 3,117 G. carolinensis adults but only 168 juveniles during a 7 yr study in central Florida. In South Carolina, Semlitsch et al. (1996) recorded 3,072 adults and 2,930 metamorphs at a temporary pond over a 16 yr period, but most captures occurred in only 3 yrs for adults and 1 yr for metamorphs. ­These studies have shown that individuals can use even small, shallow, temporary ponds for breeding and as refugia in other­wise dry habitats, despite stochastic environmental conditions. Eastern Narrow-­mouthed Frogs can be ubiquitous within a landscape when wetlands are abundant. In south Florida for example, Babbitt et al. (2006) found them at 79% of the sites they sampled. Terrestrial abundance in forests also can be high. For example, Bennett et al. (1980) captured 4,195 G. carolinensis during 2 summers trapping in 3 forest types, and Dodd et al. (2007) found Eastern Narrow-­mouth Frogs to be the dominant amphibian on the forest floor of a variety of mesic, mixed palm-­pine–­deciduous forest associations on the northern Gulf Coast. Juvenile recruitment is negatively correlated with mean ­water depth during development, but positively correlated with total rainfall and the number of breeding adults (Greenberg et al., 2017a). Average air temperature does not influence juvenile recruitment. Populations of this species vary widely in abundance among years and breeding ponds, making ­simple trend analyses difficult at the landscape level (Greenberg et al., 2018a). For example, ­these authors found that recruitment was correlated with the abundance of adult G. carolinensis breeding populations only for 4 yrs following metamorphosis. ­Because ­there is so much annual variation in breeding success among even closely situated ponds, even continuous sampling, much less sampling at longer intervals, would have difficulty detecting population trends across a regional landscape. At Greenberg et al.’s (2018a) study site focusing on 7 wetlands in Ocala National Forest in Florida, low statistical power hampered an ability to detect trends in G. carolinensis populations even for data collected annually over a 24 yr period.

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COMMUNITY ECOLOGY

The ranges of G. carolinensis and G. olivacea overlap in eastern Texas and Oklahoma. Although hybridization may occur in areas of contact, differences in call structure and body size help to minimize gene exchange. In zones of contact with G. olivacea, males and females are both larger than they are in areas where the 2 species are not sympatric (Blair, 1955a). The structure of the advertisement calls is also dif­fer­ent between the species, with G. carolinensis having calls of lower frequencies and shorter duration than the calls of G. olivacea (Blair, 1955b; Awbrey, 1965; Nelson, 1972; Loftus-­Hills and Littlejohn, 1992). Hybrids have call characteristics intermediate between the parental species. In choice tests, females prefer the calls of males of their own species to ­those of G. olivacea (Awbrey, 1965). The tadpoles of G. carolinensis are palatable, but the be­hav­ior of the tadpole aids in predator avoidance. For example, mosquitofish (Gambusia holbrooki) readily forage effectively for tadpoles in structurally complex environments. However, tadpoles of Eastern Narrow-­mouthed Frogs have greater survivorship in structurally complex habitats than in habitats with fewer places to hide (Baber and Babbitt, 2004). This is ­because the larvae are much less active than the tadpoles of other species and thus tend not to draw attention to themselves. With G. carolinensis larvae remaining immobile much of the time, fish tend to focus on more readily detectable species. Gastrophryne carolinensis tadpoles also decrease activity in the presence of predator (dragonfly larvae) chemical cues and conspecific alarm cues (Schiwitz et al., 2020). DISEASES, PARASITES, AND MALFORMATIONS

Larval Eastern Narrow-­mouthed Frogs inoculated orally with Frog Virus 3 or an FV3-­like isolate developed disease symptoms, but exposure to ­water inoculated with the viruses did not result in infection (Hoverman et al., 2010). Infected frogs ­were 3.8 times as likely to die during development as uninfected larvae, and mild to moderate edema was the only sign of infection. The fungus Basidiobolus ranarum was reported from G. carolinensis in Arkansas and Missouri (Nickerson and Hutchison, 1971). Bd has been recorded from Illinois (Phillips et al., 2014) and Oklahoma (Watters et al., 2016; Marhanka et al., 2017). Ranavirus is reported from G. carolinensis in North Carolina (Lentz et al., 2021) and Oklahoma (Davis et al., 2019). A hind limb malformation was reported by Adams et al. (2008). Gastrophryne carolinensis is parasitized by the nematode Gyrinicola batrachiensis (Pryor and Greiner, 2004). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. Selenium, mercury, silver, zinc, chromium, lead, cadmium, copper, germanium, cobalt, nickel, aluminum, and

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arsenic are highly toxic to embryos of G. carolinensis (Birge, 1978; Birge et al., 1979; Birge et al., 1983). The LC50 (7 day) for ­these metals are all 50 m) riparian zones (Rudolph and Dickson, 1990). The species also appears to tolerate prescribed burns and is frequently found in fire-­ maintained habitats (e.g., Moseley et al., 2003). This species is considered Endangered in Mary­land and Threatened in Illinois and Kansas.

Information in the lit­er­a­ture on “Gastrophryne olivacea” from Arizona actually refers to G. mazatlanensis. IDENTIFICATION

Adults. The Sinaloan Narrow-­mouthed Frog is a small, grayish-­brown, olive-­brown, or tan frog with smooth skin and a pointed snout. Dark spots are scattered on the body dorsally and dorsolaterally ­behind the eye; tiny white spots (dots) also may be pre­sent. A dark line may run from the eye to the snout. A small skin fold (transverse fold) is pre­sent across the back of the head ­behind the eyes. Dark spots and blotches may be located on the limbs to varying extent. The limbs are short, and the tympanum is not vis­i­ble. An enlarged metatarsal tubercle is located on the foot. Adults are usually 19–36 mm SUL and reach a maximum size of ca. 41 mm SUL (Murphy, 2019). Males are smaller than

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mazatlanensis was described by Taylor (1943) but synonymized with G. olivacea by Hecht and Matalas (1946). Using mitochondrial DNA analyses, Streicher et al. (2012) confirmed that G. mazatlanensis was distinct from G. olivacea by a 2.2% ge­ne­tic difference. ADULT HABITAT

The Sinaloan Narrow-­mouthed Frog is a species of desert scrub and arid grasslands. Brennan and Holycross (2006) list the habitat as Lower Colorado River Desert Scrub to Madrean Evergreen Woodland in Arizona. It occurs in the Pajarito and Patagonia Mountains in the latter habitat type (Murphy, 2019). AQUATIC AND TERRESTRIAL ECOLOGY

Distribution of Gastrophryne mazatlanensis

females, and males have a dark throat and tubercles on the chin. Larvae. In Sonora, tadpoles are grayish tan dorsally and have dark tail tips. The eyes are spaced wide apart on the sides of the head (Rorabaugh and Lemos-­Espinal, 2016). Eggs. No information is available, but presumably similar to G. olivacea.

The species is nocturnal, and is active on the surface only ­after desert rains. During the day, the frog hides ­under surface debris near ­water and in burrows. Burrows may be shared with tarantula spiders that ignore the frog and provide protection against predators. Most activity occurs during the summer monsoon season, but frogs have been observed from April to October in Sonora (Rorabaugh and Lemos-­Espinal, 2016). CALLING ACTIVITY AND MATE SE­L ECTION

The call is a short high-­pitched “peep.” This is followed by a “high-­pitched, wheezy buzz that lasts 1–4 s” (Brennan and Holycross, 2006). Calling occurs from very small crevices near the breeding site, in dense nearby vegetation, or while floating at the surface of the breeding site. Males have a ventral adhesive gland that enables them to amplex females.

DISTRIBUTION

This species occurs from Nayarit and southern Sinaloa north through southwestern Chihuahua and much of central Sonora, México, into southern Arizona, including the Pajarito Mountains (Streicher et al., 2012; Bezy and Cole, 2014). The most northern rec­ords are from extreme southeastern Maricopa County (Vekol Valley) and southwestern Pinal County. Other impor­tant distributional references include Arizona (­Sullivan et al., 1996b; Brennan and Holycross, 2006; Murphy, 2019; Holycross et al., 2021), Chihuahua (Lemos-­Espinal and Smith, 2007a), and Sonora (Rorabaugh and Lemos-­Espinal, 2016).

BREEDING SITES

As with G. olivacea, breeding occurs in a wide association of wetlands, including tinajas, temporary pools, large permanent ponds, inundated alluvial floodplains, roadside ditches, and even in man-­made habitats such as stock tanks and irrigation

FOSSIL REC­O RD

None. SYSTEMATICS AND GEOGRAPHIC VARIATION

Gastrophryne in Arizona w ­ ere considered to be conspecific with G. olivacea ­until relatively recently. Gastrophryne

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Newly hatched tadpoles of Gastrophryne mazatlanensis. Buenos Aires National Wildlife Refuge, Arizona. Photo: Brent Sigafus

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Adult Gastrophryne mazatlanensis. Photo: Dennis Suhre

Amplexing Gastrophryne mazatlanensis. Photo: Brent Sigafus

pits (­Sullivan et al., 1996b). According to Murphy (2019), breeding sites are often located in dense stands of floodplain mesquite.

ant attacks as individuals feed. Other frogs kept in containers with G. mazatlanensis have succumbed to this secretion (Murphy, 2019). Predators include the Garter Snake Thamnophis cyrtopsis (Jones and Hensley, 2020).

REPRODUCTION

POPULATION BIOLOGY

Several hundred eggs are oviposited ­after summer rains. As with other Gastrophryne, eggs are deposited in a thin surface film. Metamorphs are 9–10 mm SUL (Murphy, 2019).

According to Rorabaugh and Lemos-­Espinal (2016), sexual maturity is reached at 1–2 yrs at 19 mm SUL. DISEASES, PARASITES, AND MALFORMATIONS

DIET

The nematodes Aplectana incerta and A. itzocanensis ­were found in Gastrophryne mazatlanensis from Arizona (Goldberg et al., 1998b). Bd has been detected on this species along the Mexican-­US border (as G. olivacea: Sigafus et al., 2014).

As with other Gastrophryne, this species feeds on ants and termites.

SUSCEPTIBILITY TO POTENTIAL STRESSORS

LARVAL ECOLOGY

No information is available for Arizona.

No information is available. PREDATION AND DEFENSE

This species has a skin secretion that may irritate predators and some ­people. The secretion also likely functions to deter

STATUS AND CONSERVATION

Gastrophryne olivacea (Hallowell, 1857 “1856”    ) Western Narrow-­mouthed Frog

texana, Gastrophryne texense, Gastrophryne texensis, Microhyla areolata, Microhyla carolinensis olivacea, Microhyla olivacea Gastrophryne olivacea has had a confusing nomenclatorial history, especially in Texas where several species ­were described (Strecker, 1909) that ­were subsequently referred to this species. Hecht and Matalas (1946) and Nelson (1972) reviewed the nomenclatural history of Gastrophryne. Information in the lit­er­a­ture on “Gastrophryne olivacea” from Arizona actually refers to G. mazatlanensis. Although commonly called a toad, this species is in the ­family Microhylidae and thus is not a true toad.

ETYMOLOGY

olivacea: from Latin, referring to the olive-­green dorsal coloration. NOMENCLATURE

Synonyms: Engystoma areolata, Engystoma texense, Engystoma olivaceum, Gastrophryne areolata, Gastrophryne

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No information is available.

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Gastrophryne olivacea 413

IDENTIFICATION

Adults. This is a round, smooth-­skinned, pointy-­snouted frog with a dorsal uniform pale tan to olive to greenish-­gray coloration that is paler in females than males. ­There are no prominent dorsal markings. Temperature and moisture affect the darkening of the skin, with frogs in low moisture and high temperature situations appearing lighter than well-­ hydrated frogs at lower temperatures. ­There is a transverse dermal groove ­behind the eyes that may be covered by a fold of skin. This fold becomes more pronounced when the frog is irritated. Black spots are pre­sent dorsally and on the upper surface of the hind legs. Venters are white with slightly gray to brown pigmentation in the gular region of some individuals. No webbing is pre­sent between the digits and no tympanum is evident. Gastrophryne olivacea is sexually dimorphic. Males have a black throat patch that never ­really dis­appears, although it fades during the nonbreeding season. The color of the throat even may intensify during the breeding season. Males develop conical chin tubercles along the ventral surface of the lower jaw, and the subgular throat patch appears wrinkled (when calling, the throat sac is much larger than the frog’s head). ­These latter 2 characteristics are evident only during the breeding season. Males also develop a nuptial adhesive gland covering the thorax and inner surface of the arms (Fitch, 1956b) that secretes an adhesive substance that helps the male stay attached to the female during amplexus. Some breeding females may have prominent tubercles in the perianal region (Taylor, 1940). Males are generally smaller than females. ­There are lit­er­a­ture rec­ords for Texas males and females (means 24.1 mm SUL; 27.4 mm SUL respectively) (Blair, 1955a). Goldberg (2018f) recorded males from 26 to 31 mm SUL (mean 27.9 mm) and females from 2 to 33 mm SUL (mean 27.8 mm) based on Texas museum specimens. Blair (1955a) included mea­sure­ment data for males from 18 locations in 8 Texas counties. He noted that ­there appeared to be a geographic trend in size, with smaller individuals in the east and larger individuals in the west. Loftus-­Hills and Littlejohn (1992) recorded males with a mean SUL of 26.3 mm (range 23–30.8 mm), also from Texas. The largest recorded G. olivacea mea­sured 43 mm SUL (Powell et al., 2016). Larvae. Tadpoles are grayish brown or grayish tan dorsally and may or may not have an indistinct stripe on the side of the tail. The venter is light in coloration but may be mottled with gray. Viewed from the side, the head comes to a fairly sharp point. The jaws do not have keratinized teeth, and no oral disk is pre­sent. A single spiracle is pre­sent that opens midventrally, and the tail tip is dark. Aspects of the morphology of the tadpole ­were described by Nelson and Cuellar (1968).

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Eggs. Individual eggs are small and brown or black dorsally and white ventrally. The dia­meter of the outer envelope is 2.8–3 mm, but ­there may be a second envelope near the vitellus. The vitellus is 0.8–0.9 mm (Livezey and Wright, 1947). Eggs are oviposited in a surface film whose jelly may be truncated (i.e., flattened or appearing to be cut off) or nontruncated. Total clutches normally contain from 500 to >2,100 eggs, but partial clutches are usually deposited in a single jelly film. DISTRIBUTION

The Western Narrow-­mouthed Frog occurs from western Missouri and Kansas south throughout much of Oklahoma and eastern, central, and western Texas, then as far south as Tamaulipas and San Luis Potosí, México. It occurs eastward along the Missouri River into central Missouri and enters western Arkansas along several river courses. To the north, it occurs in a very small area of south central and southeastern Nebraska. Populations occur in southeastern Colorado and the adjacent Oklahoma Panhandle, and the range extends northward from central Texas into the eastern Texas Panhandle and adjacent Oklahoma. The range is prob­ably more contiguous than the range map indicates pending further field work. Populations occur in the Sierra Vieja and Guadalupe mountains of west Texas and in southern New Mexico. ­There is a low level of ge­ne­tic diversity within G. olivacea, suggesting rapid northern geographic expansion during the late Pleistocene or early Holocene that is ongoing (Streicher at al., 2012).

Distribution of Gastrophryne olivacea

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414  Microhylidae

Impor­tant distributional references include: range-­wide (Nelson, 1972), Arkansas (Trauth et al., 2004), Colorado (Hammerson, 1999), Kansas (Smith, 1934; Collins, 1993; Collins et al., 2010), México (Lemos-­Espinal and Smith, 2007b), Missouri (Johnson, 2000; Daniel and Edmond, 2006), Nebraska (Lynch, 1985; Ballinger et al., 2010; Fogell, 2010; Geluso and Wright, 2010), New Mexico (Degenhardt et al., 1996), Oklahoma (Sievert and Sievert, 2006), and Texas (Dixon, 2000, 2013; Tipton et al., 2012; Davis and LaDuc, 2018; Fielder et al., 2020). FOSSIL REC­O RD

Pleistocene Rancholabrean fossils are known from Texas and Sonora, México (Holman, 2003). The transverse dorsal prominence of the ilium is lower and less triangular than in G. carolinensis, making discrimination pos­si­ble. SYSTEMATICS AND GEOGRAPHIC VARIATION

Frogs of the genus Gastrophryne are derived phyloge­ne­ tically from the South American clade of microhylid frogs (Meijden et al., 2007). ­There are 4 species within this genus (Streicher et al., 2012), but only 3 enter the United States. The genus Gastrophryne is most closely related to frogs of the genus Hypopachus (Nelson, 1972; Streicher et al., 2012). Two species described from Texas (areolata, texensis) ­were synonymized with Gastrophryne olivacea and may represent hybrids, variations in color patterns, or misidentified individuals (see discussions in Smith, 1933; Burt, 1937). Laboratory-­based hybridization between G. olivacea and Chiasmocleis panamensis is not very successful (Littlejohn, 1959), and with Hypopachus may produce a very few metamorphs, although hatching success is low (Wilks and Laughlin, 1962). Natu­ral hybridization with G. carolinensis has been reported in a narrow contact area in eastern Texas and Oklahoma (Hecht and Matalas, 1946), although the larger G. carolinensis males are reluctant to clasp the much smaller G. olivacea females. Strecker’s (1909) description of Microhyla areolata appears to represent ­these hybrid individuals. Pattern phenotypes intermediate between G. olivacea and G. carolinensis are known from a number of locations, even where the species are not sympatric. Therefore, pattern variation does not result from hybridization alone (contrary to Hecht and Matalas, 1946), and care must be taken when identifying “hybrids” based solely on morphological grounds (Nelson, 1972). Most pattern variation relates to the extent of mottling on the belly, sides, and throat. Laboratory crosses with G. carolinensis may produce tadpoles (Blair, 1950). In addition to sexual size dimorphism, Nelson (1972) showed that males from lowland habitats ­were smaller than males from upland habitats. This size variation was consistent

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throughout the range of the species. He did not have sufficient data to determine ­whether this trend applied to females, however. ADULT HABITAT

Gastrophryne olivacea is a frog of the G ­ reat Plains and grassland deserts (such as Big Bend National Park; Dayton and Fitzgerald, 2006), where it occupies a wide variety of habitats and is not particularly restricted to any special type of habitat. It prefers open wooded habitats with abundant cover, such as provided by the flat limestone slabs, rock outcrops, and bluffs of eastern Kansas (Fitch, 1956b; Heinrich and Kaufman, 1985). In other areas, the species occupies mesquite flats, prairie grasslands, and overgrazed desert scrub dominated by mesquite, creosote bush, and desert grasses, sometimes with l­ittle surface cover (Smith, 1950; Degenhardt et al., 1996). It is not generally a frog of extensive river floodplains, although it may occupy sloughs along stream courses. Adjacent habitat provides terrestrial cover and foraging sites, and the frogs even may occupy cultivated fields and other agricultural habitats if ants are in abundance. The species occupies under­ground cracks and crevices, including burrows of mammals such as pocket gophers (Geomys bursarius), prairie dogs (Cynomys ludovicianus), and voles (Microtus ochrogaster), or lizards (Crotaphytus collaris) (McAllister and Tabor, 1985; Lomolino and Smith, 2003). Considering that this species often is found in limestone areas, it is not surprising that individuals are found in caves (Black, 1973a) or near the entrances to caves. The species occupies habitats below 1,342 m in New Mexico (Degenhardt et al., 1996) and 1,525 m in Colorado (Hammerson, 1999), and is considered a frog of the lowlands. TERRESTRIAL ECOLOGY

The Western Narrow-­mouthed Frog may be more widespread and abundant than it appears, due to its fossorial or semifossorial habits. Gastrophryne olivacea is active in warm weather, especially ­under conditions of high humidity or during or immediately ­after warm rains. Most activity occurs nocturnally when frogs leave their shelters to forage, but diurnal activity may occur when night temperatures are too low for normal activity or during cloud-­cover and precipitation. Preferred temperatures range from 24 to 31°C. Surface activity occurs at temperatures >20°C, although activity at as low as 16°C has been observed (Fitch, 1956b). The upper limit for activity is 37.6°C. This species is thus one of the most sensitive anurans in the ­Great Plains to cold, but one of the least sensitive to heat. Activity occurs throughout the warm season. For example, G. olivacea have been recorded from April to October in Kansas, with rare captures in December (Fitch, 1956b).

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Activity also may be curtailed by drought, causing them to cease surface movements several months ­earlier than normal. Fitch (1956b) characterized 4 movement patterns within G. olivacea: routine, short daily movements, home range shifts occurring over long or short periods ­either gradually or abruptly, movements to and from breeding ponds, and dispersal by recent metamorphs. It appears ­these small frogs are capable of considerable movement. For example, Freiburg (1951) noted a female that moved 229 m straight-­ line distance over a period of 55 days between recaptures; a male moved 102 m in 3 days. Movements >30 m appear quite common, and females and males move to the same extent and distances, although Fitch (1956b) suggested males ­were more vagile than females. This species is often found far from the nearest source of ­water, and Fitch (1956b) recorded the greatest distance moved as 610 m from ­water. During cold or other­wise inhospitable weather, this species takes refuge in loose, moist soil ­under large flat rocks, ­under the bark of fallen trees, in rock or mud crevices, in and ­under moist logs, and ­under surface debris (Freiburg, 1951). They ­will often occupy the burrows of other animals, such as invertebrates, lizards, and mammals. Strecker (1909) found them overwintering with 2 copperheads (Agkistrodon contortrix), the lizard Scincella lateralis, and the frogs Lithobates sphenocephalus (?), Dryophytes cinereus, D. squirellus, and Incilius nebulifer. Many other vertebrates and invertebrates share their terrestrial refugia (Freiburg, 1951). Like most frogs that are active both day and night, Western Narrow-­mouthed Frogs are generally photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are particularly sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). Western Narrow-­mouthed Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calls are heard throughout the warmest season of the year (generally May–­August), but they occasionally are heard as early as March in Texas (Blair, 1961b) and April in Oklahoma (Bragg, 1943a, 1950a). In Kansas, calls have been recorded from June to September (Freiburg, 1951), and females with eggs have been found in July and August (Smith, 1934). In New Mexico, calling occurs from June to August (Degenhardt et al., 1996). Thus, calling may precede and follow the primary dates of egg deposition (Wiest, 1982). Calling occurs most often from dusk ­until about midnight, ­after which it tapers off appreciably. Occasional

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calls may be heard during daylight, but most individuals withdraw to nearby retreat sites during daylight hours. Precipitation is necessary to initiate large choruses, especially at the start of the breeding season. In Kansas, such storms usually occur from May to mid-­June (Fitch, 1956b), although frogs may appear as early as mid-­April. Large choruses are sometimes heard ­after a considerable time with no chorusing; ­these ­later choruses are triggered by heavy rainfall, especially ­after substantial periods without precipitation. This suggests an opportunistic breeding strategy to take advantage of optimum breeding conditions. Wiest (1982) noted calling at 11.3–27.2°C over a 34 day period in Texas. Calls have been described as “a high shrill buzz of some 2 to 3 seconds” (Smith, 1934) or as insect-­like and difficult to describe (Fitch, 1956b). Calls are uttered at a mean rate of 7 per minute. They do not have much carry­ing capacity, and it is difficult to hear the calls at any distance away from a breeding site. The mean call duration ranges from 0.9 to 3.7 sec at a mean frequency of 2.6–5.0 kHz, depending on temperature (Bragg, 1950c; Blair, 1955b; Nelson, 1972, 1973; Loftus-­Hills and Littlejohn, 1992). The harmonic interval is 155–280 Hz. Call frequency tends to increase with temperature. ­There may be geographic differences in call characteristics, but ­these differences may reflect differences in species rather than conspecific differences. For example, the frequency is less in Arizona Gastrophryne (now considered to be a dif­fer­ent species, G. mazatlanensis) and the call duration is shorter when compared with Texas populations of G. olivacea. Nelson (1972) provided data on call characteristics from throughout the species’ range, but note that Arizona populations are not conspecific with the “G. olivacea” from Nelson’s study. Males call while floating in shallow ­water among vegetation or hidden among dense grass clumps. In Kansas, they have also been heard calling from ­under protective rock ledges adjacent to breeding sites. The calling posture is similar to that described for G. carolinensis, with males almost vertical as they call. ­There does not appear to be any calling territoriality, as many males may call in close proximity to one another (Bragg, 1943a; Freiburg, 1951). Amplexus may occur shortly ­after calling begins. Females approach calling males and likely initiate breeding. Amplexus is axillary. Males are able to remain clasped to females using their arms and by the adherence of dermal secretions produced from glands located on the male’s venter; essentially, they become glued to one another (Fitch, 1956b). ­After oviposition, it takes up to 15 min for the secretions to break down and allow the pair to separate. A short (0.5 sec), nasal-­pitched buzz has been described from G. olivacea occupying terrestrial refugia in Texas. Individuals making this call ­were not associated with a

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breeding chorus, and Dayton (2000) interpreted the calls as agonistic territorial calls. Such calls could establish calling hierarchies prior to the start of breeding choruses, or they may have nothing to do with reproductive be­hav­ior. BREEDING SITES

Breeding occurs in a wide association of wetlands, including tinajas, temporary pools, large permanent ponds, inundated alluvial floodplains, roadside ditches, and even man-­made habitats such as stock tanks and irrigation pits (Smith, 1934; Anderson et al., 1999; Dayton and Fitzgerald, 2006). Campbell (1934) noted calling from pools at the bases of trees and suggested that the tree roots offered diurnal retreats. They frequently call from bison wallows, but ­these sites may not hold ­water long enough for successful metamorphosis (Gerlanc, 1999). In the desert, breeding occurs in pools located in dense stands of mesquite shrubs in river washes. In the ­Great Plains, this species breeds in wetlands with a pH of 6.9–7.5, low aluminum concentration, low dissolved oxygen content (0.7 ppm), and dense vegetation that creates a considerable amount of cover (Anderson et al., 1999). REPRODUCTION

Based on histology, Goldberg (2018f) noted that males and females are capable of breeding from February to August in Texas. The smallest mature male was 26 mm SUL, whereas the smallest mature female was 24 mm SUL. Breeding occurs over an extended season and is largely dependent upon precipitation. Some females breed during the first rains, whereas ­others stagger breeding throughout the summer as do many xeric-­adapted anurans. Thus, each precipitation event results in some breeding. Eggs are attached to vegetation in shallow ­water at the ­water’s surface in an irregular oblong film. Clutch sizes are quite variable, with reports of several hundred to more than 2,000 eggs per female. For

Eggs of Gastrophryne olivacea. Big Bend, Texas. Photo: Dana Drake

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example, clutch size ranged from 59 to 2,174 eggs (mean 1,166) in 28 Texas females (Henderson, 1961). Except for a single female with 59 eggs, all other clutches contained >519 eggs. In Kansas, clutch size ranged from 532 to 1,217 with most clutches containing >800 eggs during the breeding season; 50 and 75 tiny, immature eggs ­were found in spent ovaries in August, presumably ­after the breeding season (Freiburg, 1951). Wright and Wright (1949) give a clutch size of 645. Some of the literature-­based variation in clutch size may reflect counts of partial clutches. Cold temperatures inhibit hatching, and eggs must be at temperatures >17–19°C in order to hatch (Hubbs and Armstrong, 1961; Ballinger and McKinney, 1966). Prolonged drought and decreased rainfall are prob­ably the biggest threats to successful reproduction, as they eliminate breeding ponds and/or decrease the duration of the hydroperiod, leaving eggs or larvae to perish. Gastrophryne olivacea may skip breeding during periods of drought. LARVAL ECOLOGY

Hatching occurs in 2–3 days following egg deposition. Tadpoles are filter feeders on phyto-­or zooplankton, and their mouthparts are incapable of rasping algae or other plant surface detritus. The length of the larval period is dependent on temperature and resource availability, and ­there are reports of 24 days (Fitch, 1956b) and 30–50 days to complete development (Wright and Wright, 1949). Newly metamorphosed froglets range from 10 to 12 mm SUL (Wright, 1932; Wright and Wright, 1949). Froglets with a small vestige of a tail stump just prior to metamorphosis may be 14.5–16 mm SUL as they aggregate around the margins of a pond (Fitch, 1956b). DIET AND FEEDING

The Western Narrow-­mouthed Frog is an ant specialist ­ rematogaster, (termed myrmecophagy), especially foraging on C

Tadpole of Gastrophryne olivacea. Photo: Greg Sievert

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a subterranean species (Smith, 1934; Tanner, 1950; Freiburg, 1951; Fitch, 1956b). Like Gastrophryne carolinensis, individuals ­will often sit near an ant-­hill or colony and feed on ants leaving the mound or forage near it (Tanner, 1950; Carpenter, 1954b). Freiburg (1951) found from 7 to 71 (mean 32.5) ant heads in the digestive tracts of 15 adults in Kansas. Other prey includes small beetles and Speodesmus falcatus (Sickled Cave Millipedes) (Owen and Pustka, 2021). PREDATION AND DEFENSE

The primary defenses of this species involve its cryptic coloration, secretive be­hav­ior, and noxious secretions. Gastrophryne olivacea tends to walk, run, or move in short hops, so saltation cannot be used to escape predators. They are capable of short, elusive bursts of movement followed by complete immobility. The Western Narrow-­mouthed Frog has noxious skin secretions that make it less prone to predation than other small frogs and also protects it from attack by the ants on which it feeds. It ­causes a burning or irritating sensation to delicate tissues such as the eyes and mucous membranes of the mouth and throat, and entangles attacking ants. If a frog is handled, copious secretions are exuded, making it slippery. As in G. carolinensis, secretions are produced by granular glands located throughout the skin, which first appear late in development coinciding with the eruption of the forelimbs. Frogs at breeding ponds are often very difficult to locate as they call hidden in vegetation. At the approach of a predator (or observer), they stop calling and do not resume ­until the predator has moved several meters away. They may remain immobile or dive ­under ­water and hide in dense bottom vegetation. Tadpoles also remain immobile in the ­water column or hang motionless at the ­water’s surface. Known or suspected predators include other frogs (Lithobates blairi, L. catesbeianus), snakes (Agkistrodon contortrix, Thamnophis cyrtopsis, T. marcianus, T. proximus), birds (Loggerhead Shrike), and possibly shrews (Blarina brevicauda) (Anderson, 1942; Freiburg, 1951; Fouquette, 1954; Fitch, 1956b; Fitch, 1960; in Collins, 1993; Kelehear et al., 2017; Ford, 2020). Eggs are fully palatable to fish (Grubb, 1972).

Adult Gastrophryne olivacea. Photo: David Dennis

phose late in the breeding season) may require a second growing season to reach maturity. ­Whether temporal differences in age at first reproduction reflect sexual differences in the timing of maturity (e.g., females reaching maturity ­later than males), individual variation in growth rates (Fitch, 1956b), or differences in the timing of metamorphosis remains to be clarified. The bulk of a population consists of 3-­yr-­olds. Gastrophryne olivacea in their fourth year are 30–38 mm SUL (males to 38 mm and females to 42 mm), and some frogs may live even longer. Frogs have been held as captives for 6 years. Like many observations on sex ratios in frogs, males appear to be more abundant than females, but this perception could be biased by sampling and differences in activity patterns between the sexes. In Kansas, Freiburg (1951) found sex ratios of 1:1–4:1 males per female depending upon month sampled. COMMUNITY ECOLOGY

POPULATION BIOLOGY

In Kansas, sexual maturity is reached by 25 mm SUL (Freiburg, 1951). This size may be reached late in the first season following metamorphosis in frogs that metamorphose early in the season. Indeed, Gastrophryne olivacea can reach 19–28 mm SUL before the first overwintering period, due to rapid growth (Freiburg, 1951; Fitch, 1956b). At least some G. olivacea prob­ably breed initially about 1 year following metamorphosis, whereas ­others (e.g., ­those that metamor-

Dodd_Canada_int_5pgs_B3.indd 417

The ranges of G. carolinensis and G. olivacea overlap in eastern Texas and Oklahoma. Although hybridization may occur in areas of contact, differences in call structure and body size help to minimize gene exchange. In zones of contact with G. carolinensis, males and females are both smaller than they are in areas where the 2 species are not sympatric (Blair, 1955a). The structure of the advertisement calls is also dif­fer­ent between the species, with G. olivacea having calls of higher frequencies and longer duration than

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418  Microhylidae

the calls of G. carolinensis (Blair, 1955b; Awbrey, 1965; Nelson, 1972; Loftus-­Hills and Littlejohn, 1992). Hybrids have call characteristics intermediate between the parental species. In choice tests, females prefer the calls of males of their own species to ­those of G. carolinensis (Awbrey, 1965). Decreases in body mass occur in larval G. olivacea when reared with Scaphiopus couchii larvae (Dayton and Fitzgerald, 2001). Data suggest that S. couchii are able to outcompete G. olivacea larvae and may exclude them from shallow ­water desert pools in some areas, such as Big Bend National Park. Western Narrow-­mouthed Frogs form an unusual association with tarantula spiders (Dugesiella hentzi) (Blair, 1936; Yeary, 1979; Hunt, 1980; Dundee, 1999; Dundee et al., 2012). Tarantulas and Gastrophryne occupy the same burrows. Despite their small size, the narrow-­mouthed frogs are not attacked by the spider. If a frog predator attacks, the frogs ­will huddle ­under the spider for protection. Likewise, narrow-­mouthed frogs ­will not eat baby tarantulas, no ­matter how small they are. In return for protection, the frogs eat ants and termites that might other­wise attack the spider’s eggs and hatchlings; they also find shelter in a favorable environment. This unlikely cohabitation is an example of mutualistic symbiosis. Many Gastrophryne may occupy the burrow of a single spider.

Breeding habitat of Gastrophryne olivacea. Photo: Mike Forstner

et al., 2017; Vhora and Bolek, 2013). Bd has been recorded from Oklahoma (Watters et al., 2016, 2019; Marhanka et al., 2017). Ranavirus also is reported from G. olivacea in Oklahoma (Davis et al., 2019; Smith et al., 2019). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. DISEASES, PARASITES, AND MALFORMATIONS

No evidence of ectoparasites has been reported. Freiburg (1951) recorded unidentified nematodes in the gastrointestinal tract. The coccidian protozoan Isospora fragosum occurs in G. olivacea (Upton and McAllister, 1988), as does an isosporan similar to I. neos (McAllister and Upton, 1987). The cestode Cylindrotaenia americana and the nematodes Aplectana hamatospicula, A. incerta, and Cosmocercoides dukae also occur in this species (McAllister and Upton, 1987; McAllister

STATUS AND CONSERVATION

Hypopachus variolosus (Cope, 1866) Sheep Frog

variolosus: from the Latin variola meaning ‘variegated’ or ‘with small spots’ in reference to the dorsal pattern.

Hypopachus cuneus, Hypopachus cuneus nigroreticulatus, Hypopachus globulosus, Hypopachus inguinalis, Hypopachus maculatus, Hypopachus ovis, Hypopachus oxyrhinus, Hypopachus oxyrrhinus ovis, Hypopachus oxyrrhinus taylori, Hypopachus oxyrhynchus, Hypopachus oxyrrhinus, Hypopachus reticulatus, Hypopachus seebachi, Hypopachus variolosus inguinalis, Systoma variolosum

NOMENCLATURE

IDENTIFICATION

ETYMOLOGY

Synonyms: Engystoma inguinalis, Engystoma variolosum, Hypopachus alboventer, Hypopachus alboventer reticulatus, Hypopachus caprimimus, Hypopachus championi,

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The Western Narrow-­mouthed Frog appears to be widespread and generally abundant, though secretive. As with all amphibians, populations may have been lost due to habitat destruction and alteration. A single study indicated no appreciable effects from a wildfire (Brown et al., 2014). This species is considered Endangered in New Mexico and of Special Concern in Colorado.

Adults. Sheep Frogs are globular in shape with pointed snouts and no neck. ­There is a small fold of skin just ­behind the eyes, and ­there is no vis­i­ble tympanum. The ground color is

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greenish brown to olive, usually with an orange or yellow mid-­dorsal stripe. The ground color shades to brown laterally, and ­there may be dark reticulations on the side. An oblique white band is pre­sent from the eye to the shoulder. The skin is smooth and exceptionally thick, but ­there are scattered white-­tipped tubercles on the dorsal and lateral surfaces and on the forelegs and the margins of the lower jaw. ­These tubercles may serve an adhesive function. The venter is mottled gray or yellowish with a midventral white line; dark reticulations may or may not be pre­sent. ­There are well-­ developed palmar tubercles at the base of the first and fifth toes of the hind foot. Throats are black in males. Adults from Texas are 27–42.5 mm SUL (mean 35.8 mm) (Nelson, 1974). Wright and Wright (1949) reported males as 25–37.5 mm SUL and females as 29–41 mm SUL. Lemos Espinal and Smith (2007a) mention individuals from Guerrero to 47 mm SUL. Larvae. Tadpoles are medium sized and darkly pigmented dorsally but immaculate ventrally on the body. This pigmentation difference is sharply delineated laterally. The tail musculature is darkly pigmented. The dorsal tail fin is highest halfway down the tail and evenly but darkly pigmented or with dark spots ­running together. The ventral tail fin has scattered spots but is not as darkly or evenly pigmented as the dorsal tail fin. The tail ends in a sharp tip. Jaws are without keratinized structures, and the oral disk and labial teeth are absent. The edges of the labial flaps are scalloped or with distinct papillae. Larvae are 27–30 mm TL. Tadpoles are illustrated by Wright (1929). Eggs. Eggs are black dorsally and white ventrally with a single jelly envelope 1.5–2 mm in dia­meter; the vitellus alone is 1 mm in dia­meter (Mulaik and Sollberger, 1938). Eggs are deposited in a surface film. DISTRIBUTION

The Sheep Frog occurs in the United States only in extreme south Texas (15 counties) along the Gulf Coast. The native range of the species extends southward to Costa Rica, but the Texas population appears disjunct from ­those farther south in México. Impor­tant distributional references include Dixon (2000, 2013), Judd and Irwin (2005), and Tipton et al. (2012).

Distribution of Hypopachus variolosus

and G. olivacea produce a small percentage of larvae and metamorphs (Wilks and Laughlin, 1962). ADULT HABITAT

Hypopachus variolosus is a frog of humid open woodlands, mature coastal brushlands, and pasturelands with short grass cover. Throughout much of its range, it is a frog of Tamaulipan thornscrub and savannas. TERRESTRIAL ECOLOGY

Sheep Frogs are fossorial and rarely observed and then only during heavy rains from April to October (Mulaik and Sollberger, 1938; Judd and Irwin, 2005). ­These authors reported ­these frogs in rodent burrows, pack rat middens, and the hollows ­under trees. They burrow backward and move deeper as moisture decreases. ­After metamorphosis, froglets migrate to upland burrows, often stopping along the way in cow dung, surface detritus, and litter (in Wright and Wright, 1933). Dispersal occurs in high humidity and during rainfall, even diurnally. Brown (1950) reported that individuals ­were found at a depth of 76 cm at the bottom of postholes during the dry season.

FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Originally described as Hypopachus cuneus by Cope (1889), the Texas Hypopachus was synonymized with H. variolosus by Nelson (1974). The genus is closely related to the genus Gastrophryne. Laboratory crosses between H. variolosus

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CALLING ACTIVITY AND MATE SE­L ECTION

Males call while floating in the ­water or from the shorelines of ponds within 24 hrs ­after heavy rainfall during the warm season; Garrett and Barker (1987) give the breeding season as March–­September. The call of H. variolosus is described as a bleat, similar to a sheep, and lower in tone than that of G. olivacea (Mulaik and Sollberger, 1938). Calls last

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420  Microhylidae

0.8–6 sec (mean range 1–4.5 sec) and are repeated at >15 sec intervals. The dominant frequency is 1,500–3,900 Hz (mean 2,620) with a harmonic interval of 100–220 Hz (mean 180) (Nelson, 1973, 1974). Calling occurs from 18 to 30°C. BREEDING SITES

Breeding occurs in shallow temporary to permanent pools. They also breed in roadside ditches, railroad right-­of-­way ditches, and pothole basins (Judd and Irwin, 2005). REPRODUCTION

Reproduction is stimulated by heavy rainfall but can occur in response to the sudden irrigation of agricultural fields (Judd and Irwin, 2005). Oviposition occurs within 24 hrs of rainfall. The eggs are oviposited in a surface film that is loosely held together. The film has a somewhat truncate or flattened appearance similar to the surface films of Gastrophryne. Mulaik and Sollberger (1938) reported a single clutch of 700 eggs, whereas Wilks and Laughlin (1962) obtained 452 eggs from a single female. Eggs hatch in ca. 12 hrs a­ fter deposition (Mulaik and Sollberger, 1938).

Adult Hypopachus variolosus. South Texas. Photo: Clint Guadiana

LARVAL ECOLOGY

The larval period lasts about 30 days (Mulaik and Sollberger, 1938). Newly metamorphosed froglets are 11–16 mm SUL. DIET

The diet of adults consists of ants, termites, and small flies (Mulaik and Sollberger, 1938; Garrett and Barker, 1987). Larvae are likely generalist feeders of organic and inorganic ­matter.

Breeding habitat of Hypopachus variolosus. Willacy County, Texas. Photo: Gary Nafis

PREDATION AND DEFENSE

Predators include Eastern Ribbon Snakes (Thamnophis saurita) (Wright and Wright, 1949).

SUSCEPTIBILITY TO POTENTIAL STRESSORS

POPULATION BIOLOGY

STATUS AND CONSERVATION

No information is available.

Sheep Frogs are considered Threatened in the state of Texas but are abundant in several counties in south Texas (Judd and Irwin, 2005). No information is available on status or population trends.

DISEASES, PARASITES, AND MALFORMATIONS

The mite Caeculus hypopachus was described from this species (Mulaik, 1945).

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No information is available.

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F ­ amily Rhinophrynidae

Rhinophrynus dorsalis Duméril and Bibron, 1841 Mexican Burrowing Toad ETYMOLOGY

dorsalis: from the Latin dorsalis, possibly referring to the dorsal pattern. NOMENCLATURE

Synonyms: Rhinophryne rostratus, Rhinophrynus rostratus IDENTIFICATION

Adults. Rhinophrynus dorsalis is a rotund frog with very short but power­ful limbs, pustulose and loose skin, a short ­little head, tiny eyes with vertical pupils, and no tympanum. The ground color is dark brown to black and appears somewhat translucent. ­There is a distinctive, light yellow to red-­orange mid-­dorsal line down the back with similarly colored small spots on both sides. Venters are lighter than the dorsum but still dark in coloration. The hind legs are partially enclosed in loose body skin. Large, spade-­like tubercles are pre­sent on the rear feet. The snout is protuberant. ­These frogs reach about 50–90 mm SUL at maturity, with females larger than males (in Fouquette, 2005). Larvae. The head is broad and depressed, with small eyes. The mouth is unique in that it is a wide slit bordered by 11 short barbels, giving it the appearance of whis­kers. ­There are no keratinized structures associated with the jaws, and the oral disk and labial teeth are absent. The spiracles are paired and laterally positioned unlike in other native North American frogs. The tail fins and musculature are well developed and taper to a narrow-­pointed tip. Larvae grow to 39.5 mm TL. Orton (1943) describes the tadpole and illustrates the unique oral barbels surrounding the mouth. Eggs. The eggs have not been described.

known only from Jim Hogg, Starr, and Zapata counties, Texas. Impor­tant distributional references include James (1966) and Dixon (2000, 2013). FOSSIL REC­O RD

Pleistocene fossils of R. dorsalis are known from a cave in Tamaulipas, México (Holman, 2003). The genus also is represented by Eocene fossils from Saskatchewan (Holman, 1963) and Oligocene fossils from Florida (Blackburn et al., 2019). SYSTEMATICS AND GEOGRAPHIC VARIATION

Rhinophrynus is a monotypic genus allied within a highly derived, primitive group of frogs, the aglossal pipids of the suborder Pipoidea. The diploid chromosome number is 22 (Bogart and Nelson, 1976). ADULT HABITAT

Garrett and Barker (1987) noted that Mexican Burrowing Toads prefer areas with loose soil for digging. In Texas, the species is found in agricultural areas and gardens in the

DISTRIBUTION

The Mexican Burrowing Toad occurs from south Texas to northwestern Costa Rica. In the United States, R. dorsalis is

Dodd_Canada_int_5pgs_B3.indd 421

Distribution of Rhinophrynus dorsalis

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422  Rhinophrynidae

Matamoran District of the Tamaulipan Biotic Province. The area in Texas consists of rolling hills of sand and gravel over thin soils. ­These hills are interspersed by deep arroyos fed from shallow ravines and washes. Vegetation is arid to semiarid trees and shrubs. James (1966) recorded breeding in arroyos surrounded by very thick tangles of thorny vegetation consisting of cacti, acacias, and retama (Parkinsonia aculeata). TERRESTRIAL ECOLOGY

As its common name implies, R. dorsalis is almost entirely fossorial and rarely comes to the surface. They can remain for long periods under­ground. Surface activity is opportunistic and occurs only in response to rainfall. They dig backward into the soil with digging spades on the rear feet. They also twist their bodies extensively back and forth and inflate their bodies to facilitate penetration into the soil. Body inflation allows them to maintain a cavity space as the soil ­settles around them. They can affect some forward motion using their widely spaced, power­ful spatulate forelimbs and tuberculate hands (Trueb and Gans, 1983). Nothing is known concerning migration or movement patterns.

Tadpole of Rhinophrynus dorsalis. Photo: Seth Patterson

CALLING ACTIVITY AND MATE SE­L ECTION

The call of R. dorsalis is a loud but low-­pitched guttural moan (James, 1966; Garrett and Barker, 1987) described as a low-­pitched “wh-­o-­o-­o-­e” by Crump (2015). Calling occurs from within burrows, with males emerging ­after heavy rains to form large breeding choruses. Males then call from the surface of the ­water or on soil or among short vegetation bordering flooded sites. When calling, James (1966) likened males to inflated balloons floating on the ­water. James (1966) mentioned very large and loud breeding choruses.

Adult Rhinophrynus dorsalis. Photo: Seth Patterson

BREEDING SITES

Breeding takes place in shallow temporary pools and flooded areas formed ­after heavy rainfall. ­These ponds may contain extensive vegetation. James (1966) also mentioned breeding in stock tanks and drainage ditches. REPRODUCTION

Eggs are deposited in small clumps, which then separate and float to the surface. No information is available on clutch size or any other aspect of reproduction. LARVAL ECOLOGY

Adult Rhinophrynus dorsalis peering from burrow. Photo: Seth Patterson

The duration of the larval period is at least 2 months (in Fouquette, 2005). In other parts of the species’ range, tadpoles form large aggregations that are maintained by visual or olfactory cues.

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Rhinophrynus dorsalis 423

DIET

DISEASES, PARASITES, AND MALFORMATIONS

Rhinophrynus dorsalis is an ant and termite specialist. The species has a number of unique morphological specializations for feeding on ­these prey, including epidermal armor on the snout, ornately folded buccal and esophageal lining folds, an ability to “double-­close” the lips, and a specialized tongue apparatus for ­handling small prey in subterranean burrows (Trueb and Gans, 1983). The spade-­like tubercles on the hind feet are used to dig into termite mounds. Larvae feed on phytoplankton.

Parasitic opalinid protozoans have been reported from this species in México. SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Texas protects this species as Threatened, but t­ here is nothing known concerning its status and trends (Fouquette, 2005). COMMENT

PREDATION AND DEFENSE

When threatened, the Mexican Burrowing Frog inflates its body thus obscuring the head and limbs. ­There is no information available on predators. POPULATION BIOLOGY

No information is available.

Dodd_Canada_int_5pgs_B3.indd 423

The Ma­ya of México considered R. dorsalis to be messengers, attendants, and musicians to the chacs, the rain deities. The call of this frog summoned the chacs to tip their gourds, thus causing rain to fall from the sky. According to Crump (2015), the chacs controlled all of nature, including ­water, clouds, thunder, lightning, earth, the heavens, plants, and animals. As such, this species was of considerable importance to the Ma­ya.

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F ­ amily Ranidae

Lithobates areolatus (Baird and Girard, 1852) Crawfish Frog ETYMOLOGY

areolatus: from the Latin aesopus meaning ‘dwarf.’ The name refers to the hunchback appearance of the frog. NOMENCLATURE

Conant and Collins (1998): Rana areolata Dubois (2006): Lithobates (Lithobates) areolatus Fouquette and Dubois (2014): Rana (Lithobates) areolata Synonyms: Rana areolata, Rana circulosa IDENTIFICATION

Adults. Lithobates areolatus is a large, stocky frog with a large head and prominent eyes. The ground color is light gray to brown, with the dorsum covered by large, round dark spots that are surrounded by light halos. The spots are interspersed by dark reticulations. Dorsolateral folds are pre­sent, and may or may not be prominent. The skin is smooth or rugose. The rear limbs are banded with dark markings bordered by light stripes. Venters are off-­white. During the breeding season, males have enlarged thumbs and paired vocal sacs, which are evident ­behind and below the tympanum. Females are generally larger than males. Rec­ords include males 64–117 mm SUL (mean 93 mm) and females 75– 118 mm SUL (mean 98 mm) in Illinois (Smith et al., 1948), and adults 57–110 mm SUL, also in Illinois (Smith, 1961). In another Illinois population, males averaged 82.8 mm SUL (range 71–90 mm) and females ­were 89.6 mm SUL (range 79–102 mm) (Redmer, 2000). Based on museum specimens from Oklahoma, Goldberg (2019a) reported males from 68 to 98 mm SUL (mean 79.8 mm) and females from 76 to 91 mm SUL (mean 84 mm). Length is positively correlated with body weight and age. Collins et al. (2010) report a Kansas individual of 122 mm SUL. Larvae. The tadpole of L. areolatus is large (to 65 mm TL; Smith et al., 1948) and deep bodied, and dark brown to

Dodd_Canada_int_5pgs_B3.indd 424

vari­ous shades of green (light, dark, olive) dorsally. Viewed dorsally it is ovoid and chunky with a rounded, blunt snout. Upper and lower tail fins may or may not be heavi­ly marked with small diffuse spots, and the dorsal tail fin is attenuated and elongate. In general, the tail fin is equally pigmented above and below the tail musculature. The lower jaws are wide, and the mouth’s beak is broadly marginated. Throats are unpigmented or uniformly pigmented. The gut may or may not be vis­i­ble through a white venter. Bragg (1953) offers a number of additional identification characters. The larvae are similar to ­those of leopard frogs (L. blairi, L. pipiens, L. sphenocephalus) and may be impossible to distinguish. Eggs. Eggs are medium in size and dark colored. Two gelatinous envelopes are pre­sent; the outer envelope is 3.3–5 mm in dia­meter and the inner envelope is 1–3.3 mm in dia­meter; the vitellus is 1–3.8 mm in dia­meter (Smith, 1934; Livezey and Wright, 1947; Smith et al., 1948; Bragg, 1953). Egg mea­sure­ments are smaller in Illinois than in Kansas, indicating the potential for regional variation, although differences in the condition of the individuals may account for the disparity. Eggs are deposited in an ovoid, firm jelly mass and are attached to vegetation below the ­water’s surface. ­These masses may be invaded by algae, giving them a green appearance. DISTRIBUTION

The distribution of the Crawfish Frog is not contiguous within its range, with many disjunct populations occurring from the Midwest to the Texas Gulf Coast. The species occurs from southern Iowa, Illinois, and western Indiana, south to central Mississippi, southeastern Arkansas, and areas in Louisiana. Disjunct populations occur in western and central Alabama, northwestern Louisiana, and southeastern Indiana. High ge­ne­tic differentiation between sites in southwest and southeast Indiana suggest historical isolation of ­these sites from one another rather than recent isolation (Nunziata et al., 2013). In the west, the species occurs in eastern Kansas and Oklahoma south to the Texas Gulf Coast. This species is often associated with river floodplains, such as along the Arkansas and Missouri rivers.

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Lithobates areolatus 425

Distribution of Lithobates areolatus. Dark gray indicates extant populations; light gray indicates extirpated populations.

Impor­tant distributional references include: Alabama (Holt, 2015), Arkansas (Bacon and Anderson, 1976; Trauth et al., 2004), Illinois (Smith, 1961; Phillips et al., 1999; Palis, 2018), Indiana (Swanson, 1939; Minton, 2001; Brodman, 2003; Engbrecht and Lannoo, 2010; Engbrecht et al., 2013), Iowa (Bailey, 1943; Christiansen, 1998; Hemesath, 1998), Kansas (Collins et al., 2010), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Missouri (Johnson, 2000; Daniel and Edmond, 2006), Oklahoma (Sievert and Sievert, 2006), Tennessee (Redmond and Scott, 1996; Niemiller and Reynolds, 2011), and Texas (Dixon, 2000, 2013). Range-­wide distribution is covered by Lannoo and Stiles (2020a). FOSSIL REC­O RD

Fossils of L. areolatus are reported from Miocene (Kansas) and Pliocene (Texas) deposits (Holman, 2003). SYSTEMATICS AND GEOGRAPHIC VARIATION

Crawfish Frogs are in the Nenirana clade of North American ranid frogs, a group that includes L. palustris, L. capito, and L. sevosus (Hillis and Wilcox, 2005). They are only distantly related to the Scurrilirana clade. Two subspecies have been identified historically, L. areolatus areolatus (Southern Crawfish Frog) and L. areolatus circulosus (Northern Crawfish Frog). Lithobates a. areolatus is said to be larger

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with a more rounded snout and a greater dorsal rugosity than L. a. circulosus. Other differences involve body proportions and the prominence of the dorsolateral folds. Bragg (1953) found some of ­these characters useful in separating subspecies (but see Lannoo et al., 2018, for corrections), but noted a large degree of overlap. Calls of ­these subspecies are similar and, as with other Nenirana, are characterized by long snore-­like calls with lower frequencies and short interpulse periods. More information on distribution and molecular phylogeny are needed to better delineate the relationships and validity of ­these taxa. For example, the subspecies do not ­really line up as “northern” and “southern,” more like eastern and western in many re­spects. Lithobates areolatus does not appear to hybridize with other Lithobates in nature. ­Under laboratory conditions, ­viable hybrids have been produced between ♂ L. sphenocephalus and ♀ L. areolatus. However, crosses between ♂ L. areolatus and ♀ L. sphenocephalus ­were inviable or only produced a small number of larvae (Cuellar, 1971). Other laboratory studies produced a small to large number of larvae in crosses between L. areolatus and L. blairi, L. sphenocephalus, L. palustris, L. pipiens, and a number of Mexican Lithobates (Moore, 1949a; McAlister, 1961; Mecham, 1969; Cuellar, 1971; Hillis, 1988). Crosses between L. berlandieri, L. catesbeianus, L. clamitans, or L. sylvaticus and L. areolatus ­were unsuccessful (Moore, 1949a, 1955; Mecham, 1969; Cuellar, 1971). ADULT HABITAT

Southern Crawfish Frogs are found in open damp areas, wooded valleys, oak-­hickory woodlands, floodplains, and open and brushy fields (Bragg, 1953; Clawson and Baskett, 1982). The Northern Crawfish Frog is a species of tall-­grass prairies and grasslands in some areas (Johnson, 2000), although Engbrecht and Lannoo (2010) noted that many of ­these areas ­were forested prior to Eu­ro­pean settlement. Thompson (1915) recorded the species in Illinois in an area of rolling hills and agriculture with few streams and no natu­ral ponds or lakes. In eastern Kansas, Crawfish Frogs are found in gentle terrain with deep (0.9–1.5 m) clay soils, whereas in western Tennessee, the species was considered common in flat sandy or semiswampy areas (Gentry, 1955). Areas with shallow soils and intensive mechanized agricultural activity are not favored. TERRESTRIAL ECOLOGY

During the nonbreeding season, the Crawfish Frog is largely fossorial, although frogs may be active throughout the day and night at burrow entrances and even leave the burrow for long periods of time (Hoffman et al., 2010; Stiles et al., 2017b). For example, Engbrecht and Lannoo (2012) noted that frogs ­were

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Lithobates areolatus in burrow entrance. Indiana. Photo: Nate Engbrecht

Lithobates areolatus entering burrow. Indiana. Photo: Nate Engbrecht

active from ca. 10–11 hrs daily in a study lasting from August to September. Activity is seasonal at the burrow, with most activity around its entrance occurring in May and September when frogs are active around the clock. In early spring and late fall, the frogs are primarily diurnal, but they switch to a nocturnal activity cycle during the summer. Activity is best explained as a daily response to temperature (average temperature of 19.4°C) and vapor pressure gradient (a minimum VPG of 1.2 hPa and a maximum VPG of 23.3 hPa) (Stiles et al., 2017b). Emergence from burrows occurs at temperatures as low as 9°C (but normally about 16.5°C), and frogs reenter burrows at about 13.3°C, but as low as 9.7°C (Engbrecht and Lannoo, 2012). Total precipitation should be at least 3.1 mm. ­There are no differences in adult and juvenile preferences for ­these environmental variables. Crawfish Frogs are philopatric to upland crayfish burrows, which they inhabit for >10 months of the year (Heemeyer et al., 2012). They have an excellent ability to home to par­tic­u­lar crayfish burrows, and ­will use the same migratory pathways to reach their burrow (Heemeyer and Lannoo, 2012). Migration movements may be quite long

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since breeding sites can be >1 km from the home burrow. Of 48 radio-­tracked adults, migration distances averaged 0.5 km, with 1 frog moving 1.2 km. Crawfish Frogs prefer the burrows of the crayfish Cambarus diogenes—­which is often not associated with wetlands—­but also have been found in mammal burrows, ­under logs, and in tunnels in road cuts. Some burrows may serve as more or less perma­ thers are used along migratory nent homes, whereas o pathways or while foraging away from the home burrow (Heemeyer et al., 2012). Home ranges average only 0.05 m2. Occupied crayfish burrows may extend 90 cm below the surface (Thompson, 1915; Bailey, 1943). The sides of the burrow tend to become slick as the frog moves up and down, and the bottom of the burrow ­will be full of frog feces. They likely ­will utilize any burrow available, including tree root channels. More than 1 frog may occupy a burrow, and burrow occupancy can be extensive; Bailey (1943) reported that all 12 burrows observed within a 15 m radius ­were occupied by 1 or more frogs. Juveniles in par­tic­u­lar are quick to use preexisting burrows, which helps to minimize moisture loss; they are capable, however, of digging their own burrows using only their hind limbs (Parris, 1998). In Illinois, individuals have been found in agricultural fields 15 cm below the surface not long ­after the breeding season (Smith et al., 1948). Crawfish Frogs forage at the entrance of their burrows as ambush predators. They frequently leave the burrow and forage in terrestrial litter. For example, Hoffman et al. (2010) reported a Crawfish Frog in Indiana that spent 87% of its time (237 hrs) outside the burrow and only 13% (36 hrs) in its burrow over a period of just ­under 13 days. Another frog spent as much as 91% of its time (162 of 179 hrs) within its burrow. From midsummer to late summer, periods of inactivity

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tended to be shorter than periods of activity outside burrows. As the season progresses, activity periods become shorter. Hoffman et al. (2010) observed 1 frog outside its burrow as late as 13 December. Emergence in spring is related to increasing temperatures and rainfall. Several days with temperatures ≥15°C followed by moderate rainfall are enough for frogs to emerge and move to breeding sites. Cool temperatures early in the spring may inhibit activity, even if breeding is underway. Activity occurs from March to September in Kansas (Collins et al., 2010) and from March to October in Missouri (Johnson, 2000). Frogs may occasionally be killed by cold temperatures as they overwinter in their burrows (Heemeyer and Lannoo, 2011). Colonization of new habitats does not occur once a frog has settled into a burrow; that is, they do not move long distances between upland burrows. Instead, new habitats are colonized by the adults in varying wetland breeding sites across years and through juvenile dispersal (Heemeyer and Lannoo, 2012). As a prairie species, Crawfish Frogs ­were undoubtedly subject to periodic fires. Fires do not seem to have any effect on survivorship, but the frogs change their be­hav­ior in post-­burn areas that have become more exposed. Nighttime be­hav­ior does not change, but during the day frogs stay in or very close to their burrow entrances instead of foraging away from them as they do in vegetated unburned grasslands (Engbrecht and Lannoo, 2012). They also emerge ­later in the day in burned habitats than they do in unburned habitats. CALLING ACTIVITY AND MATE SE­L ECTION

Rainfall is necessary to stimulate movement to breeding ponds, and calling occurs nocturnally from January to July, depending on weather and latitude. Calling may occur immediately ­after arrival, but ­there is conflicting information about ­whether it is necessary for rainfall to have occurred during the previous 24 hr period for chorusing to take place (Williams et al., 2012a, 2013). The apparent conflict may arise from the dif­fer­ent environmental conditions and sampling time frames between studies. ­There is no relationship between drought conditions (high temperatures, low precipitation) and ­either the onset of breeding or peak breeding periods (Lannoo and Stiles, 2017). As with other species, Crawfish Frogs do not occupy ­every available habitat within a region. For example, Williams et al. (2012a) found them at 22 of 45 grasslands in 2010 and 24 of 45 grasslands in 2011 during call surveys. Detection was associated with time of night (>60% 3 hrs ­after sunset), precipitation (before and during the survey), and temperature (60% >8°C), and occupancy was associated with grassland size (mean 57 ha, range 11–133 ha). Chorusing

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begins at about 19:00 hrs, peaks before midnight, and tapers off at ca. 03:00 hrs (Williams et al., 2013). The call of both subspecies is a deep, guttural snore, but Bragg (1953) also noted a short call, which he likened to a dog barking. Bragg (1953) stated that the call of L. a. areolatus had lower dominant frequencies than L. a. circulosus, with a longer duration that ­rose in frequency and often terminated in a low whistle. In fact, the opposite is true (Lannoo et al., 2018). Lithobates a. areolatus actually have higher frequency calls of shorter duration. Both subspecies’ calls rise in frequency, and Lannoo et al. (2018) never heard a terminal, high frequency whistle in ­either subspecies. ­These authors reported a mean dominant frequency of 1,133 Hz, a mean call duration of 0.51 s, a mean interpulse period of 0.017 s, and a mean of 32.9 pulses per call in L. a. areolatus at 15°C in Texas. In L. a. circulosus, ­these values ­were a mean dominant frequency of 806 Hz, a mean call duration of 0.84 s, a mean interpulse period of 0.019 s, and a mean of 40 pulses per call at 15°C in Indiana. Calls are made from shallow ­water while sitting on the bottom, although some ­will call from the bank. In deeper ­water, males call while floating on the surface. Bragg and Smith (1942) noted that calling occurred while the male was “sprawled out” in the ­water. The call produces a distinct vibration as the male’s paired vocal sacs beat the ­water’s surface. Chorusing is most intense during and shortly ­after nightfall, and during peak chorusing can last all night. Occasional males ­will call during cloudy or rainy days. Males ­will call at temperatures as low as 2–8°C, but temperatures ≥13°C are necessary to initiate calling (Busby and Brecheisen, 1997). McKnight and Ligon (2016) found no correlation between rainfall and calling in this species, but found that frogs called at temperatures between 11.7 and 16°C; calling peaked at 22:00 hrs. Males arrive 5–14 days prior to females and remain at the breeding site throughout the breeding season, which lasts 22–55 days (Smith et al., 1948; Bacon and Anderson, 1976; Busby and Brecheisen, 1997; Trauth et al., 2004). Peak chorusing lasts only 3–4 consecutive nights and usually occurs about a week ­after chorus initiation. McKnight and Ligon (2016) recorded chorusing males on only 9 and 17 nights at 2 ponds in Oklahoma. Choruses often are small and consist of 8°C for breeding to commence, with most activity occurring at >12°C. Diminished calling and breeding may occur at ≤6°C (Smith et al., 1948; Bacon and Anderson, 1976). Eggs are oviposited in a plinth 120–210 mm in dia­meter and ca. 25 mm thick. The mass is attached ­under ­water, and ­water depths are usually 150–200 mm (Busby and Brecheisen, 1997). Egg masses may be communally located (Bragg, 1953), with Busby and Brecheisen (1997) reporting 22 masses in a 1 m2 area. Clutch size ranges between 2,000 and 7,000 eggs per egg mass. Specific counts include 3,192–6,807 in Illinois (Smith et al., 1948) to >7,000 in Indiana (Wright and Myers, 1927), a mean of 6,320 in Indiana (Lannoo and Stiles, 2020a), 3,208–6,807 (mean 4,868) in Illinois (Redmer, 2000), single clutches of 2,233 in Arkansas (Trauth et al., 1990), and 3,801 in Oklahoma (Bragg, 1953). Clutch size is strongly positively correlated with female SUL but only weakly positively correlated with age; clutch size is negatively correlated with ovum size (Redmer, 2000). Hatching occurs in 3–4 days, although Johnson (2000) indicated that it took 7–10 days. LARVAL ECOLOGY

Larvae grow rapidly, with Bragg (1953) reporting growth rates of 1.06 mm/day during the first 15 days. Growth rates gradually decrease to 0.9–0.6 mm/day. Bragg (1953) estimated that the growth rate for 59 days following the initiation of feeding (4 days ­after hatching) was 0.76 mm/day. The length of the larval period was 63 days ­after hatching in Bragg’s (1953) study, and 56–61 days in Indiana, depending on the extent of intra-­and interspecific competition (Lannoo and Stiles, 2020a). Recent metamorphs are 22–30 mm SUL (Wright and Myers, 1927; Cagle, 1942; Smith, 1961). Bailey (1943) reported finding large larvae in early spring in Iowa, suggesting that some larvae overwinter and transform the following summer. In a within-­pond field enclosure experiment, Williams et al. (2012b) noted that larvae reared at low densities ­were larger at metamorphosis and survived better than larvae reared at high densities. When released terrestri-

Tadpole of Lithobates areolatus. Photo: Laurie Vitt

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ally, large juveniles resulting from large tadpoles had higher survivorship than small juveniles resulting from small tadpoles. Increased density also extends the larval period, which could be detrimental during drought years. Whereas ca. 98% of eggs hatch successfully, only about 1% survive to metamorphosis (Lannoo and Stiles, 2020a). DISPERSAL

Juveniles disperse randomly from breeding sites. Lannoo et al. (2017) found that average daily movements varied between 27 and 35 m depending upon year, with maximum daily movements of 114–297 m. DIET

The diet includes beetles, spiders, crickets, ants, millipedes, centipedes, and small crayfish (Thompson, 1915; Smith, 1934; Smith et al., 1948). The type of beetles eaten indicates nocturnal feeding. Any animal that can fit into the mouth ­will likely be consumed. PREDATION AND DEFENSE

When disturbed, Crawfish Frogs quickly retreat down their burrows (e.g., Engbrecht et al., 2012; Lannoo and Stiles, 2020a). The hind limbs in par­tic­u­lar are used as wedges to prevent extraction from the burrow. Crawfish Frogs also have a defensive posture whereby the head is lowered and the body is elevated and inflated to prevent ingestion (Thompson, 1915; see Fig. 4.7 in Lannoo and Stiles, 2020a). Altig (1972b) also provided a photo­graph and noted that the posture was assumed in response to snake and small mammal predators and was often accompanied by a loud scream. This species has a noxious odor that may serve an antipredator function. They are extremely wary in breeding ponds and readily become quiet and submerge at the approach of an intruder. Crawfish Frogs also have antimicrobial peptides in their skin which may assist in protecting the frog against microorganisms (Ali et al., 2002). Larvae and postmetamorphs are likely eaten by a variety of invertebrate and vertebrate predators, but no information is available. Adults are eaten by hognose snakes (Heterodon platirhinos) (Engbrecht and Heemeyer, 2010), Black Racers (Coluber constrictor) and Common Garter Snakes (Thamnophis sirtalis) (Stiles et al., 2017b; Lannoo and Stiles, 2020a), Plain-­bellied Watersnakes (Nerodia erythrogaster), Loggerhead Shrikes (Baecher et al., 2014), and raccoons (Heemeyer et al., 2010). POPULATION BIOLOGY

Populations of Crawfish Frogs at the northern portion of their range in Indiana have a high degree of ge­ne­tic diversity based on analyses of heterozygosity at 10 microsatellite loci

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Adult Lithobates areolatus circulosus. Photo: David Dennis

(Nunziata et al., 2013). The degree of population subdivision was low regionally, and ge­ne­tic differentiation was related to geographic distance, as expected. At a very local level, ­there was no ge­ne­tic differentiation among individuals from nearby (within 250 m) breeding ponds, with only slight differentiation from a pond 750 m away. Not surprisingly, sex ratios at breeding ponds are highly skewed ­toward males. For example, Smith et al. (1948) recorded a sex ratio of 6.08:1 in an Illinois pond, and Lannoo and Stiles (2020a) reported a sex ratio of 1.22:1 in 1,102 breeding adults from 2009 to 2016 at 2 Indiana sites. In terms of overall survivorship, juveniles only had ca. 3% survivorship in a long-­term Indiana study, whereas adult survivorship was ca. 70%, but with considerable annual variation (Lannoo and Stiles, 2020a). Lannoo and Stiles (2017) found that ­there was no relationship between drought and adult survivorship. However, the lowest survivorship estimates occurred during the wettest years, which may relate to the frog’s occupancy of crayfish burrows. Drought is also inversely correlated with body condition in both males and females and, as such, drought affects fecundity with serious demographic consequences. In wet years, females oviposited an estimated 7,554 eggs/ female, whereas in drought years, only 4,907 eggs/female ­were estimated to have been produced. Thus, the average difference in fecundity between wet and dry years was 2,647 eggs/female (Lannoo and Stiles, 2017). At some ponds, populations appear to have been quite large at one time. For example, Cagle (1942) noted a pond in southern Illinois that contained 500 breeding adults that deposited 179 egg masses; the pool only mea­sured 36 m × 91 m. In 3 other pools, ­there ­were 115, 125, and 75 egg masses. Cagle (1942) also reported large numbers of adults being collected (289 from several small ponds) and removal of the 179 egg masses. Such large-­scale collecting

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could have adversely affected the local population. Smith (1961) noted breeding aggregations that contained as many as several hundred animals, and Barbour (1971) noted that breeding populations ­were large in western Kentucky. Males mature ­earlier than females, with a mean male age of 3.53 yrs (range 2–5 yrs) and a female mean of 3.83 yrs (range 3–5 yrs) in Illinois (Redmer, 2000). Longevity can range to at least 10 yrs (Lannoo and Stiles, 2020a). COMMUNITY ECOLOGY

Lithobates areolatus may breed in ponds occupied by other ranid species. In experimental ponds, interspecific competition with L. blairi and L. sphenocephalus resulted in an increased larval period and a decreased body mass of metamorphic L. areolatus (Parris and Semlitsch, 1998). However, neither intra-­nor interspecific competition with L. clamitans or L. sphenocephalus larvae affected the survivorship of L. areolatus larvae (Lannoo and Stiles, 2020a). Competition has a density and species-­specific component. For example, survivorship of L. blairi larvae decreased in the presence of L. areolatus larvae at high density. In contrast, the presence of L. areolatus seems to facilitate growth of L. sphenocephalus larvae. In experimental ­trials with the salamander Ambystoma texanum, the presence of salamanders severely reduced the survivorship of L. areolatus tadpoles. As tadpole density decreased as a result of predation, the survivors ­were actually longer and heavier and metamorphosed quicker than tadpoles raised at the same initial density but without the predator (Lannoo and Stiles, 2020a; Stiles et al., 2020). Still, when this predator was absent, high densities of intra-­ and interspecific competitors (see below) had no effect on larval survivorship of Crawfish Frogs. In a further series of mesocosm and outdoor enclosure experiments, Stiles et al. (2020) showed that intra-­and interspecific (with larval L. clamitans and L. sphenocephalus) competition reduced the size (both length and mass) of Crawfish Frog larvae at metamorphosis when densities ­were high. ­There ­were also density-­dependent fitness effects on the size of newly transformed metamorphs that affected length and mass (high larval densities produced small metamorphs) of Crawfish Frogs. ­These effects ­were carried over into juvenile survival, adult size, and breeding adult numbers—­ small metamorphs produced smaller adults with reduced survivorship, which resulted in fewer breeding adults. Seemingly paradoxically, a decline in population size might be compensated for by an increase in adult body size, and thus increase reproductive potential and adult fitness, at least up to a point (Stiles et al., 2020). Complex interactions such as ­these are impor­tant components of larval amphibian communities and lend insights into conservation strategies.

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DISEASES, PARASITES, AND MALFORMATIONS

The fungal pathogen Batrachochytrium dendrobatidis (Bd) has been found on L. areolatus from Oklahoma (Watters et al., 2016) and Indiana (Lannoo and Stiles, 2020a). A total of 53% of adults tested over a 2 yr period tested positive for Bd in Indiana, with more adults exiting the pond testing positive than adults entering the pond. Mortality occurred especially when frogs contained >10,000 zoospores. Infection rates ­were near zero at the end of summer but increased to >25% following overwintering in crayfish burrows; rates then doubled again following breeding, when mortality occurred (Kinney et al., 2011). Further reports of Bd in this species are in Terrell et al. (2014b), who documented that drought reduced mortality and the intensity of infection, but not the prevalence of Bd infection. Ranavirus was reported in captive-­raised L. areolatus released in Indiana (Stiles et al., 2016). Amelia, ectromely, and micropthalmia have been reported in this species (Terrell et al., 2014a; Stiles et al., 2017a). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Chemicals. Death occurred in about 17 hrs at 30 mg/L of carbaryl, a broad-­spectrum insecticide, and ­there ­were significantly reduced larval activity levels at 2.5 mg/L (Bridges and Semlitsch, 2000). This is among the most sensitive species to carbaryl among ranids so far tested. STATUS AND CONSERVATION

The Crawfish Frog was once widely distributed and reasonably common, but through the years many populations have been lost due to habitat loss and degradation. For example, many rec­ords in Indiana are >50 yrs old, and ­there are no current rec­ords for 7 historic counties (Minton, 2001; Brodman, 2003; Engbrecht and Lannoo, 2010; Engbrecht

Breeding site of Lithobates areolatus in Illinois. Photo: John Palis

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et al., 2013; Lannoo and Stiles, 2020a). Engbrecht et al. (2013) estimated that ­there ­were fewer than 1,000 Crawfish Frogs remaining in Indiana. Christiansen (1998) reported no recent rec­ords at 9 historic locations in Iowa. Minton (2001) indicated that populations began declining in about 1970 for no obvious reason. In contrast, Florey and Mullin (2005) noted increasing detection during road-­call surveys in Illinois from 1986 to 1989; Phillips et al. (1999) ­later indicated many populations ­were no longer extant in much of the state. In southern Illinois, however, Palis (2014b) found them at 90 of 220 breeding sites during auditory surveys, including 80 sites where they had not been previously recorded. Threats to Crawfish Frogs include pond draining, road mortality, and habitat loss and fragmentation. They may be evicted from burrows during plowing and mowing, and this appears to have occurred commonly in the past (Hurter, 1911; Thompson, 1915). Even in 1913, the species was reported as becoming increasingly rare in Illinois due to agricultural activities (Thompson, 1915). Minton (2001) also noted that the species was used for food in Indiana and Illinois, where it was easily collected. Although pre­sent in some agricultural settings, the species is more abundant in natu­ral habitats and absent from mined habitats (Anderson and Arruda, 2006); populations in Indiana ­were destroyed by mining (Minton et al., 1982). However, Terrell et al. (2014a) and Stiles et al. (2017a) observed 2,592 individuals at 2 ponds over a 2 yr period at a mine reclaimed site in Indiana. Bragg (1953) noted that the lights and noise from a nearby highway did not seem to disturb the frogs.

In order to conserve this species, breeding and upland habitats must be protected and properly managed, especially by the prevention of lowered ­water ­tables due to groundwater pumping. Management could include prescribed burns to maintain optimal habitat, but must involve the maintenance of crayfish burrows, fishless breeding ponds, and dispersal corridors (Lannoo et al., 2017). Crawfish Frogs ­will occupy newly created, large, open-­canopied mitigation wetlands as long as a source population is nearby (Palis, 2007; Baecher et al., 2018; Drayer et al., 2020). In several cases, Crawfish Frogs seemed to prefer created wetlands to natu­ral wetlands (Baecher et al., 2018; Drayer et al., 2020). Johnson (2000) noted that the Missouri Department of Conservation was in the pro­cess of constructing fishless ponds in managed prairies for this species. Engbrecht et al. (2013) suggested that ­there is enough grassland habitat remaining in Indiana that the species could be restored in many of its former sites. Williams et al. (2013) provide guidelines for conducting road surveys as part of monitoring using auditory cues. This species is considered Endangered in Indiana (Engbrecht and Lannoo, 2010) and Iowa. A popu­lar account of the biology of this species and the research conducted on it by Michael Lannoo, Rochelle Stiles, and their colleagues is in their 2020 book The Call of the Crawfish Frog. Much additional information on the biology of Crawfish Frogs is in this enjoyable book. Lannoo and Stiles (2020a) further review the status and habitat requirements of this species in Indiana, and suggest management recommendations that are likely apropos elsewhere throughout its range.

Lithobates berlandieri (Baird, 1859) Rio Grande Leopard Frog

References to this species in the lit­er­a­ture are often incorrect. For example, Smith and Sanders (1952) discussed the distribution of “Rana pipiens berlandieri” in Texas but included individuals from what is now known to be Lithobates blairi. The nomenclatural history of this species is discussed by Hillis (1988). Readers should verify the locations of individuals in order to ensure correct species identification. Degenhardt et al. (1996) noted that the date of the description of this species is often incorrectly listed as 1854.

ETYMOLOGY

berlandieri: a patronym honoring Jean Louis Berlandier (1805–1851). Berlandier was a French naturalist who worked for the Mexican Government surveying eastern Texas in 1827–1828. His extensive collections ­were the first made in Texas. NOMENCLATURE

Conant and Collins (1998): Rana berlandieri Dubois (2006): Lithobates (Lithobates) berlandieri Fouquette and Dubois (2014): Rana (Lithobates) berlandieri Synonyms: Rana austricola, Rana halecina, Rana pipiens berlandieri, Rana virescens

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IDENTIFICATION

Adults. This light brown, olive-­green, tan, or grayish-­colored leopard frog has well-­developed dorsolateral folds that are discontinuous and curve medially ­toward the rear of the frog. The inset dorsolateral fold is singular and similar to the primary dorsolateral fold (see Fig. 1(1) in Pauly et al., 2020a). A supralabial stripe is pre­sent, but it is indistinct anterior to the eye. Light borders may or may not surround the dark dorsal spots, but they are usually faint when

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pre­sent. Usually, no spots are pre­sent on the nose in front of the eyes, although a few individuals may have a single spot or faint mottling. A light tympanic spot is usually absent. Throats and the anterior portions of the chest are mottled, especially in older frogs, and the level of mottling appears to increase ­after dark (Sanders and Smith, 1971). Venters are cream colored. The posterior part of the thigh has sharply contrasting dark and light reticulations, sometimes with a bluish background coloration. Males have paired external vocal sacs and prominent vestigial oviducts. Hind toes are webbed. Males are smaller than females, with a mean body length of 64.4 mm SUL for males and 73.5 mm SUL for females (in Degenhardt et al., 1996). Based on museum specimens, males had a mean of 69.5 mm SUL (range 44–94 mm) and females had a mean of 77.5 mm SUL (range 47–103 mm) (Hughes and Meshaka, 2018); also based on museum specimens, Texas males averaged 73.7 mm SUL (range 63–83 mm) and females 88.6 mm SUL (range 74–102 mm) (Goldberg, 2020d). The maximum SUL is 114 mm SUL (Brennan and Holycross, 2006). Hughes and Meshaka (2018) suggested that adults from northern populations ­were smaller than ­those from southern populations, based on museum specimens from Texas. Larvae. The tadpole is long and slim and has a variable color pattern. The overall color is a dark grayish black in small tadpoles that becomes olive with a yellowish cast as the tadpoles grow. The lateral line system is generally obscure, especially on the head. Spots on the side of the body are gray. The tail is moderately deep with a narrow, pale gray tail muscle. The belly and throat are white. The tail pattern consists of discrete, dark olive and pale or golden spots, or it may have a strikingly dark olive reticulated pattern enclosing pale spots. The iris is gold and contains black flecks. The tadpole was described by Hillis (1982), Scott and Jennings (1985), and Rorabaugh and Lemos-­Espinal (2016). Eggs. Eggs are deposited in a firm jellied mass that is about 70–90 mm across (in Degenhardt et al., 1996). The eggs likely are bicolored, white on the bottom and dark on top, although Dayton et al. (2007) describe them simply as black. The egg capsule is 3.2–5.1 mm in dia­meter (mean 3.9 mm) (Grubb, 1972). In Texas, ovum dia­meter averaged 1.31 mm (range 0.9–2.0 mm) (Hughes and Meshaka, 2018). Ovum size is not correlated with body size or clutch size. DISTRIBUTION

Lithobates berlandieri occurs naturally throughout much of central, southern (including North Padre Island; Duran and Hall, 2013), and southwest Texas and southeastern New Mexico (lower Pecos drainage of Eddy County) into the

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Distribution of Lithobates berlandieri. Populations of L. berlandieri in Arizona and California are introduced.

adjacent Mexican states of Coahuila and Chihuahua and well south to Veracruz and Oaxaca. Although “L. berlandieri” has been identified as far south as Nicaragua, it is likely that frogs in southern México and Central Amer­i­ca represent diverse taxa (reviewed by Rorabaugh, 2005b). The species has been introduced into the southwestern United States across southern Arizona (Gila, Salt, and Agua Fria drainages) and along the southern Colorado River of Arizona and California (Clarkson and Rorabaugh, 1989; Platz et al., 1990; Rorabaugh et al., 2002; Murphy, 2019; Holycross et al., 2021). It has also spread into interior southeast California as far as Indio (Goodward and Wilcox, 2019; Pauly et al., 2020a), Utah, and México (Kraus, 2009; Lemos Espinal and Smith, 2007b). Impor­tant distributional references include: Platz et al. (1990), Degenhardt et al. (1996), Dixon (2000, 2013), Brennan and Holycross (2006), Tipton et al. (2012), Davis and LaDuc (2018), Murphy (2019), and Pauly et al. (2020a). Although L. yavapaiensis has been extirpated in California since 1965, leopard frogs resembling this species have been observed in habitats it formerly occupied in the state. Despite similarities, ge­ne­tic analy­sis has confirmed that ­these leopard frogs are the invasive L. berlandieri. Morphological characters used to distinguish ­these species overlap in the frogs now found in California, which complicates field identification. Lithobates berlandieri in this region show extensive variation in the condition of the inset of the dorsolateral folds, a key character, more so than they do in their native range (see Fig. 1 in Pauly et al., 2020a).

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Lithobates berlandieri 433

FOSSIL REC­O RD

Pleistocene fossils referred to as “Rana pipiens” by Holman (1969) from Llano and Bexar counties, Texas, may be referable to L. berlandieri. Similar fossils reported for Denton County, Texas, are in the contact zone between L. blairi and L. berlandieri (Dixon, 2000), and may be referable to ­either species. SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates berlandieri is a member of the Novirana clade of North American ranid frogs, and it is classified within the Pantherana, a group that includes the leopard and gopher frogs and comprises the Lithobates pipiens complex. Mating calls within the Pantherana are highly complex in structure, and include ele­ments described as chuckles, grunts, and snores (Hillis and Wilcox, 2005). Mating calls are only produced during breeding choruses, whereas other ele­ments of the call are produced at other times of the year in the Pantherana. Rio Grande Leopard Frogs are more closely related phyloge­ne­tically to L. sphenocephalus, L. blairi, L. yavapaiensis, L. onca, and several Mexican species (the Scurrilirana, frogs that produce a chuckle-­like call) than they are to L. pipiens and L. chiricahuensis. Lithobates berlandieri hybridizes in nature with other members of the Leopard Frog complex such as L. sphenocephalus and L. blairi in central and west Texas (McAlister, 1962; Littlejohn and Oldham, 1968; Sage and Selander, 1979; Platz, 1981; Kocher and Sage, 1986). Contact hybrid zones may be rather narrow in extent (8 km in Texas) and stable through time, although hybridization can occur over a wider area (36–75 km) as demonstrated by biochemical analy­sis (Sage and Selander, 1979). Backcrossing into the parental populations is not extensive. Hybridization rates are low; for example, Platz (1981) recorded 5.8% hybrids at a contact zone between L. blairi and L. berlandieri in Texas. He further noted a significant change in the ratio of L. berlandieri to L. blairi through time (2 to 1 in 1969 and 19 to 1 in 1975), which suggested a rather dynamic interaction between the species. Differences in premating isolating mechanisms likely make hybrids unsuccessful in spreading into habitats occupied by the parental species. In laboratory crosses, ♂ L. berlandieri can produce successful larvae when crossed with ♀ L. palustris but not with ♀ L. montezumae (Mecham, 1969). Larvae of the ♂ L. berlandieri × ♀ L. palustris and ♂ L. berlandieri ×  ♀ L. blairi crosses developed macrocephaly and several tail fin abnormalities, but ­these dis­appeared with further development. Larvae resulting from crosses between L. berlandieri and L. forrei, L. magnaocularis, L. spectabilis, or L. sphenocephalus exhibited mild to severe hybrid inferiority (references in Hillis, 1988). Crosses between

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L. berlandieri and L. areolatus, L. chiricahuensis, L. megapoda, or L. pipiens ­were unsuccessful (Mecham, 1969; Cuellar, 1971; references in Hillis, 1988). ADULT HABITAT

This species is associated with a wide variety of mostly clear-­water aquatic habitats, including ponds, springs, streams, rivers, permanent pools in intermittent streams, temporary pools with extended hydroperiods, and stock tanks. Sharma et al. (2011) provide a detailed analy­sis of ­water quality associated with this species’ habitat in Big Bend National Park, Texas. Habitats should contain nearby retreat sites, such as permanent ­water, root systems, rock cracks, or burrows. Along rivers, this species is most often found sitting on mud banks and rarely on rocks, sand, or in the ­water (Jung et al., 2002). On the Rio Grande, they ­were observed most often in open habitats or near seepwillow (Baccharis salicifolia) and ­giant reeds, but much less often among willows, tamarisk, or mesquite. In the Chisos Mountains of Texas, Minton (1958) found this species at elevations 2–3 km; Franz et al., 1988; Smith et al., 2021). Phillips (1995) recorded 2 frogs in Georgia moving distances of 95–102 m from their breeding ponds, and Humphries and Sisson (2012) tracked dispersing adults 0.5–3.5 km (mean = 1.3 km) to summer refugia following breeding in North Carolina. In the latter study, frogs moved from 263 to 1,200 m in a single night (mean 743 m). Many frogs moved directly to their refugia in 1 night, but Humphries and Sisson (2012) noted a few frogs that took from 9 to 27 days before settling down in their summer refugia. At 4 sites in Florida, Gopher Frogs ­were found 141–3,402 m from the nearest breeding wetland (Smith et al., 2021). Gopher Frogs make use of Gopher Tortoise burrows, longleaf pine stump holes, and other retreats for temporary shelter as they migrate to and from breeding ponds (Bailey, 1989), and ­will return to the same refugium from 1 year to the next (Humphries and Sisson, 2012). Under­ground burrows are also crucial for dispersing juveniles, with survivorship greatly increased in areas with high numbers of burrows (Roznik and Johnson, 2009b). Precipitation over a period of days likely facilitates such movements. Juvenile dispersal occurs during the hot summer months and may not be correlated with rainfall during the humid nights (Greenberg, 2001b); some juveniles may be active even in fall and winter, however. Juveniles emigrate from breeding ponds nonrandomly ­toward fire-­maintained longleaf pine habitats rather than to habitats that are fire suppressed. Fire-­maintained habitats have more open-­canopy forest (ca. 18%; Phillips, 1995), fewer hardwood trees, smaller amounts of leaf litter, and larger amounts of wiregrass than fire-­suppressed habitats. They also contain more Gopher Tortoise and small mammal burrows than fire-­

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suppressed habitats, thus offering migratory Gopher Frogs shelter and permanent retreats. In Florida, dispersing juveniles moved to 691 m (mean 173 m) from the natal pond and sometimes used dirt roads as movement corridors (Roznik and Johnson, 2009a, 2009b). The mean distance between successive moves was 60.4 m. Long movements sometimes occur in a short period, and juveniles ­will use other ponds in the vicinity as stopovers during dispersal. Gopher Frogs are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). They likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males and females arrive si­mul­ta­neously at breeding ponds, but males arrive in greater numbers early in the breeding season. Although movements may occur over an extended time, most frogs move on only a few nights. Movements to breeding sites are correlated with rainfall, and most frogs arrive 1–5 hrs ­after dark or around sunrise (Bailey, 1990). Males remained longer at a breeding site in Alabama than females (males, mean 25 days; females, mean 9 days), although males spent up to 59 days and females 37 days at the pond (Bailey, 1990, 1991). Nearly half the males remained at the pond ≥30 days, but only 6% of the females did so. In Florida, mean residency for males was 14.6 days (range 1–78 days) and for females was 9.5 days (range 1–95 days) (Palis, 1998). Movements occur back and forth between the breeding site and terrestrial habitats throughout the breeding season, depending on rainfall (Humphries and Sisson, 2012), and ­these frogs usually emigrate in the same general direction as they arrived. Calling occurs from thick grass tussocks and sedges in shallow ­water, pondside debris, and from the bases of stumps, gum trees, and cypress trees. Males call at night, with peaks ­after dark and before dawn, and calls may occasionally be heard in the late after­noon. The call of L. capito is a deep snore lasting about 2 sec. Lannoo et al. (2018) reported a mean dominant frequency of 708.8 Hz, a mean call duration of 1.82 sec, a mean interpulse period of 0.045 sec, and a mean of 42.4 pulses per call at 15°C in Georgia. The call has good carry­ing capacity and may be heard 0.4 km distant (Wright and Wright, 1949). Gopher Frogs also call from underwater, in which case the call may not be audible to ­human observers >10 m away (Jensen et al., 1995). BREEDING SITES

Breeding occurs in temporary to semipermanent, mostly fishless ponds (Holbrook and Dorn, 2016, but see Phillips, 1995) dominated by short herbaceous vegetation (maiden-

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Lithobates capito 447

cane, panic grass, bluestem, yellow-­eye, pipewort, vari­ous rushes). Short woody vegetation often surrounds the pond margins, but ­there should be an open canopy throughout much of the pond. Depression wetlands range in size from 0.10 to 33.5 ha (Bailey, 1990; Dodd, 1992; Cash, 1994; Palis, 1997; Greenberg, 2001b; Greenberg and Tanner, 2005b). Other sites include sinkhole ponds, cypress ponds, cypress/gum ponds, pine savanna wetlands, Carolina Bays, ditches, and borrow pits. Ponds are usually shallow (80% of the area within 1 km of the home pond (Herrmann et al., 2005). In the ­Great Plains along the western extent of its natu­ral range, bullfrogs prob­ably followed forested riparian habitats along river floodplains into the prairies. ­Today in the West, however, bullfrogs are widely introduced into many types of habitats, from California rivers to isolated ­cattle tanks in Arizona deserts to prairie potholes throughout the upper ­Great Plains. They are found at densities of 9/km along the lower reaches of the Colorado River, primarily in areas with >50% of the riverbank in vegetation (Clarkson and DeVos, 1986). American Bullfrogs have occasionally been found in wells or caves (Lee, 1969a; Black, 1973a; Mount, 1975; Garton et al., 1993; Rimer and Briggler, 2010; Niemiller et al., 2016; Zigler et al., 2020; Camp and Jensen, 2021). ECOLOGY DURING NONBREEDING SEASON

Adults and juveniles presumably occupy the same general types of habitats, but ­there may be some degree of habitat partitioning among the life stages. For example, Goin (1943) reported that adult bullfrogs avoided the hyacinth-­covered ponds and streams of Florida, but that juveniles ­were common in this habitat in spring. Juveniles also hide ­under rocks and in crevices of dry wetlands during unfavorable weather conditions (Carlson, 2014b). American Bullfrogs are active usually from spring through the autumn in the North, depending on weather conditions

Dodd_Canada_int_5pgs_B3.indd 454

and latitude. They tend to emerge from overwintering sites when ­water temperatures reach 14°C and air temperatures reach 20–24°C (Willis et al., 1956; Treanor and Nicola, 1972; Wiese, 1985; Meshaka et al., 2015b). However, Meshaka et al. (2015b) observed activity in ­water in April when the air temperature was only ca. 6°C in Pennsylvania, although terrestrial activity did not begin ­until air temperatures reached 14°C. In the southern parts of their range, they are active all year, both day and night (e.g., George, 1940), or they become active sometimes during the midwinter depending on temperature. Lit­er­a­ture reports of first activity (summarized by Bury and Whelan, 1984; also see Willis et al., 1956; Smithberger and Swarth, 1993; Gibson and Sattler, 2020) include January along the Gulf Coast and in ­Virginia and California; February in Kansas, Florida, Missouri, and Texas; March in Colorado, Connecticut, Texas, Missouri, Illinois, and Ohio; April in Indiana, Mary­land, New York, and Pennsylvania; and May in Colorado (Wiese, 1985). Some of ­these reports prob­ably represent sampling biases, as bullfrogs are active year-­round in the South, even as far north as eastern North Carolina (Gaul and Mitchell, 2007). In the central and northern parts of their range, American Bullfrogs are active ­until the cold weather of autumn, for example, mid-­to late October in Illinois (Durham and Bennett, 1963), Ohio (Walker, 1946), Colorado (Wiese, 1985), and Connecticut (Klemens, 1993); late October to early November in Missouri and ­Virginia (Willis et al., 1956; Gibson and Sattler, 2020); and as late as December in Montana (Sepúlveda and Layhee, 2015). Gorham (1964) found juveniles ­under stones along a lake in October and around cold springs in November in New Brunswick. Adults usually begin winter dormancy well before the onset of freezing weather, whereas juveniles may be active ­until the first freezing temperatures of winter. Still, a few individuals may be observed in northern locations almost any time ­there is an unusual warm spell in winter. For example, Nussbaum et al. (1983) reported a massive ball of 186 semi-­torpid bullfrogs in an Oregon pond in February. Winterkill can be a major source of mortality. Lithobates catesbeianus are most active in the spring, and terrestrial activity is associated with night and rainfall (George, 1940; Raney, 1940; Gibbons and Bennett, 1974). This may be ­because adult bullfrogs are unable to tolerate the loss of much body ­water (Thorson and Svihla, 1943; Thorson, 1955), so they must remain close to ­water or move when humidity is high. ­Unless exposed directly to sunlight, larvae and adults tend to reflect the temperature of the ­water in which they are located (Brattstrom, 1963). They spend more time on land on warm rainy nights than on cold rainy nights (Currie and Bellis, 1969). On cool or windy nights, they tend to float lower in the ­water than they do on warm

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Lithobates catesbeianus 455

and still nights, perhaps to avoid unfavorable temperatures (Wiese, 1985). Basking has been reported in American Bullfrogs, and they tolerate high thermal temperatures (26–33°C, mean 30°C, CTmax = 38.2°C in California; Lillywhite, 1970). Postmetamorphs often sit exposed in sunlight near ponds with ­little indication of heat stress. Juveniles and adults have similar preferred body temperatures (roughly 30°C), although acclimation history affects preferences such that frogs acclimated at lower temperatures prefer cooler environmental temperatures (Lillywhite, 1971a). Temperature regulation is by behavioral means and is mediated by the hypothalamus. Juveniles and adults move back and forth between the pond shoreline and ­water in order to regulate their body temperatures, although juveniles also change postures more than adults to achieve thermal preferences. The heads of bullfrogs are particularly sensitive to heat. As they warm, the anterior hypothalamus directs mucous glands in the skin to increase mucous secretion, which modulates evaporative ­water loss as the temperature rises (Lillywhite, 1971b). The circulatory system also plays a vital role in maintaining skin hydration, perhaps more so than mucous secretions during basking (Lillywhite, 1975). Movements within and between seasons are sometimes extensive. An adult was recorded moving 1,219 m over a period of 19 days in New York, and another moved 380 m in 8 days (Ingram and Raney, 1943). In this same study, an individual was recaptured 1,600 m from where it was marked the previous year, with many other individuals moving >350 m from 1 season to the next. In Missouri, Willis et al. (1956) found that most bullfrogs remained within a wetland but that overland movement could occur to 1.25 km. In addition, many individuals also remained close (within 30 m) to where originally marked, both within and between seasons (Raney, 1940; Ingram and Raney, 1943; Wiese, 1985). In Washington, 1 bullfrog moved 2,794 m in 58 days (to 849 m in 1 week), and another moved 2,172 m (Rowe et al., 2021); movements averaged 156 m/week during midsummer. Other frogs stay in 1 general location for a season or more, then change locations, or they move back and forth between a series of locations (Durham and Bennett, 1963). Raney (1940) noted that bullfrogs inhabiting small wetlands and streams tended to remain at ­these locations during the course of the summer, although he also noted occasional long-­distance exceptions. In arid conditions, adults and juveniles may not leave the shoreline for more than a few meters, except when they are dispersing, due to a lack of suitable terrestrial habitats with favorably humid microenvironments. Movements may occur from 1 side of a large pond to the other (Willis et al., 1956). In a postbreeding study in

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Ontario, Currie and Bellis (1969) recorded a mean activity radius of 2.6 m (0.61–11.3 m) over a ca. 2 month observation period, with males having a slightly larger activity radius than females. When ponds ­were at a low density of bullfrogs, individual activity radii ­were larger than they ­were when bullfrog densities ­were high. Displaced bullfrogs home readily (McAtee, 1921; Schroeder and Baskett, 1965), moving distances of 205 m in 48 h (Ingram and Raney, 1943), although some animals move to other locations (Durham and Bennett, 1963). Based on a small number of recaptures, Raney (1940) initially thought the tendency to home was not well developed, but ­these results prob­ably reflected biases resulting from a short study season and a small sample size. In the Deep South, bullfrogs do not become dormant in the winter. In the North, overwintering occurs in ­water (Smith, 1934; Willis et al., 1956; Treanor and Nicola, 1972; Stinner et al., 1994; Minton, 2001; Chance, 2002; Collins, 1993; Sepúlveda and Layhee, 2015), although ­there is a report of dormancy ­under leaf litter on land throughout the winter (Bohnsack, 1952). Bullfrogs bury into the mud, and their cavities have been described as pits or cave-­like holes (in Bury and Whelan, 1984). They tend to overwinter in shallow areas of a pond and do not bury into the substrate; although such areas are colder than deeper ­waters, they have a higher dissolved oxygen content (Chance, 2002). Bullfrogs tend not to remain in 1 location but move around considerably ­under ­water, even ­under ice, despite the cold conditions (Friet, 1993; Stinner et al., 1994; Chance, 2002). Bullfrogs also ­will leave drying pools or stream channels and move to dormancy sites in springs, wells, and moist crevices in the ground, or to other wetlands if available. Juveniles may excavate single-­occupancy pits, which serve a dual function of concealment and retarding desiccation (Thrall, 1971). Juvenile bullfrogs may be quite active in the autumn in Montana. They generally remain within 6 m of the shore initially, but as the season progresses, they can be found at 15 m from the shoreline. Sepúlveda and Layhee (2015) found that juveniles moved greater distances in late summer and early autumn (4.5 m/day), but that as temperatures became colder, activity became more localized. For example, movements of 13 juveniles covered an area of 15,384 m2 during the active season but shrank to 130 m2 in the east cove of their study pond as the site froze over. Distances moved (median cumulative movement of 107.5 m) ­were not correlated with SUL. ­These authors suggested that the late season concentration of bullfrogs during cold northern winters might be used to eliminate them from areas where they had been introduced, such as their study site in Montana. During the active season, American Bullfrogs do not exhibit any characteristics of sleep be­hav­ior, and appear alert

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456  Ranidae

at all times despite intervals of rest (Hobson, 1966). Like most frogs that are active both day and night, American Bullfrogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are particularly sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open horizon above lakes and ponds (Hailman and Jaeger, 1974). Bullfrogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Male American Bullfrogs are highly territorial (Wiewandt, 1969; Emlen, 1977), and both male calling be­hav­ior and territorial defense may be influenced by hormone levels. Levels of androgen and luteinizing hormone generally increase at the start of the breeding season, and corticosterone peaks as the most intense period of chorusing begins. However, hormone levels fluctuate throughout the breeding season, even in nonterritorial males. Mendonça et al. (1985) suggested that stresses associated with territorial be­hav­ior and aggressive encounters might result in an inhibitory effect on androgen production. Thus, the effects of hormones on reproductive activity and be­hav­ior in American Bullfrogs are not as clear as they are in some other animals. ­There are a number of distinct types of calls in the American Bullfrog: mating (or advertisement), territorial (sex specific), territorial (produced by both sexes), release, warning, and distress (Capranica, 1968). As their names imply, each has a specific function, with their own call characteristics and sounds. Mating calls are ­those most familiar to ­people—­that is, the deep sonorous snore (vaarrhummm) of the male’s voice vibrating across a ­water body at dusk. Mating calls are composed of 3–15 rising and falling notes in rapid succession, during which the male’s paired vocal sacs are inflated. Individual notes last 0.6– 1.5 sec, with intervals between notes of 0.5–1.0 sec. Calls have a periodicity of 90–110/sec; low frequencies of 200– 300 cps rise to high frequencies of 1,400–1,500 cps. Capranica (1965, 1968) and Bee and Gerhardt (2001a) provide comprehensive summaries of the mating call characteristics. The male uses the advertisement call to attract females to mate, although it can be induced in laboratory settings at other times of the year. ­There is an inverse relationship between the size of the male and the resonance of his mating call. Thus, the call transmits information to the female of his potential as a mate (assuming larger males have greater fitness) as well as his location. In addition, male bullfrogs are able to discriminate the calls of familiar and unfamiliar males (Bee and Gerhardt, 2002), and thus become familiar with the location and characteristics of adjacent (and

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perhaps rival) males. Bee and Gerhardt (2001b) suggested such neighbor recognition could lead to reduced aggression among adjacent males, as males become habituated to a neighbor’s call. Bee (2001) could provide only partial support for this hypothesis. However, American Bullfrogs do exhibit some degree of stimulus-­specific habituation in regard to encounter calls, aggressive movements, and the tendency to approach a rival (Bee, 2003). Territorial calls are used by males and females to establish the bounds of their territories to members of their own sex. The male’s territorial call challenges nearby or rival males and asserts owner­ship of the territory, but calls are not used to assess the fighting ability of rivals as they are in other species (Bee, 2002). If the territorial call is unheeded by an approaching frog, an attack may follow (Wiewandt, 1969; Ryan, 1980; photo of wrestling in Howard, 1988b). As might be expected, most aggressive encounters occur early in the breeding season as territories are being established, and they are usually won by the older and larger male (Howard, 1978b). A female’s territorial call serves the same function. In both instances, however, the opposite sex does not respond by calling in response to the other’s territorial call, although they may move away from the area. A third call, termed the male/female territorial call by Capranica (1968), is a more generalized call issued by the winner of a contest or at the distant approach of a conspecific. Territorial males occur at regular intervals (­every 2–6 m) around the breeding pond, and readily defend their calling station. Wiewandt (1969) noted that the most responsive bullfrogs in his study defended 9–25 m of shoreline. According to Emlen (1976), centrally located territories are occupied by larger (older) males, with smaller males peripheral. Howard (1978b), however, disputed a size-­based distribution pattern to chorus structure, and noted that it is often difficult to define the spatial configuration of a chorus. Calling occurs during a discrete breeding season beginning in late spring and continuing through midsummer, depending on latitude. Bullfrogs are able to call both entirely on land and while partially submerged. Partial submergence allows the calls to resonate throughout the breeding pond (Boatwright-­Horowitz et al., 1999), and allows males to transmit information about their location and physical attributes farther than if calling occurred solely through the atmosphere. Males arrive first at the breeding ponds, and larger males usually arrive before smaller males. Calling begins when spring air temperatures are >21°C and ­water temperatures are >20°C (Fitch, 1956a; Oseen and Wassersug, 2002). However, Meshaka et al. (2015b) reported nocturnal calling at temperatures of 15–22°C in Pennsylvania when humidity was high. Bullfrogs prefer calling when wind speeds are low (Oseen and Wassersug, 2002).

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Lithobates catesbeianus 457

Choruses draw bullfrogs of both sexes ­toward a breeding pond (Emlen, 1976). At first, calling is sporadic and begins in the late after­noon, although calls may occur at any time during the day. As the season progresses, however, choruses (defined by Howard, 1978b, as a group of acoustically interacting males) become much more intense, with peak calling occurring ­after midnight (e.g., >12:00 in Michigan; 02:50–08:00 in Québec; 01:30–06:30 in South Carolina). Chorusing occurs all night, tapering off at dawn (Oseen and Wassersug, 2002; Mohr and Dorcas, 1999). In contrast, Cook et al. (2011) reported that most calling occurred from an hour or 2 ­after sunset till midnight on Cape Cod; the peak chorus time lasted 222 minutes. Chorusing occurs steadily throughout the breeding season but may be inhibited by low temperatures and high winds along the ­water’s surface. On Cape Cod, Cook et al. (2011) noted a peak chorusing time of 27 days in an 88 day calling season. A chorus begins with a few males calling, followed by a steadily increasing number of males calling ­until a veritable roar emanates from the breeding site. Males begin to drop out of the chorus ­after a short while, and calling decreases to only a few males. The chorus may be ­silent for several minutes before the cycle starts again. Peak calling occurs over a period of 3–5 nights within an individual chorus, but choruses may form and dissolve at dif­fer­ent locations around a breeding pond during the course of the breeding season. Emlen (1976) provides a diagram of chorus location on 10 dif­fer­ent eve­nings, illustrating the fluctuating position of American Bullfrog choruses around a single pond. Chorus shifts may occur in response to leech population buildup, changes in vegetation structure, or thermal changes as the season progresses. Not all bullfrog populations have shifting chorus locations, however. The positions of chorus locations also can change annually. Males sometimes move between chorusing aggregations. In contrast to Emlen’s (1976) assertion, the structure of bullfrog choruses in not analogous to lek formations in birds, inasmuch as chorus structure is not easily defined spatially, nor is it size based (Howard, 1978b). Females initiate mating by approaching and physically contacting the male. They may not accept the first male contacted, but instead may approach several males before choosing to mate, a pro­cess that may take several hours (Emlen, 1976). Sexual se­lection is weakly size related, with large females selecting large males, although smaller females are more generalist in their mate choice (Howard, 1988b). As might be expected, larger males tend to mate more often than smaller males, presumably in part ­because of their higher-­quality territories and physical superiority. Small males, however, may become satellite males, trying to intercept females attracted to the high-­quality territories of

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large males. Satellites remain in a crouched or “low” position within ca. 1 m of a territorial male, do not call, and avoid encounters with large males; their mating success is very low. A third group of opportunistic males may call, but they do not establish territories and readily flee when challenged by territorial males. ­These males tend to move around a ­great deal, avoiding aggressive encounters; they usually have intermediate mating success between territorial and satellite males. The number of nights a male ­will engage in territorial be­hav­ior and calling varies, and shifts by males among choruses do not affect the amount of time spent chorusing. Dominant males spend about 50% of their time chorusing (that is, on 26 nights of a 46 night breeding season in Ontario, with 15 nights in dominant tenure; Judge and Brooks, 2001), whereas many males spend much less time ­doing so. Males in better initial body condition tended to have longer dominant tenures at choruses and lost their body condition slowly, but had poorer body condition at the end of the season than males with shorter periods of dominant tenure. Thus, ­there appears to be an energy constraint to chorusing that affects the extent a male ­will remain in dominant tenure during the breeding season. Release calls are issued by males or unreceptive females in response to amplexus attempts by other males or would-be suitors. The sound is usually made ­after a short strug­gle, and the amplexed animal is quickly released. Release calls last 0.5–1.0 sec and are repeated ­every 1.5–2.0 sec. Calls are repeated at 60–85/sec at about 500 cps (Capranica, 1968). BREEDING SITES

Breeding sites are large, permanent bodies of ­water, such as lakes, oxbows, natu­ral and artificial ponds and reservoirs, and quiet ­waters of rivers, streams, and canals; bullfrogs also may occupy semipermanent wetlands with long hydroperiods (Eason and Fauth, 2001; Babbitt et al., 2003). In a typical example, most American Bullfrogs found in New Hampshire occurred in permanent ponds (95%), with only 5% found in temporary wetlands (Herrmann et al., 2005). In addition, they are frequently found in association with agricultural and urban settings, perhaps at reduced abundance (Picone, 2015), but much less so with silviculture (Surdick, 2005). Brodman (2009) noted considerable variation in annual occupancy of breeding sites over a 14 yr study in Indiana. In Missouri, American Bullfrogs occupied 55% of 210 ponds sampled over an 11 yr period (Drake et al., 2015). In northern Idaho, Lucid et al. (2020) found them at 23 of 433 (5%) potential sites. In the South, American Bullfrogs are associated with cypress, tupelo, buttonbush, willows, and grasses, although it is difficult to identify preferred habitats by vegetation alone (George, 1940). They readily breed in human-­created

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wetlands, including reclaimed surface mines (Myers and Klimstra, 1963; Turner and Fowler, 1981; Lacki et al., 1992; Ultsch et al., 1999; Anderson and Arruda, 2006), farm and stock ponds (Galois and Ouellet, 2005; Anderson and Arruda, 2006), quarries, golf course ponds (Boone et al., 2008; Scott et al., 2008; Mifsud and Mifsud, 2008), artificial urban wetlands (Neill, 1950a; Delis et al., 1996; Surdick, 2005), fish hatcheries, power-­line wetlands (Fortin et al., 2004b), constructed or restored wetlands (Briggler, 1998; Merovich and Howard, 2000; Pechmann et al., 2001; Foster et al., 2004; Brodman et al., 2006; Henning and Schirato, 2006; Palis, 2007; Shulse et al., 2010; Denton and Richter, 2013; Terrell et al., 2014a; Walls et al., 2014a; Drayer and Richter, 2016; Kross and Richter, 2016; Mitchell, 2016: Stiles et al., 2017a; Baecher et al., 2018; Drayer et al., 2020), and retention ponds (Foster et al., 2004; Surdick, 2005; Ostergaard et al., 2008; Birx-­Raybuck et al., 2010; McCarthy and Lathrop, 2011). Proximity to riparian and terrestrial habitats facilitates retention pond use. Large permanent ponds are particularly favorable for colonization. American Bullfrogs oviposit in shallow ­water (15–60 cm), preferably with substantial, uniformly distributed emergent and submerged vegetation. They tend to choose warmer sites early in the breeding season, but cooler sites as the ­water temperature increases as the season progresses (Howard, 1978b). Breeding sites are usually open canopied and relatively devoid of tree cover. REPRODUCTION

Breeding begins in early to late spring and occurs through the summer, depending on latitude. However, adults ­will become active much ­earlier than when calls are first heard. ­There also appears to be some confusion in the lit­er­a­ture as

Egg mass of Lithobates catesbeianus. Photo: Dana Drake

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to the breeding season, as calling is often equated with breeding; hence, some “breeding” dates in the lit­er­a­ture may not be accurate. For example, bullfrogs in east central Illinois first appear at breeding sites in late March to mid-­April, but eggs are not observed ­until late May (Durham and Bennett, 1963). In Québec, calling begins in May, but eggs are not observed ­until mid-­June (Bruneau and Magnin, 1980b). In Louisiana, calling occurs in February, but egg deposition occurs in mid-­April (George, 1940), and in Indiana, calling occurs from April to early July, although peak calling occurs from early May to mid-­June (Brodman and Kilmurry, 1998). Lit­er­a­ture rec­ords of calling dates include: February–­ August (Louisiana: George, 1940), March–­October (Florida: Carr, 1940a; Texas: Blair, 1961b), March–­June in Texas (Car­ter et al., 2018), April–­June (New Jersey: Ryan, 1980), April–­July (Indiana: Brodman and Kilmurry, 1998; ­Virginia: Mitchell, 1986; Gibson and Sattler, 2020), May–­July (Michigan: Emlen, 1976; North Carolina: Gaul and Mitchell, 2007; Ohio: Walker, 1946; Québec: Bruneau and Magnin, 1980b; Rhode Island: Raithel, 2019); May–­August (Mas­sa­chu­setts: Cook et al., 2011; Oregon: Nussbaum et al., 1983), June–­July (Minnesota: Oldfield and Moriarty, 1994; New York: Raney, 1940; Nova Scotia: Gilhen, 1984; Ohio: Varhegyi et al., 1998; Ontario: Piersol, 1913; Judge and Brooks, 2001; Pennsylvania: Meshaka and Morales, 2020; Québec: Lepage et al., 1997; South Dakota: Fischer, 1998; Wisconsin: Vogt, 1981); and July–­September (Colorado: Wiese, 1985). Parker (1937) also reported hearing calls of American Bullfrogs as late as the first week of September ­after a period of rain in western Tennessee. Egg deposition occurs from April to May in Alabama (Brown, 1956), May–­June in Tennessee (Gentry, 1955), June in Pennsylvania (Meshaka and Morales, 2020), June–­July in Kentucky (Viparina and Just, 1975), April–­July in Arizona (Dowe, 1979), and May–­August in Oregon (Brown, 1972); in Oregon, however, most eggs are oviposited from late June to mid-­July. The normal breeding season (i.e., with egg deposition) extends over a period of 1 to 2 months throughout most of the range of the American Bullfrog, despite differences in when it is initiated. However, ­there may be 2 peaks of egg laying within the breeding season, the first in spring to early summer and the second about 3 weeks ­later as large females get ready to deposit a second clutch (Bruggers and Jackson, 1974; Emlen, 1977; Dowe, 1979); Emlen noted that 5 of 73 bullfrogs followed during a breeding season in Michigan produced a second clutch. George (1940) suggested that choruses only lasted a few days ­after egg deposition in Louisiana, although occasional calls could be heard ­until August. Females ovulate over a very short period of time (one night), but ovulation within the population is asynchronous and occurs throughout the

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breeding season (Emlen, 1976). Thus, males may stay at the breeding pond throughout the breeding period, whereas females stay just long enough to mate and move back and forth to the main breeding pond. The sex ratio around a breeding pond is highly male skewed at any one time during the breeding season since females are receptive for only a very brief time and do not remain in close proximity to the highly territorial males. However, the sex ratio may be 1:1 for frogs sampled throughout the breeding season at multiple sites (Schroeder, 1966; Schroeder and Baskett, 1968; Suhre, 2010). Still, Wiese (1985) found a sex ratio of 1.3 females per male in a Colorado population. Amplexus is axillo-­pectoral, occurs in ­water, and lasts from 17 to 155 min (mean 49 min) in Michigan (Howard, 1978b). During oviposition, the female extends her legs backward, downward, and laterally; her body arches concavely, her head is ­under ­water, and her cloaca is arched upward (illustration in Aronson, 1943a). The male appears to stimulate the female using his hind limbs to stroke her body, at which time the female’s abdominal muscles begin to contract and the first eggs are extruded. The male fertilizes the eggs, then uses his rear legs to spread them out in a surface film. The duration of amplexus (ca. 2 hrs in Aronson’s observations) is positively correlated with female body size, but not with male body size. Males mate with more than 1 female during a breeding season, although Howard (1988b) found that 48% of the males in his population ­were unsuccessful at mating. Females, however, all mated at least once, and some twice, over a 3 yr period. Mating success is low in the first year of maturity (11% for males, 10% for females), but increases from 55 to 100% for 2–5 yr old males. Females >1 yr in age had a 100% chance of mating successfully (Howard, 1988b). Eggs are normally deposited within the territory of the fertilizing male (Ryan, 1980). The number of eggs produced varies with female size, such that larger and older females produce far more eggs than younger and smaller females (Collins, 1975; Howard, 1978a; McAuliffe, 1978; Bruneau and Magnin, 1980b). For example, Howard (1988b) recorded Michigan females as producing means of 2,007 eggs in yr 1, 3,372 in yr 2, 7,228 in yr 3, 10,238 in yr 4, and 11,147 in yr 5; the maximum number was ca. 20,000 eggs. Female body weight is correlated with clutch weight and the number of eggs per clutch, but not with female SUL (Woodward, 1987a). In Québec, the number of eggs ranged from 3,826 to 23,540 (Bruneau and Magnin, 1980b). In Nebraska, a 128 mm SUL female deposited 16,640 eggs and a 179 mm SUL female contained 47,480 eggs (McAuliffe, 1978), whereas a 161 mm SUL female deposited 29,281 eggs in Pennsylvania (Rep et al., 2015). George (1940) estimated clutch sizes at 8,000–15,000 (mean 12,000) eggs in Louisiana; Trauth et al.

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(1990) recorded 12,756–43,073 (mean 22,944) eggs per female in Arkansas; Ryan (1980) recorded a mean of 7,360 eggs/mass in New Jersey; Meshaka et al. (2015b) estimated a mean of 15,237 eggs (range 4,742–35,800) in Pennsylvania; 11,585–21,510 eggs ­were counted in Nova Scotia (Gilhen, 1984); Brown (1972) counted 16,491 eggs in a single Oregon clutch; and Woodward (1987a) recorded a mean of 11,126 eggs per clutch in New Mexico. Numbers >20,000 or so could represent multiple clutches. Willis et al. (1956) provide detailed descriptions of the eggs, oviducts, and ovaries of bullfrogs throughout the reproductive cycle. Males generally are able to fertilize more eggs as they grow larger and older, from 2,732 in yr 1 to 19,346 in yr 5 (Howard, 1988b). Large females also produce larger eggs (1.58 mm in dia­meter) than smaller females in their first clutch of the year, but not in the second clutch (1.48 mm in dia­meter). Female size and total fecundity are not correlated, although such a relationship exists if first and second clutches are distinguished (Howard, 1978a). The second clutch of small females, however, also had smaller eggs than their first clutch of the season. Eggs fertilized by larger males also may have higher hatching success in some years than eggs fertilized by small males. American Bullfrogs have a rather narrow range of temperatures in which normal embryo development takes place. Eggs do not hatch or develop in ­water 32°C, with death at 33–36°C (Moore, 1942a; but see Dowe, 1979, where eggs developed at 33°C). The egg mass temperature may be slightly lower than the surrounding ­water temperature, thus providing a mea­sure of protection from high temperatures (Ryan, 1978). LARVAL ECOLOGY

Larvae are considered primarily herbivorous, grazing on algae and plant material. Larvae ­were considered detritus feeders by Thrall (1972), whereas ­others have reported them grazing on algae and bacteria as well as detritus (Brown, 1972; Kupferberg, 1997a), or acting as suspension feeders on phytoplankton (Seale, 1980). Thrall (1972) found no evidence of morphological or physiological specializations for herbivory. In contrast, Pryor (2003, 2008) described the gut as containing bacteria (Edwardsiella tarda, Clostridium sp.), ciliates, and nematodes, particularly the nematode Gyrinicola batrachiensis. ­Under laboratory conditions, the presence of the nematode aided in fermentation, except for cellulose, and it is considered a mutualistic symbiont of

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460  Ranidae

Tadpole of Lithobates catesbeianus. Photo: David Dennis

Tadpole of Florida-­Georgia “Lithobates catesbeianus.” Photo: Ronn Altig

bullfrog tadpoles (Pryor, 2003; Pryor and Bjorndal, 2005). Munz (1920) recorded diatoms, mud, a number of species of algae, and epidermis in tadpole guts. In Oregon, American Bullfrogs tend to feed on algae in proportion to its availability within the pond. However, ­there may be among-­habitat differences in the percentage of species consumed, a diet that changes during the season. Larvae may eat species in proportion to their occurrence in 1 pond, but not in adjacent ponds. In other situations, some algae (e.g., Spirogyra, Mougeota) may be found in much greater proportions in guts than in pond ­water, suggesting preferential se­lection (Brown, 1972). Types of algae in the diet of bullfrogs include members of the Oedogoniales, Bacillariophycea, Desmidacea, and Zygnematales. Light, of course, influences primary production of algae, periphyton, and phytoplankton. As a result, one might expect that tadpole growth would be enhanced in open-­canopied wetlands that receive large amounts of sunlight. Light does have strong effects on resource quality, but this does not predict bullfrog larval growth or development. Instead, it is the

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nutrients that fuel the algal growth, and thus accounts for the larger size of tadpoles and the faster rate of tadpole development in high nutrient mesocosms (Rowland et al, 2016). Larval American Bullfrogs are opportunistically carnivorous, feeding on the dead bodies of other animals, including larval conspecifics. They sometimes consume the eggs and hatchlings of other ranid species (Ehrlich, 1979), despite the inability of Funk­houser (1976) to induce them to take animal material. Disagreements as to the categorization of the diet of bullfrog larvae may reflect regional differences among populations of bullfrogs or in food availability. ­Under laboratory conditions, larvae are coprophagous, a situation that enhances growth rates (Steinwascher, 1978a; Pryor, 2003). In the North, larvae grow from around 19–26 mm body length in midsummer to 32 mm by autumn of the first year (Willis et al., 1956; Bruggers and Jackson, 1974), to 40–52 mm by the second year, and to 53–56 mm ­after the third winter (in Québec: Bruneau and Magnin, 1980a). Larval growth rates are dependent on temperature, oxygen levels, larval density, food resource amount and availability, and possibly extent of sedimentation (George, 1940; Raney and Ingram, 1941; Licht, 1967; Brown, 1972; Bruggers and Jackson, 1974; Corse and Metter, 1980). ­These latter authors found that supplemental feeding greatly increased growth rates and size at metamorphosis and decreased the length of the larval period. Larvae do not grow in the winter, but they ­will continue to feed at much reduced levels. For example, nearly all growth occurs from May to November in Oregon (Brown, 1972). Larger larvae tend to choose ­water temperatures of 24–30°C (Brattstrom, 1962, 1963; Lucas and Reynolds, 1967), but preferences and developmental rates change with previous thermal history and stage of development (Hutchison and Hill, 1978; Crawshaw et al., 1992). The length of the photoperiod does not affect tadpole temperature preferences (Smith, 1999). In terms of other thermal effects, McCallum et al. (2020) found that swimming activity declines with temperature and also decreases as food resources increase. Swimming is more pronounced in low feeding groups ­under laboratory conditions, but feeding be­hav­ior does not increase significantly as temperature increases. As food resources decrease, tadpoles also tend to scrape the substrate more as they search for food. In short, hungry, warm tadpoles scrape the substrate more than satiated, cool tadpoles. In simulations of sudden ambient temperature decreases such as occur in autumn, McCallum et al. (2020) found that tadpoles are capable of altering their foraging be­hav­ior to compensate for decreased metabolism during the stress of suddenly declining ­water temperatures. As they approach metamorphosis, premetamorphic larvae move to the warmest part of the pond prior to leaving. In Michigan, small bullfrog tadpoles tended to prefer warmer, more open habitats in medium-­depth ­water, whereas larger

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tadpoles prefer cooler, deeper ­water and show no preferences regarding the extent of cover, at least ­under laboratory conditions (Smith, 1999). ­These results seem ­counter to other observations obtained from field observations of temperature and habitat use. During the summer, tadpoles may be found in both deep and shallow areas of ponds and lakes, a habitat choice that may result from dif­fer­ent predation pressures in shallow ­water habitats; that is, if predation pressure is high, larvae move to deeper ­waters. Inasmuch as deep ­water may be anoxic or severely hypoxic, bullfrog larvae switch from facultative air breathing to obligate air breathing in deep-­water lakes and ponds (Ultsch et al., 1999). However, Nie et al. (1999) suggested that diurnal and annual shifts in habitat use between deep and shallow ­water is driven by temperature se­lection rather than predation pressure. In any case, larger larvae tend to choose more structured habitats than smaller larvae, but ­there is a ­great deal of individual variation in habitat use and activity (Smith, 1999; Smith and Doupnik, 2005). American Bullfrog larvae often aggregate in large numbers, but ­these aggregations do not appear to have a social context as they do in some other ranids (Wassersug, 1973). Bullfrog larvae are more active in groups when in proximity to predators than are solitary tadpoles (Smith and Awan, 2009); this suggests that presence in larval groups reduces individual predation threat. Individual bullfrog tadpoles may have dif­fer­ent personalities, particularly as regards boldness and exploratory be­hav­ior, but not in terms of activity levels (Carlson and Langkilde, 2013). Observations on exploratory be­hav­ior ­were repeatable over multiple ­trials, but for boldness only marginally so, perhaps ­because of the small sample size tested. Bullfrogs are able to use plane-­polarized light in spatial orientation (Auburn and Taylor, 1979) as well as using extraocular photoreceptors for sun-­compass orientation (Justis and Taylor, 1976), both of which help in learning the physiography of the pond and in setting biological clocks. In this way, they are able to ascertain the location of shorelines, an impor­tant consideration for escape to deeper ­waters and for foraging and thermoregulation in shallower ­water. Larval bullfrogs are photopositive ­toward the color green, which may help them associate with green plants which offer a source of both food and cover (Jaeger and Hailman, 1976). Generalizations about the length of the larval period of this wide-­ranging species are misleading, as American Bullfrogs occur from the subtropics to the northern hardwood forest. Much more research is available on northern bullfrogs than southern bullfrogs, and statements on generalized life history characteristics are often based on northern populations. In addition, lit­er­a­ture rec­ords are sometimes unclear in their use of the term “year.” Thus, “transformation occurs in 2 years” could mean ­after 2 calendar years (i.e., in the third summer of life) or in the second season (i.e., in the second summer of life).

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American Bullfrogs overwinter 1 or 2 years prior to metamorphosis (i.e., metamorphosis in their second or third summer) throughout much of their range but may require 3 overwintering periods in Canada (Bruneau and Magnin, 1980a) and New Hampshire (Oliver and Bailey, 1939). In some areas, however, larvae do not overwinter. In Louisiana and Oregon, metamorphosis can occur 4 months ­after hatching (George, 1940; Cook et al., 2013); in Arizona, transformation occurs ­after 3 months (Dowe, 1979); Seale (1980) suggested transformation at 70%) at concentrations >30 mg/L (Hall, 1990). The insecticide fenitrothion kills or leaves bullfrog larvae para­lyzed at exposures of 4.0 and 8.0 ppm; larvae are unresponsive to touch at exposures as ­little as 0.5 ppm, and they do not recover (Berrill et al., 1994; Berrill et al., 1997). Azinphos-­ methyl (Guthion®) kills bullfrog tadpoles at 1.8 kg/ha in field applications (Mulla, 1962), but Meyer (1965) found no effect at 1.0 mg active ingredients per liter. TFM (a lampricide) has an LC50(96 hrs) of 3.55 mg/L (Chandler and Marking, 1975). The now banned DDT caused both mortality and delayed tail regeneration in American Bullfrog larvae (Weis, 1975). Other insecticides known to kill bullfrog larvae are endrin, heptachlor, dieldrin, aldrin, toxaphene, thiodan, Bayer 38920, trithion, and GC-3582 (Mulla, 1962, 1963). Fenthion at 5 mg/L for an exposure of 96 hrs had no effect on bullfrog larvae, but mallard ducklings that fed on the tadpoles died (Hall and Kolbe, 1980). Some insecticides also have sublethal effects beyond lethal toxicity. For example, the LC50(16 day) for malathion is 1.5 mg/L, which suggests that it is not lethal to American Bullfrogs ­under field conditions (Relyea, 2004b). However, concentrations of 0.1–1 mg/L can reduce activity by 19–27% (Relyea and Edwards, 2010; Hanlon and Relyea, 2013). Unlike Green Frogs, ­there are no additive effects between predator presence and malathion effects, and ­there was no effect on larval survival. Likewise, carbaryl at the same concentrations also significantly reduces activity, likely making larvae more susceptible to predation (Relyea and Edwards, 2010). Interestingly, parasite species richness and diversity are lower in American Bullfrogs from wetlands where parasites have been sprayed than in insecticide-­free wetlands (King et al., 2010). The insecticide endosulfan is highly toxic to American Bullfrogs even at low concentrations, with an LC50(4 day) of 1.3 ppb (Jones et al., 2009). In contrast, endosulfan had no significant effect on larval activity except in the presence of

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potential aquatic predators. In the presence of 10 ppb endosulfan and predators, survival was reduced initially in experimental ­trials, but ­there was no interaction between the 2 ­factors (Hanlon and Relyea, 2013). As time went on, an interaction became apparent as endosulfan reduced tadpole survival to 0% when predators ­were absent. ­These results suggest that predators have a greater influence on activity than the low concentrations of endosulfan used in the experiment. American Bullfrogs do not avoid areas where herbicides have been sprayed, particularly, diquat dibromide (Reward®), glyphosate (Roundup®), and chelated copper that acts as an algaecide (Picone, 2015). The herbicide acetochlor appears to affect thyroid hormone gene expression, and therefore possibly brain function, in American Bullfrog larvae. However, no effects on metamorphosis or escape be­hav­ior could be detected (Helbing et al., 2006). The herbicide hexazinone has no lethal effects on American Bullfrog embryos or tadpoles, although they may not respond to stimuli (Berrill et al., 1994, 1997). However, the herbicide triclopyr kills newly hatched tadpoles at 2.4 and 4.8 ppm (Berrill et al., 1994, 1997). Atrazine ­causes decreased hatching and survivorship (LC50 = 0.41 mg/L; Birge et al., 1980) and is lethal to larvae at 200 mg/L (Boschulte, 1993). Atrazine also has indirect effects on larvae at a much lower dosage, particularly on larval biomass when other grazers are pre­sent (DeNoyelles et al., 1989). The LC50 for the herbicide Roundup® is 2.1–2.2 mg a.e./L at low and medium tadpole densities (Jones et al., 2011). Increasing concentrations of the herbicide cause a sharp decrease in tadpole survival associated with an increase in periphyton abundance. Increasing concentrations of glyphosate (1–3 mg a.e./L) decrease larval survivorship, but adding predators had ­little interactive effect on survival (Relyea, 2018). As might be expected, increasing tadpole density ­causes decreases in tadpole growth, but this also makes the herbicide significantly more lethal, with an LC50 of only 1.6 mg a.e./L. The American Bullfrog appears resistant to the herbicide paraquat, perhaps due to stress-­ induced increases in antioxidant enzyme activity (Jones et al., 2010). American Bullfrog embryos and larvae are sensitive to toxicity from: carbon tetrachloride (LC50 = 0.90 mg/L), methylene chloride (LC50 = 17.78 mg/L), nitrilotriacetic acid (NTA) (LC50 = 113.4mg/L), and phenols (LC50 = 0.23 mg/L) (Birge et al., 1980), as well as 2-­chloroethanol, hexachloroethane, pentachlorophenol, permethrin, and 2,2,2-­trichloroethanol (Thurston et al., 1985); other chemical geometric mean LC50 values include: acridine (1.24 mg/L), CCl4 (1.26 mg/L), methylene chloride (22.9 mg/L), NTA (63.8 mg/L), and

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B-­naphtal (18.1 mg/L) (Westerman et al., 2003a). ­Under natu­ral conditions, American Bullfrogs do not selectively avoid sites contaminated by polychlorinated biphenyls (PCBs) (Gibbs et al., 2017). American Bullfrog tadpoles experience a suite of adverse behavioral and morphological changes when exposed to crude oil (McGrath and Alexander, 1979). They ­will float on the ­water’s surface, regardless of oil concentration, and they swell, causing bulges to appear on the lateral body surfaces. Tadpoles become lethargic and are unable to dive. Larvae exposed to high oil concentrations rapidly swim at the ­water’s surface with their heads above ­water while vigorously fanning their tails. ­After a short period of time, the tadpoles collapse back ­under the ­water’s surface. The eyes become bloodshot, indicating hemorrhage, and the forepart of the body takes on an unusual heart shape when viewed dorsally, due to grossly inflated lungs. Upon dissection, oil is common in the digestive tract, and the liver appears bright yellow (“fatty liver”) due to the presence of oil droplets. Later-­stage tadpoles are more sensitive to oil than early stage larvae. Taken together, ­these results suggest that crude oil in the environment is detrimental to American Bullfrog larvae. Per-­and polyfluoroalkyl substances (PFAS—ex. PFOS, PFOA, PFHxS) are widely used in a variety of ­house­hold applications. The mean LC50(96 hr) for PFOS was 99 mg/L for American Bullfrogs, whereas for PFOA it was ca. 1038 mg/L (Flynn et al., 2019; Tornabene et al., 2021). The specific chemical used and ­whether larvae are exposed chronically or acutely also influences the effects of exposure to PFAS. For example, chronic exposure to PFOS more adversely affects larval SUL, mass, and developmental stage than PFOA, but PFOA had more of an effect on SUL from a single exposure (Flynn et al., 2019). Combining ­these substances ­causes an additive effect. Survivorship declines with increasing concentrations of PFHxS, although American Bullfrogs ­were less sensitive to this chemical than to PFOS. UV light. American Bullfrog eggs have high levels of hatching success ­under field conditions of incident light, incident light with UVB light blocked, and even with artificially enhanced UVB light (Crump et al., 1999). Significant mortality occurs ­after 10 min exposure at high levels of UVB (>936 mJ/cm2) (Licht, 2003). At 300 nm, 40% of UVB is absorbed by the jelly capsule surrounding the egg. Salinity. In experimental ­trials, larval L. catesbeianus had 100% survival at salinities ≤ 5 ppt. However, no larvae survived at salinities of 14 or 16 ppt over a 72 hr period (Brown and Walls, 2013). Larval survival was ca. >70% at 10 ppt at 70 hr, but was declining rapidly by 72 hr.

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COMMERCIAL USE

American Bullfrogs are one of the most heavi­ly exploited amphibians in the world and have been so for more than a ­century. The primary purpose of commercial use has been for frog’s legs for ­human consumption. They have also been used extensively in science teaching labs and for medical research (Culley, 1973). Although frog “gigging” continues to supply frog legs for local consumption, much of ­today’s commercial trade results from vast “frog farms,” both in the United States (Arkansas, Louisiana, Florida, Hawai’i) and in numerous other countries. For example, Rana Ranch Commercial Bullfrogs in Idaho ships frogs to 43 states for not only biological research, but “to stock lakes, wetlands, ornamental koi ponds, ­water gardens, and golf course ­water features” (Capital Press, Salem, Oregon, July 12, 2019: https://­www​.­capitalpress.com​/nation​_world​/profit​/western​ -­innovator-­frog-­ranch-­keeps-­owners-­hopping/article​ _207a1dd0​-­3c7d​-­11e9​-­8bea​-­eb888f2b1dfe​.­html; Accessed January 29, 2020). Attempts at frog farming began in the United States and Canada before 1900, and perhaps ­earlier than 1888 in Ontario (Meehan and Andrews, 1908; Priddy and Culley, 1971; Dodd and Jennings, 2021). For example, Anonymous (1899) reported on a small frog farm in New York, and I have brochures or references for frog farms in Arkansas, California, Hawai’i, Illinois, Louisiana, Mas­sa­chu­setts, New Hampshire, New York, North Carolina, Ontario, and Texas, among other locations; doubtless many ­others existed throughout the US and Canada. Chamberlain (1897) noted that a frog farm in the Trent River Basin, Ontario, had been in operation for “about 20 years.” However, frog “farms” have often operated more like a vast network of persons collecting frogs and transporting them to a central location rather than true closed-­cycle operations. Since the late 1800s, American Bullfrogs have been recognized by private, state, and federal agencies as one of the primary species to fulfill the demand for frog’s legs (Meehan, 1906, 1908a, 1908b; Dyche, 1914; Louisiana Department of Conservation, 1931; Viosca, 1931; Stoutamire, 1932; Hannaca, 1933; AFCC, 1936; Baker, 1942; Brashears and Brashears, 1950; Broel, 1950; Florida Department of Agriculture, 1952; Brown, 1953; USDI, 1965; also see Storer, 1933; for an opposing view, see Schmidt, 1946). ­These publications supply a wealth of information on the life history of L. catesbeianus in addition to their marketing. State and federal publications also provide evidence of the massive dispersal of frogs throughout the United States and elsewhere. The earliest trade statistics ­were supplied by Chamberlain (1897), who reported an annual catch of about

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1 million frogs worth $50,000. Anonymous (1892) noted that shipments of 30,000–40,000 at a time ­were sent to Vienna, Austria, and that New Yorkers consumed 60,000 pounds/yr at 0.30¢ per pound. In Texas, the annual consumption was estimated at 300,000 pounds (Baker, 1942). Storer (1933) noted that ­there was already a plentiful supply of American Bullfrogs in California, and that attempts to start frog farms in that state began in 1898, based on a supply of imported bullfrogs “from somewhere in the eastern States.” American Bullfrogs ­were known from a frog farm in Contra Costa County, California, in 1896; they received imported stock from Mary­land and Florida (Heard, 1904; Jennings and Hayes, 1985). Frog farms in New York, North Carolina, and Florida imported their breeding stock from Louisiana (Lucas, 1965; Dodd and Jennings, 2021). Undoubtedly many frogs escaped and bred with local bullfrog populations. More than 5,000 live American Bullfrogs ­were shipped to Japan in the 1920s to create frog farms, although Japa­nese frog populations ­were subsequently seriously depleted by DDT. In 1945, 300,000 pounds of frog’s legs, worth >$100,000, ­were exported to Cuba, even though L. catesbeianus had been released in Cuba in 1915 and had spread throughout the island (Martinez, 1948). Broel (1950) lists American Bullfrogs as being shipped to China, France, Germany, Hungary, Italy, the Philippines, and South Amer­i­ca ­either as food or as stock for frog farms. The value of the frog industry has been substantial, even before the advent of commercial farms (see, for example, discussion in Dundee and Rossman, 1989). In 1908, for example, 113,636 kg of frog’s legs (worth $42,000) ­were reported taken in the United States. Canadians consumed CAN $200,000 worth of frog’s legs in Montreal alone in 1911, with CAN $100,000 worth ­going to the “country ­people” of Québec (Sibley, 1912). In 1928, 325,245 kg of bullfrogs (worth $107,331) ­were harvested from Louisiana, and ­these figures increased to 2.75 million pounds, worth $650,000, in 1936 (George, 1940). Some of the figures, however, include other ranid species, particularly L. pipiens. By the early 1970s, 9 million ranid frogs ­were harvested annually (360 tons), nearly all from wild-­caught stock. Major suppliers ­were in Wisconsin, Minnesota, and Vermont. Commercial take in the Midwest, in par­tic­u­lar, led to regional declines and to calls to enact conservation mea­sures (Gibbs et al., 1971). Concern for wild populations had been expressed much ­earlier, however. Chamberlain (1897) noted that bullfrog populations near areas of market and transport ­were decimated by un­regu­la­ted harvest. Meehan and Andrews (1908) even stated that Northern Leopard Frogs might be the best species to use in a frog farm, due to “the

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Lithobates catesbeianus 475

natu­ral supply [of bullfrogs] being apparently doomed to exhaustion.” The primary frog markets ­were in New York, Chicago, St. Louis, San Francisco, Boston, Philadelphia, Washington, and New Orleans. Pennsylvania established the first major state-­led commercial frog farming effort in 1899, but many prob­lems ­were encountered, and the attempt was not successful (Meehan and Andrews, 1908). By the 1930s, interest in commercial frog farming appeared to gain strength, in part due to the state of Louisiana’s interest in aquaculture, which had been initiated around 1917 (Viosca, 1931, 1934). Louisiana State University began extensive research into bullfrog culture in the late 1960s (Priddy and Culley, 1971). Successful frog culture was carried out in Arkansas and Louisiana by the early 1970s. ­Today, American Bullfrogs are highly sought ­after for commercial farms, and they have been introduced throughout the world, often with known or suspected deleterious effects on native fauna (Orchard, 1999; Mazzoni et al., 2003; Giovanelli et al., 2008; Wang and Li, 2009). Commercial bullfrog farming is still promoted by the UN/FAO (http://­www​.­fao​.­org​/­fishery​/­culturedspecies​/­Rana​_­catesbeiana). STATUS AND CONSERVATION

Despite a long history of exploitation, American Bullfrogs remain a common and widely distributed species. Griffis-­ Kyle et al. (2018) considered this species as least vulnerable throughout the Desert Southwest. Most surveys indicate populations that are stable or increasing, even when surveys of par­tic­u­lar areas ­were conducted de­cades apart (e.g., Christiansen, 1981; Busby and Parmelee, 1996; Weller and Green, 1997; Mierzwa, 1998; Mossman et al., 1998;

Habitat of Lithobates catesbeianus. Eastern North Amer­i­ca. Photo: John Bunnell

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Habitat of Lithobates catesbeianus. Desert Southwest. Photo: C.K. Dodd, Jr.

Brodman et al., 2002; Florey and Mullin, 2005). Still, Villena et al. (2016) suggested populations in the South ­were declining based on probability of occurrence through time. On the eastern seaboard, populations ­were considered increasing in 4 states (Delaware, New Jersey, ­Virginia, West ­Virginia) based on 7 yrs of data using occupancy modeling (Weir et al., 2009). Throughout many areas, introductions from a variety of sources (but principally by state wildlife and fisheries agencies) have resulted in expanding populations (Lannoo et al., 1994; Lannoo, 1996; Christiansen, 1998). Still, habitat loss to urbanization, agriculture, silviculture, transportation corridors, wetlands drainage, and other ­causes is the primary threat to most bullfrog populations. For example, American Bullfrogs dis­appeared around Baton Rouge in the 1930s as ponds and wetlands ­were drained (George, 1940); wetland modification leading to premature or unusual pond drying contributed to population declines. High levels of toxic agricultural pesticides are associated with smaller sizes, lower body masses, shorter tibia lengths, smaller sizes of the tympanum, and younger ages than conspecifics found in nonagricultural areas (Spear et al., 2009). In addition, agricultural chemicals can alter physiological activity and compromise immune function, and ­these in turn can be mediated through parasite activity (Marcogliese et al., 2009). ­These data suggest that a soup of agricultural chemicals ­causes decreased longevity and growth rates in impacted populations, even if populations are not outright eliminated. The presence of ­cattle in breeding ponds also reduces the abundance of postmetamorphic American Bullfrogs (Burton et al., 2009). In Ontario, evidence suggests that some populations are declining. Hecnar (1997) found American Bullfrogs in only 2.9% (5 of 174) of ponds in southern Ontario and noted declines or extirpations in areas where they ­were formerly

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abundant. Shirose and Brooks (1995a, 1997) also noted unstable population structures and variable age-­specific mortality rates in Algonquian Provincial Park, suggesting increased mortality prior to 1985 and from 1987 to 1991. Declines have been attributed to the effects of otters, overharvest, shoreline modification, and the use of pesticides (Shirose and Brooks, 1997; Weller and Green, 1997). Declines elsewhere have been reported for Wisconsin (Casper, 1998), and the species is listed as of Special Concern in Minnesota (Oldfield and Moriarty, 1994). Bullfrogs may be found in forested wetlands and then dis­appear following clearcutting. According to some studies, initial declines are followed by eventual recolonization (e.g., Clawson et al., 1997). However, it is often difficult to assess the impacts of clearcutting, as follow-up studies usually are of short duration, and they often use presence rather than abundance or demographic data to evaluate the effects of habitat disturbance. This is a poor metric to use, since it tends to ignore the functional context of a species, opting instead for mere occurrence. In studies on the effects of silviculture on bullfrogs, it is sometimes impossible to know ­whether 1 frog is involved or 10,000. However, American Bullfrogs are occasionally found in open habitats created during forestry operations, such as in canopy gaps and along skidder trails (Cromer et al., 2002). They ­will colonize artificial ponds, but Monello and Wright (1999) found that they ­were not particularly successful at ­doing so in northern Idaho. Road traffic undoubtedly takes a heavy toll on populations (e.g., Cunnington et al., 2014), and ­little is understood of its long-­term effects. In Indiana, bullfrogs ­were the most common species found as roadkill over a 17 month study, with 1,671 recorded (Glista et al., 2008), whereas 1,345 bullfrogs ­were killed on 3.6 km of the Long Point Causeway in Ontario over a 4 yr period (Ashley and Robinson, 1996). In the latter study, mortality was seasonally bimodal, as frogs immigrated in spring and emigrated in the autumn. On impervious roads, L. catesbeianus populations are negatively impacted at small local scales, but not over a more widespread regional scale (Marsh et al., 2017). As with other amphibians, bullfrogs are frequently surveyed using road-­ based call surveys (Burton et al., 2006). Acute artificial lighting alters calling be­hav­ior by decreasing the number of males calling in a chorus and decreasing the intensity of calling (Hall, 2016). Vargas-­Salinas et al. (2014) found that American Bullfrogs, a species with a low call peak frequency, called more often when traffic noise was low.

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In addition, American Bullfrog populations undoubtedly ­ ere decimated or extirpated by the un­regu­la­ted harvest for w frog’s legs throughout vast areas of North Amer­i­ca. Schroeder and Baskett (1968), for example, noted an absence of large individuals in areas heavi­ly harvested in comparison with areas not harvested. It is clear that many authors ­were concerned about the effects of the trade, although no data are available on specific populations. Most states and provinces regulate the take of bullfrogs ­today, although ­there are still few data on the population effects of harvest. Despite a variety of potential threats, however, the American Bullfrog remains one of the most widespread and common of North American amphibians, and it has become a considerable nuisance throughout the western states and provinces. Bullfrog populations may be transient, depending upon local conditions. Hecnar and M’Closkey (1996a) noted a small regional decline among ponds surveyed in Ontario over a 3 yr period. Rather than indicate concern, such changes may be indicative of normal population turnover, especially near the northern limit of the species’ range. The US Fish and Wildlife Ser­vice developed a Habitat Suitability Index (HSI) model for this species for use in habitat evaluation procedures on federal lands (Graves and Anderson, 1987). Such models incorporate lit­er­a­ture reports of habitat requirements, in this case employing 11 variables, to determine relationships between habitat variables, model components (“life requisites”), and HSI values that can be used to ascertain ­whether a habitat supports American Bullfrogs. American Bullfrogs usually are sampled through visual encounter surveys of larvae or adults, or by detecting them using automated frog call recorders. In France, however, researchers have developed the ability to identify American Bullfrog presence by detecting its DNA in ­water samples. Environmental DNA is amplified using PCR techniques, and tests reveal it is highly reliable at identifying wetlands where the species is pre­sent (Ficetola et al., 2009). Attempts to eliminate American Bullfrogs from areas where they have been introduced have met with mixed success. Efforts have included direct capture, trapping, electroshocking, poisons, and shooting, particularly in the Desert Southwest (e.g., Orchard, 2011; Snow and Witmer, 2011; Underwood and Letchworth, 2016; Lukey, 2017). Most efforts are ­labor intensive and only partially effective for a ­limited period of time.

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Lithobates chiricahuensis 477

Lithobates chiricahuensis (Platz and Mecham, 1979) Chiricahua Leopard Frog ETYMOLOGY

chiricahuensis: the name is derived from the Apache word Chiricahua in reference to the Chiricahua Mountains where the holotype was collected, and in recognition of the Chiricahua Apaches who inhabited the region. NOMENCLATURE

Stebbins (2003): Rana chiricahuensis Dubois (2006): Lithobates (Lithobates) chiricahuensis Fouquette and Dubois (2014): Rana (Lithobates) chiricahuensis Synonyms: Rana (=Lithobates) fisheri) in part, Rana chiricahuensis, Rana subaquavocalis Leopard frogs have long been recognized for their phenotypic variation. Dif­fer­ent phenotypes have been considered to reflect polytypic variation of a wide-­ranging species (Lithobates pipiens), to be the result of clinal variation, or to be members of a wide-­ranging multispecies complex (the Leopard Frog complex) (Ruibal, 1957; Moore, 1975; Hillis, 1988). Some assessments (e.g., Moore, 1944; Ruibal, 1957) mixed individuals from a variety of locations, resulting in considerable taxonomic confusion. In addition, ­these species often may be sympatric (e.g., Frost and Bagnara, 1977; Frost and Platz, 1983), further complicating identity. Much of the scientific lit­er­a­ture uses the name Rana pipiens for frogs now recognized as Lithobates chiricahuensis. In addition, lit­er­a­ture references to L. chiricahuensis inhabiting Arizona’s central Mogollon Rim and in the San Francisco Mountains on the Arizona–­New Mexico border may actually refer to L. fisheri. Readers should verify locations when using older lit­er­a­ture. IDENTIFICATION

Adults. This is a rather stocky leopard frog. The ground color is light to dark olive or brown with numerous small black spots. ­These spots usually lack a light halo, and they are pre­sent anterior to the eye. ­There is an incomplete white stripe on the upper lip which is diffuse in front of the eye. Dorsolateral folds are pre­sent; ­these folds are interrupted and tend to align medially ­toward the frog’s posterior. Blunt tubercles are pre­sent between the dorsolateral folds. The skin is rough. A light stripe on the upper lip is ­either faint or absent. Throats are mottled gray (unlike other Southwestern leopard frogs), and this gray color may extend onto the chest. Yellow pigmentation occurs in the groin region and ventrally on the

Dodd_Canada_int_5pgs_B3.indd 477

thighs, and often extends onto the rear part of the venter. Venters are dull and melanistic; gray mottling may be pre­sent. The posterior concealed portion of the thigh is darkly pigmented except for scattered small light spots, each containing a tubercle. Rear toes are broadly webbed. Males usually have small external vocal sacs (but see Platz et al., 1997, who characterized the vocal sacs of the Huachuca Mountain population >80 mm SUL as “well-­developed”), and the folds of the vocal sacs may be darkened. Additional information on morphology and coloration is provided by Ruibal (1957), Mecham (1968), Platz and Platz (1973), and Platz (1976). Females are generally larger than males, growing to 125 mm SUL in the Huachuca Mountains; males grow to slightly more than 100 mm SUL (Platz et al., 1997). ­There, the mean male length was 83 mm SUL, whereas the female mean was 105 mm SUL (Platz, 1993). In New Mexico, Fritts et al. (in Degenhardt et al., 1996) gave a mean male size of 64.3 mm SUL and a mean female size of 76.9 mm SUL. Brennan and Holycross (2006) gave the maximum size as 135 mm SUL. In a sample of museum specimens, Goldberg (2020g) reported males as 72–93 mm SUL (mean 78.7 mm) and females as 64–108 mm SUL (mean 84.9 mm) from Arizona. Larvae. Larval L. chiricahuensis are darker than other leopard frog tadpoles in the Southwest. The dorsal coloration is a dusky olive gray, and the dorsum contains faint black spots. The lateral sides of the tadpole are olive with large dark spots, and ­there are large bronze splotches ­toward the venter. The venter itself is grayish white with a pinkish-­ bronze sheen. Tails tend to be olive gray with large but dull olive spots. The iris is bronze. The maximum size is ca. 80 mm TL, and the tail is about 1.5 times the body length. A detailed description of the larvae and mouthparts is in Scott and Jennings (1985), and Jennings and Scott (1993) illustrated color and morphological differences between stream and pond larvae (see Larval Ecol­ogy). Eggs. ­There are no descriptions of the eggs of this species. Eggs masses are ­spherical and are attached to vegetation in shallow ­water. Clutch sizes range from 300 to 1,485 (in Sredl and Jennings, 2005). DISTRIBUTION

­ here are 2 main populations of L. chiricahuensis that are T separated into northern and southern groups by the Gila River drainage (roughly north and south of the Interstate 10 corridor; reviewed by Rorabaugh and Sredl, 2014). The northern group is found from the eastern bajada of the Black Range in the Rio Grande drainage of Socorro and Sierra counties, New Mexico (mostly the Gila and San Francisco river drainages) westward through the San Francisco Mountains

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Distribution of Lithobates chiricahuensis

barely entering Arizona. An isolated population also is found near Camp Verde in central Arizona. A southern group occurs in southwest Arizona from the Baboquivari Mountains and Altar Valley in Pima County eastward to Hidalgo County, New Mexico, in the Animas, Peloncillo, Huachuca, Dragoon, Pajarito, and Chiricahua mountains. This species has dis­ appeared from much of its historic range in the US, and currently separated populations may have been historically contiguous over a much wider range. In México, Chiricahua Leopard Frogs are found in the Sierra Madre Occidental of Chihuahua, northern Durango, and eastern Sonora. Impor­tant distributional references include: Campbell (1934), Frost and Bagnara (1977), Clarkson and Rorabaugh (1989), Degenhardt et al. (1996), Sredl and Jennings (2005), Brennan and Holycross (2006), Lemos Espinal and Smith (2007a), Hekkala et al. (2011), Rorabaugh and Sredl (2014), Murphy (2019), and Holycross et al. (2021). FOSSIL REC­O RD

No fossils are known. Holman (2003) noted Miocene (Hemphillian) fossils from Navajo County in northern Arizona that belonged to the L. pipiens complex. SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates chiricahuensis is a member of the Novirana clade of North American ranid frogs. It is an associate of the mostly Mesoamerican Lithobates montezumae group (or Lacusirana) (Hillis and Wilcox, 2005). Its closest relative in the United States is the Northern Leopard Frog (L. pipiens). Hekkala et al. (2011) extracted DNA from preserved frogs collected from the Vegas Valley, described as L. fisheri in 1893, and determined that L. fisheri was conspecific with populations of leopard frogs from the central Mogollon Rim in Arizona and the San Francisco Mountains on the Arizona–­New Mexico border, heretofore considered

Dodd_Canada_int_5pgs_B3.indd 478

L. chiricahuensis. Additional molecular research is ­under way to elucidate the relationships of L. chiricahuensis in the west central New Mexico mountains and populations in southern Arizona and México. Variation in leopard frog phenotypes from the American Southwest has been recognized for some time, with L. ­chiricahuensis often referred to as the “southern type” or form (Mecham, 1968; Mecham et al., 1973; Frost and Bagnara, 1976, 1977). The species can be separated from other leopard frogs by a combination of morphological, biochemical, auditory, and ge­ne­tic characteristics (Mecham, 1968; Platz and Platz, 1973; Platz and Mecham, 1979; Frost and Platz, 1983; Hekkala et al., 2011). Frost and Bagnara (1976) presented a ­table comparing vari­ous phenotypes among leopard frog populations. In their description, Platz and Mecham (1979) noted regional variation in the presence of vestigial oviducts, with northern males tending to lack them and southern males having rudimentary oviducts. Populations of frogs in the Huachucas, Dragoon, Pajarito, and Chiricahua mountains ­were thought to form a southern clade within L. chiricahuensis, with populations in the White Mountains and along the Mogollon Rim forming a northern clade. ­These northern clade forms have now been identified as conspecific with L. fisheri, a species known previously only from the Vegas Valley and thought to be extinct (Hekkala et al., 2011). In 1988, Platz (1993) described a leopard frog from the Huachuca Mountains in southern Arizona as Rana subaquavocalis (Ramsey Canyon Leopard Frog). One of the most distinctive characteristics of the species was its underwater advertisement call (hence its name), a character unique among leopard frogs. The range included only Ramsey Canyon and 2 other locations. Both individual and population-­level ge­ne­tic heterozygosity was low in the Ramsey Canyon and Barachas Ranch populations, and ­these populations dis­appeared by 1996 (Platz and Grudzien, 2003). The results of further molecular analyses have demonstrated that L. subaquavocalis and L. chiricahuensis are conspecific (Goldberg et al., 2004a). Natu­ral hybrids have been identified between L. chiricahuensis and L. pipiens in Arizona (Mecham, 1968; Platz and Platz, 1973; Platz, 1976; Platz and Mecham, 1979; Green and Delisle, 1985). No evidence of hybridization between L. chiricahuensis and L. blairi has been demonstrated in wild populations (Frost and Bagnara, 1977). In laboratory crosses, L. chiricahuensis produces hybrids with L. blairi, L. berlandieri, L. magnaocularis, and L. pipiens; the percentage of embryos that develop and the levels of abnormalities are variable (Mecham, 1968; Purcell, 1968; Frost and Bagnara, 1977; Frost and Platz, 1983). Surviving hybrids have very low sperm counts.

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Lithobates chiricahuensis 479

ADULT HABITAT

Lithobates chiricahuensis is found in semidesert grassland, Madrean evergreen woodland, pinyon-­juniper conifer forest, and montane conifer forest habitats. Streams and associated plunge pools in rocky canyons constitute the preferred habitat for this species. It also inhabits grassy streams, slow-­moving creeks, rocky pools, springs, pools alongside streams, beaver ponds, and even stock tanks and ditches. The species occurs to 1,034 m in the Altar Valley (Rorabaugh and Sredl, 2014). AQUATIC AND TERRESTRIAL ECOLOGY

This is a highly aquatic species that rarely ventures far from ­water. At night, however, individuals may venture from the ­water’s edge to forage. Chiricahua Frogs are sometimes observed floating on algal mats or other floating vegetation. Frogs are generally inactive from November to February, but frogs at geothermal sites may be active year-­round. During the warm season, activity follows temperature, with frogs being most active in the morning, prior to when temperatures increase. As the season progresses and ­water temperatures warm, nocturnal activity becomes more prevalent. Activity also is associated with calm winds. Refugia from cold weather and drought have not been described, but Chiricahua Leopard Frogs likely seek refuge in rock cracks and crevices, ­under tree roots, and in undercut stream banks. Long-­distance movements take place along watercourses. Sredl and Jennings (2005) offer the following information on home range. The home range size varies between the dry and wet seasons. For males, the mean is 161 m2 in the dry season and 375.7 m2 in the wet season. The largest male home range recorded was 23,390 m2 for an individual that used a stream corridor 10 m wide × 2,339 m in length. Another male was recorded to move 3.5 km. The largest female home range recorded was 9,500 m2 (10 m wide × 950 m in length). Males tend to have greater home ranges than females. In another movement study (Hinderer et al., 2017), frogs moved from breeding ponds during rainfall, with movements averaging 97 m/day along a stream drainage, albeit with much individual variation. The mean distance of 30 tracked individuals was 2,427 m. One individual moved 1,658 m in 1 day, whereas another radio-­tracked frog moved 9,888 m (total displacement of 8,506 m) in 36 days. Dispersing frogs tended to move upstream more than downstream.

tions are short (6–28 msec) and rise with time (0.4–6.6 msec) (Platz and Mecham, 1979; Platz, 1993), producing a brief audible rise in pitch. Frost and Platz (1983) and Platz (1993) provide sonograms comparing this species’ advertisement call with ­those of other Southwest leopard frogs. Most L. chiricahuensis call above ­water from the shorelines of streams and associated ­water pockets, tanks, or pools. In the Huachuca Mountains, however, Chiricahua Leopard Frogs call from 1.0 to 1.3 m underwater. This call is inaudible in air. ­Whether other populations of this species, particularly ­those of the southern clade, have an underwater call has not been published, although the possibility is mentioned by Norman J. Scott (in Degenhardt et al., 1996). Aggressive be­hav­ior between males has been recorded during the breeding season (Sredl and Jennings, 2005), but apparently does not occur at other times. BREEDING SITES

Breeding occurs in a wide variety of slow-­moving or lentic ­waters, including streams, rivers, pools in intermittent streams, beaver ponds, marshy wetlands, and springs. They also use man-­made ­water sources, such as stock tanks, irrigation sloughs, wells, and backyard ponds. Thermal springs may be a particularly impor­tant breeding site for this species, since such springs may allow year-­round breeding and activity and are ­free from predaceous nonindigenous fish. Occasionally, however, sunfish (Lepomis) have been found at sites occupied by this species, but nothing is known about the nature of the interactions between ­these species (Howell et al., 2019). REPRODUCTION

In a very small series of museum specimens, Goldberg (2020g) recorded spermiogenesis from frogs in April, July,

CALLING ACTIVITY AND MATE SE­L ECTION

Advertisement calls consist of a long, snore-­like trill. Calls consist of a single note lasting 1–2 sec with a dominant frequency of 0.9 kHz (Frost and Platz, 1983). Calls are characterized by a high pulse repetition rate (17–39 pulses/ sec) and a high number of pulses (19–68/call). Pulse dura-

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Egg mass of Lithobates chiricahuensis. Photo: Brent Sigafus

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480  Ranidae

and September. The smallest mature male was 72 mm SUL. Females considered ready to spawn ­were observed from March to November, but 2 collected from June to July ­were not ready to spawn. The smallest mature female was 64 mm SUL and was collected from November. Goldberg (2020g) interpreted ­these results to indicate a spring and autumn reproductive season, similar to L. yavapaiensis. Eggs may be deposited throughout the warm season, with rec­ords in New Mexico from April and September (Scott and Jennings, 1985) and from February to November in Arizona (Brennan and Holycross, 2006; Goldberg, 2020g). Indeed, elevation plays an impor­tant role in the timing of reproduction. At low elevations (1,800 m), it occurs mostly from June to August (Frost and Platz, 1983). According to Sredl and Jennings (2005), breeding may occur year-­round in warm-­ water geothermal springs. In Arizona, the northern clade of frogs breeds ­later than the southern clade of frogs. An extended breeding season prob­ably allows Chiricahua Leopard Frogs to take advantage of favorable environmental conditions for larval development in an arid land. Eggs are attached to vegetation and are deposited within 5 cm of the ­water’s surface. ­Under experimental conditions, embryos can develop at ­water temperatures of 12–31.5°C (Zweifel, 1968b). Egg mass temperatures in nature have been mea­sured from 12.6 to 29.5°C (Sredl and Jennings, 2005). Hatching occurs within 14 days of deposition in the Huachuca Mountains populations of this species, but at geothermal springs hatching may occur within 8 days (Sredl and Jennings, 2005). LARVAL ECOLOGY

Tadpoles have been found in New Mexico from February to November, although most prob­ably metamorphose by September. Metamorphosis occurs at 35–40 mm SUL. The bimodal breeding season and differences in thermal conditions prob­ably account for the protracted proportion of the year when tadpoles may be found. According to information in Sredl and Jennings (2005), the larval period lasts 3–9 months. The difference in the length of the larval period may reflect thermal conditions during development. In warm-­ water springs, continuous growth allows for rapid development ­toward metamorphosis, whereas in cold-­water habitats development proceeds slower and larvae may overwinter. Tadpoles have been observed ­under ice in ­water 5°C (Sredl and Jennings, 2005). Tadpoles of this species may be found in a variety of habitats, from streams to ponds. The habitat in turn influences larval coloration and morphology. Larvae from streams have more contrasting and blotched patterns on the tail. They have

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Tadpole of Lithobates chiricahuensis. Photo: Cecil Schwalbe

thicker dorsal tail fins and larger tail muscles than larvae taken from ponds. Even within streams, the level of tail thickening can vary, perhaps in response to stream flow. ­These differences are illustrated by Jennings and Scott (1993). Ponds may produce substantial numbers of metamorphs. Hinderer et al. (2017) recorded 2,459 juvenile captures from 2 stock ponds over a 2 yr period. Most frogs appeared to orient ­toward a nearby creek. Movements from ponds depended on daily rainfall, with the number of daily captures proportional to rainfall amount. DIET

Larvae are herbivorous. The diet of postmetamorphic Chiricahua Leopard Frogs has not been examined. Like other leopard frogs, it prob­ably feeds on a variety of invertebrates and even small vertebrates in relation to their availability. PREDATION AND DEFENSE

This species is preyed upon by garter snakes (Thamnophis cyrtopsis, T. elegans, T. eques) and Sonoran Whipsnakes (Coluber [= Masticophis] bilineatus) (Rorabaugh and Sredl, 2014; Drost, 2020; Jones and Hensley, 2020; Jones and ­Sullivan, 2020; Jones et al., 2020). Adults are likely eaten by a wide variety of other snakes, birds, and mammals. Larvae are eaten by aquatic invertebrates and Tiger Salamanders (Ambystoma mavortium). Upon the approach of a potential predator, postmetamorphs jump into the ­water but do not have an alarm call. In low temperatures and reflectance, the ventral skin coloration tends to darken, which may make this species less obvious to potential predators. POPULATION BIOLOGY

Chiricahua Leopard Frog populations consist of a series of small subpopulations (85 mm SUL, but that Bronze Frogs rarely exceeded 75 mm SUL. Males and females are about the same maximum size in some areas (Martof, 1956a; Smith, 1961; Fleming, 1976), although females have been reported to be larger than males in ­others (Ryan, 1953; Jenssen and Klimstra, 1966; Meshaka et al., 2009a, 2011c). The reverse is also sometimes true. Differences in mean sizes are small, however, and not much more than about 3 mm. Thus, this species cannot be generalized to be sexually size dimorphic. Larvae. Tadpoles are large (80–100 mm TL; Logier, 1952) but not deep bodied, and are olive green with small to large dark markings. Throats are usually white, and bellies are a deep cream color with no iridescence (but see Systematics and Geographic Variation). The tail is green and mottled with brown, and without pinkish to buff spotting. The dorsal fin terminates posterior to the spiracle, which is located on the left side of the body. ­There are no stripes on ­either the dorsal fin or on the tail musculature. The eyes are dorsally located, and the oral disc is emarginate and narrowly pigmented. The anus is dextral rather than located medially. Green Frog larvae may be in the same developmental stages, but still be very dif­fer­ent in size, depending upon the pond conditions in which they develop (Rogers, 1999). Descriptions of larvae are in Wright (1914, 1929, 1932), Altig (1970), and Priestley et al. (2010). Yellowish-­white tadpoles ­were reported from Tennessee (Dyrkacz, 1981), an unusually colored specimen from Michigan (Bowen and Beever, 2010), and albinos from Québec (Saumure, 1993).

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Eggs. The black-­and-­white eggs are laid in a large (for example, 12 cm × 17 cm or 15 cm × 21 cm), irregular surface film; each film (maximum dia­meter 30 cm) contains from 1,000 to >5,000 eggs amid emergent or floating vegetation. The surface film may float ­free, or it may be attached to aquatic vegetation. Surface films sometimes break apart, giving the impression of smaller clutch sizes, or films oviposited by separate females may merge, extending the extent of the film and increasing perceptions of the numbers of eggs (Wright, 1914). Although somewhat similar to the surface film egg masses of American Bullfrogs, ­there are many fewer eggs in Green Frog masses. Surface films allow for better oxygen­ ater temperatures in which this species ation in the warm w breeds. The eggs are deposited in relatively shallow ­water (36°C. Martof (1953a) occasionally found frogs at temperatures below 15°C, but such frogs ­were inactive or rarely encountered. Adult and juvenile Green Frogs often remain near ­water during the postbreeding season, where they occupy ephemeral wetlands, swamps, marshes, and stream habitats (Pitt et al., 2017). However, they forage extensively in terrestrial habitats, particularly in areas with deciduous leaf ground cover (Pitt et al., 2017), where food resources are greater than in proximity to ­water. During ­these forays, they gain body mass. In New York, Lamoureux et al. (2002) recorded 0–7 forays per frog from mid-­August ­until late October. Movements between pond and terrestrial foraging areas occurred with precipitation and in less than 24 hrs. Frogs moved from 10 to 99 m (mean 36 m), and forays lasted from 7 to 408 hrs (mean 88 hrs). Foraging sites had dense terrestrial vegetation with thick layers of leaf litter; frogs ­were usually observed well hidden, with only their noses vis­i­ble. In South Carolina, Pitt et al. (2017) tracked 6 Green Frogs for a mean of 51 days over a total path distance of 43–394 m. Green Frogs move to protected sites in the fall and enter winter dormancy from mid-­September to mid-­November in the North. Such sites may not be in the immediate proximity of the summer activity centers. For example, frogs in New York moved 100–560 m to reach overwintering sites in distant streams, seeps, and a beaver pond (Lamoureux and Madison, 1999), and in Missouri, they moved from 550 to 900 m (Birchfield, 2002). The length of dormancy changes with latitude and environmental conditions but can last more than 5–6 months; overwintering in Michigan occurs from late October to early November ­until late March–­early April (Martof, 1953a, 1956b). Frogs become active when daily maximum temperatures reach 15°C for several days and mean daily temperatures are >4°C. Rainfall helps stimulate an end to winter dormancy. Overwintering normally occurs in aquatic habitats in mud and bottom debris (Dickerson, 1906; Walker, 1946; Wright and Wright, 1949; Logier, 1952; Lamoureux and Madison, 1999; Birchfield, 2002). Streams seem to be preferred, as the flowing ­water remains unfrozen and provides oxygenation during the long winter months. Males and females choose the same overwintering sites and remain ­there ­until dormancy ends. Gorham (1964) noted in New Brunswick that L. clamitans are observed around springs in November. Green Frogs also overwinter in terrestrial situations in soil pockets below the leaf litter, or in crayfish burrows (Badje et al., 2016). In such a situation, Bohnsack (1951) noted air temperatures of 0–2.5°C surrounding an overwintering Green Frog in Michigan, and that ice crystals ­were occasionally found in the soil

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near the frog. Winterkill may occur in such sites, however; Lannoo et al. (1998) recorded a mass mortality event of subadult Green Frogs found adjacent to a wetland in Iowa, and suggested that winterkill resulting from a poorly chosen terrestrial site was the most likely explanation. Green Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Pearse, 1910; Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Green Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling occurs over an extended period in spring and summer, but Green Frogs become active several weeks prior to the breeding season. Male Green Frogs are territorial, with larger males occupying larger and better-­quality habitats than smaller males. During the course of a breeding season, males often occupy more than 1 territory (usually 2–5, mean 3.7; Wells, 1977b), and they may spend from 1 to as long as 7 weeks within a par­tic­u­lar territory. Males move back and forth between defended territories. Large males also occupy their territories longer throughout the breeding season than small males. Occasionally, a group of males ­will maintain a spatial relationship to one another (Martof, 1953b), but with the shifting of territories, most spatial relationships change constantly, especially during the breeding season (Shepard, 2004). This may explain the seemingly random nonterritorial distribution pattern reported by Shepard (2002), especially since his study was conducted for only a short-­term period. Territories (4–6 m in dia­meter) are defended during the breeding season, and males aggressively challenge intruders, who sometimes remain as satellite males in the anticipation of a territory becoming available (Wells, 1978; Shepard, 2004). In such encounters, males use patrolling, splashing displays, vocalizations (“growling”: Jenssen and Preston, 1968), chases, attacks, and wrestling to establish dominance over their territory (Brode, 1959; Schroeder, 1968; Wells, 1978). Aggressive encounters often involve the display of the yellow throat patch ­toward the intruder, and vocalizations occur continuously during wresting and shoving matches. Most encounters are short lived, but some can last a considerable amount of time (45 min). Larger males are normally successful in aggressive encounters. Territory quality is determined by the extent of vegetation in the shallow ­water of the potential oviposition site, and females choose males based on the quality of the habitat and male size. In turn, males with high-­quality territories mate

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more often (to 5 times) than ­those with lower-­quality territories, although some males are not successful despite the quality of their territory. Wells (1977b) suggested that the intense male-­male competition for mates evolved as a result of the prolonged breeding season, and described the territorial be­hav­ior of males in detail. Calling sites are located in shoreline vegetation where ample cover is provided. Males call from vegetation near the shoreline or while floating on the ­water’s surface, and they may swim between nearby vegetation mats between calls. Males usually are spaced about 2–3 m from one another, but they may call from much closer proximity (Martof, 1953b). If calling from the shoreline, the male often creates a small pool to sit in by rotating the hind limbs back and forth. Wells (1977b) noted that pool construction occurred when pond ­water levels ­were low or decreasing. Wells (1978) identified 4 types of calls, in addition to a release call. ­These are: Type I (spontaneous calls given day or night, 1 single note); Type II (a high intensity advertisement call of 3–4 notes delivered in rapid succession, usually in response to a disturbance in the territory); Type III (similar to Type I but more explosive, directed ­toward an opponent in an agonistic encounter); and Type IV (a long, low-­ frequency call often given by the winner of a wrestling bout, perhaps as a warning). The release call (Type V) may be given both by males and females. The dominant frequency of the call of male Green Frogs is 416–544 Hz (Given, 1990), and the dominant frequencies of the advertisement calls are inversely correlated with the caller’s snout-­vent length (Ramer et al., 1983). Males can assess the dominant frequency of the calls of conspecific males and are thereby able to assess the size of a rival. If the rival is a large male, they ­will lower the dominant frequency of their call. However, if the rival is a small male, they do not change their own dominant frequencies (Bee et al., 1999). Ramer et al. (1983) found that small males increased the rate of agonistic vocalizations directed ­toward other small males, whereas large males decreased their rate of baseline calling. Large males directed agonistic calls to other large males but responded ­little if at all to small males. ­These be­hav­iors suggest that males know the sizes of males in adjacent territories, and this helps mediate or direct territorial aggression. The calls used in territorial aggression are Type III calls, which have lower dominant frequencies and a longer note duration than the somewhat similar Type I advertisement calls (Bee and Perrill, 1996). Males do not respond to heterospecific calls, despite the close proximity of calling males (Given, 1990). Calling occurs both diurnally and nocturnally, but the greatest intensity of calling occurs from shortly ­after midnight ­until dawn (Mohr and Dorcas, 1999; Cook et al.,

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2011). Northern Green Frogs generally call in the spring to midsummer, whereas Bronze Frogs ­will extend calling into late summer or even early autumn. In New Brunswick, for example, calling occurs from late April to mid-­June (Gorham, 1964), in Michigan and Pennsylvania from May through mid-­August (Meshaka et al., 2015b), in Mas­sa­chu­ setts from April to August (Cook et al., 2011), in West ­Virginia from March to July (Meshaka et al., 2009d), and in Texas from March to September or October, depending upon location (Meshaka et al., 2011c; Car­ter et al., 2018). In Louisiana, calling occurs from March to September (Dundee and Rossman, 1989; Meshaka et al., 2009a), although calls are heard as early as January (Meshaka et al., 2009b). Males often begin calling from pools and isolated portions of streams early in the season, but they ­will leave ­these temporary calling locations ­after 2–4 weeks and move to larger lakes and ponds, where calling begins in earnest. At Cape Cod, for example, the peak duration of the calling period lasted 42 days out of a 141 day calling season (Cook et al., 2011). Males frequently move during dry weather, but females appear to require both warm weather and precipitation for movement and breeding activity. Calling occurs at air temperatures from 11 to 29°C, with most activity at ca. 22–24°C (range 18–29°C) (Meshaka et al., 2009b, 2011c); the level of calling increases as temperatures increase (Steen et al., 2013). Environmental conditions influence the timing of calling, but the way they impact call activity varies seasonally. At the beginning of the season in Nova Scotia, for example, males generally wait for warm ­water temperatures (>22°C) between sunset and sunrise before beginning to call in earnest, and calling is associated with high relative humidity and high barometric pressure (Oseen and Wassersug, 2002). As the season progresses, ­water temperature becomes less impor­tant as summer temperatures climb, and frogs call at lower barometric pressures associated with rainfall. Humidity also becomes unimportant ­later in the season, as frogs attempt to reproduce before cool weather sets in. Precipitation is impor­tant in initiating spring movements to breeding ponds, and ­later in the season in initiating calling when ­water levels are decreasing; it is not impor­tant in triggering calling activity in the spring. Calling is not affected much by windy or cloudy conditions, although most calling in Texas occurs at still or very low wind speeds (Meshaka et al., 2011c). At Cape Cod, the mean calling time was 206 minutes (Cook et al., 2011). Green and Bronze Frogs breed at sites frequented by many other frog species, including some with loud and frequent calls of their own. In order to avoid sound interference, Green Frogs actively space their calls during ­silent gaps in the calls of other species, such as American Bullfrogs

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(Herrick et al., 2018). This is especially obvious when the 2 species are in close proximity to one another (nearest neighbors), in which case the timing of the calls does not overlap. In Michigan, males tend to congregate in ponds or other suitable sites during the entire breeding season, whereas females are found more along small streams when they are not at breeding sites (Martof, 1956b). Martof (1956b) suggested that females rarely spent more than a week at the main breeding sites, quickly returning to nonbreeding habitats ­after ovipositing their eggs. Thus, ­there appears to be a degree of adult habitat partitioning which allows spent or nonreproductive females to avoid contact with reproductive males. At the breeding site, females in a head-­down posture approach the territory of a calling male, and they may visit several territories prior to selecting a mate. The cues by which females choose males may vary, but Schulte-­Hostedde and Schank (2009) have shown that males with larger forearms and intermediate shades of yellow on the throat are in better body condition than other males. In addition, the size of the tympanum may provide the female with cues for mate se­lection. ­These multiple cues likely help the female refine her choice based on the condition of a calling male. The male Green Frog approaches the female from ­behind while holding her hind limbs, and then guides her body under­neath his forelimbs as he floats in the ­water. Amplexus is axillo-­pectoral—­that is, halfway between axillary and pectoral. The male holds on firmly to the female by pressing his thumbs into the female’s lateral sides. Spawning takes place in the male’s territory as the pair floats just beneath the ­water’s surface. Just prior to spawning, the female lowers her head below the ­water’s surface, arches her back concavely, and brings her cloaca to just above the ­water’s surface. The male then slides backward and rotates his thighs upward. This brings the cloacas of the pair into near (3 mm) contact. Eggs are extruded 30–50 at a time, and the male uses his rear feet to direct the eggs to his cloaca, where they are fertilized. The male then uses his toes and feet to push the eggs away from the amplexed pair. The pair then repeats the deposition/fertilization pro­cess ­until all the eggs are oviposited. Oviposition lasts 10–25 min. The males releases the female as soon as she abandons the oviposition posture. Males and spent females have a warning croak indicating nonreceptivity to a suitor male (call Type V), but the female does not regain her voice ­until about 24 hrs ­after oviposition. In the meantime, she may be clasped by another male. The mating sequence and postures of Green Frogs are described and illustrated in detail by Aronson (1943a). Amplexus by males with females of other ranids (L. pipiens, L. palustris, L. sylvaticus) has been reported (Wright, 1914).

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BREEDING SITES

Green Frogs are found in a wide variety of breeding sites, usually with long hydroperiods. They prefer lakes, ponds, and slow-­moving permanent streams, although they can be found in relatively small woodland pools. They use artificial breeding sites, such as retention ponds (Birx-­Raybuck et al., 2010; Brand and Snodgrass, 2010), and canals or ditches with slow-­moving ­water. McCarthy and Lathrop (2011), however, noted that while calls ­were heard at retention ponds, successful breeding did not occur ­there, perhaps ­because of the presence of fish and other predators. In experimental mesocosms, increased depth and area led to lower survivorship but greater growth of the survivors when densities ­were kept constant (Pearman, 1993), but observations on Green Frogs in the field do not suggest that wetlands are selected on the basis of size for breeding, much less that small ponds produce metamorphs of higher fitness. In Newfoundland, the presence of Green Frogs is negatively correlated with dissolved oxygen content (preferred DOC 5–10 mg/L), but positively correlated with pond permanence and the presence of ­human residences nearby (Campbell et al., 2004). The positive association between Green Frogs and ­human occupation, however, likely reflects human-­mediated dispersal in this area. In Ontario, Hecnar and M’Closkey (1996b) could find no appreciable differences in pond ­water chemistry between occupied and unoccupied sites. REPRODUCTION

The breeding period of Green Frogs is prolonged throughout the spring and summer at many locations (but see Berven et al., 1979), rather than occurring over a short period of days to weeks as in many other anurans. Testis size was

Eggs of Lithobates clamitans. Photo: Dana Drake

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492  Ranidae

largest in spring, and females ­were gravid from April to September in Texas, mirroring an extended reproductive season (Meshaka et al., 2011c). In contrast, testis size was largest in July and August in Pennsylvania, with gravid females observed from May to July (Meshaka, 2013b). Lit­er­a­ture rec­ords of breeding dates include the following: March–­July (northern Louisiana: Meshaka et al., 2009a), March–­September, with a peak of April and May (southern Louisiana: Meshaka et al., 2009b), April–­June (Rhode Island: Anonymous, 1918; Paton and Crouch, 2002), April–­August (­Virginia: Gibson and Sattler, 2020; North Carolina: Gaul and Mitchell, 2007; Rhode Island: Raithel, 2019), May and June (Indiana: Brodman and Kilmurry, 1998), May–­July (Mary­ land: Lee, 1973a; New Hampshire: Oliver and Bailey, 1939; Ohio: Walker, 1946; Pennsylvania: Meshaka, 2013b), May–­August (Michigan: Martof, 1956a, 1956b; Minnesota: Fleming, 1976; Oldfield and Moriarty, 1994; New Brunswick: Gorham, 1970; New ­England: Klemens, 1993; New York: Wright, 1914; Gibbs et al., 2007; Ontario: Piersol, 1913; Pennsylvania: Meshaka and Morales, 2020), May–­September (Mary­land and ­Virginia: Berven et al., 1979), June and July (Illinois: Cagle, 1942; Nova Scotia: Gilhen, 1984; Ohio: Varhegyi et al., 1998; ­Virginia: Berven et al., 1979; West ­Virginia: Rogers, 1999), and June–­August (New York: Wells, 1977b; Tennessee: Gentry, 1955). It is likely that the ­actual breeding season (that is, when eggs are deposited) extends beyond or is less than some of the monthly rec­ords found in the lit­er­a­ture, depending on weather conditions. Calling usually precedes mating by up to a month or so, which in turn affects perceptions of the timing of the breeding season. For example, calling begins in April in lowland sites in the Shenandoah Valley, ­Virginia, although breeding does not actually occur ­until mid-­May (Berven et al., 1979). Many lit­er­a­ture reports do not make the distinction between when calls are heard and when breeding actually takes place. In addition to a latitudinal effect on breeding dates in such a widespread species, elevation also affects the amount of time available for reproduction. In lowland areas of Mary­ land and ­Virginia, for example, breeding is prolonged from mid-­May to September. In the higher elevations of the Alleghany Mountains in ­Virginia, the time of egg deposition is much shorter, extending only between July and August (Berven et al., 1979). The timing of the ovarian cycle likely varies according to region. In Louisiana and Texas, observations of gravid females begin early in the season and increase rapidly in spring and summer. Some females yolking eggs may be found at any time of the year, but advanced-­stage gravid females are usually not found in any frequency in the autumn. It is at this time that they begin acquiring the lipids needed for their

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eggs to be deposited the following breeding season, a pro­cess that ­will continue right up to egg deposition (Meshaka et al., 2009b). Fat reserves are usually depleted by July or August. Clutch size may vary with phenotype (and hence latitude) and size of the female, but data are ­limited. Trauth et al. (1990) recorded clutch sizes of 4,924 and 5,730 in two Bronze Frogs, and 2,851 from a single Green Frog; Martof (1956b) recorded 3 clutches of 3,800, 4,100, and 4,300 eggs in Michigan; clutch size ranged from 1,401 to 5,289 in 10 Green Frog clutches from Nova Scotia (Gilhen, 1984); Wright (1932) recorded a single clutch of 1,451 eggs from a Bronze Frog in the Okefenokee Swamp; Meshaka et al. (2009a) estimated a mean of 2,550 eggs (range 1,600–4,200) in Louisiana; and the clutch size of Green Frogs averaged 4,631 (range 2,334–6,467) in Pennsylvania (Meshaka and Hughes, 2014) and 5,830 (range 3,786–9,215) in West ­Virginia (Meshaka et al., 2009d), and was estimated as 2,750–3,300 in a second Pennsylvania study (Meshaka, 2013b). Small females may deposit as few as 1,000 eggs, perhaps reflecting regional variation in female size (Pope, 1964). Green Frogs often have 2 sets of eggs developing within an ovary, 1 set mature and the other in a much less advanced state of development. Martof (1956b) therefore suggested that maturation of eggs requires 1 full year prior to oviposition. Green Frogs normally oviposit 1 clutch per season, but double clutching has been reported (Wells, 1976). Egg deposition occurs at ­water temperatures of 22–32°C in Pennsylvania (Meshaka and Morales, 2020). Eggs kept in cold ­water (23°C than they do at 20°C or less (Warkentin, 1992a), and thus feed more actively during the ­middle of the day than at night or in the morning when temperatures are cooler. It is not surprising, then, that larvae exhibit a daily rhythm in their tolerance to high temperatures, a rhythm that mirrors daily ­water temperatures. Larvae acclimated to specific temperatures have a higher CTmax than larvae that are not acclimated, and this allows them to adjust rapidly to changing temperatures within a pond (Willhite and Cupp, 1982). Unlike temperature, illumination per se has no effect on larval feeding, a result that is not unexpected for a suspension feeder (Warkentin, 1992a). Microhabitat choice plays an impor­tant role in larval feeding ecol­ogy, and likely on subsequent growth and the

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time to metamorphosis as well. During the day, tadpoles tend to concentrate in vegetated areas to feed, but at night some tadpoles, particularly larger tadpoles, move to deeper ­waters where feeding efficiency is not as ­great. Feeding rates are not dif­fer­ent between vegetated and more open ­water areas during the daylight, and small tadpoles do not move to open areas at night (Warkentin, 1992b). ­These results suggest that feeding efficiency is not the only consideration in the choice of microhabitat se­lection. Perhaps larvae use the vegetated areas during the day ­because of increased cover, with larger larvae moving to deeper ­waters at night to escape shoreline predators. Variation in growth rates is genet­ically based, such that larvae are adapted to metamorphose as rapidly as pos­si­ble given ambient temperature constraints (Berven et al., 1979; Watkins and McPeek, 2006). Larvae grow slower but longer at high elevations, and presumably in northern latitudes. Larval size is also affected by length of development. In montane ponds, larvae are consistently larger at any developmental stage than are low-­elevation larvae, and thus juveniles resulting from high-­elevation ponds are larger at metamorphosis than ­those from lower-­elevation ponds. Berven et al. (1979) noted that size at metamorphosis and mean pond temperature ­were significantly negatively correlated ­under field conditions. Stress can help speed up metamorphosis. Martof (1956a) noted that Green Frogs transformed within the first season (within 70–85 days of oviposition) in Michigan when the developmental pond dried up. Presumably, such metamorphs would be smaller than ­those that develop normally, and this may account for some of the extremes in variation observed in the size of metamorphs. For example, I found very small (15 mm SUL) metamorphic Green Frogs in shallow drying woodland pools in the ­Great Smoky Mountains National Park, yet large (35 mm SUL) metamorphs emigrating from a permanent pond about 20 km away. The large metamorphs likely originated from the previous year’s reproductive cohort, whereas the small metamorphs ­were escaping from a currently desiccating pool (Dodd, 2004). As a result of variation in larval period, the age of metamorphs and age at maturity ­will vary with the conditions ­under which larvae developed, even within a geographic region. Large spring-­transforming metamorphs ­will be at least 9 months older than small fall-­transforming metamorphs, and they ­will prob­ably reach sexual maturity faster ­after metamorphosis than small metamorphs. In addition, the larger juveniles should have greater survivorship than the smaller juveniles, especially as winter approaches. Still, transforming at a smaller size and delaying maturity may be less of a risk than remaining as larvae in a drying pond.

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In situations where larvae develop in the absence of environmental stress, the size at metamorphosis is the same for individuals completing development within a single season as for individuals that overwinter (Martof, 1956a). Size at metamorphosis may vary with latitude and elevation in accordance with developmental pond temperatures, and the fact that tadpoles and metamorphs can be of greatly dif­fer­ent sizes is often confusing to naturalists. Lit­er­a­ture rec­ords of size at metamorphosis include: a mean of 33 mm SUL and 3.6 g in Rhode Island (Raithel, 2019); 24–36 mm (mostly 26–30 mm) in West ­Virginia (Rogers, 1999); 28.4–36.3 mm (mean 32.6 mm) in Michigan (Martof, 1956a); 28–38 mm (mean 32 mm) in New York (Wright, 1914; also see Ryan, 1953); 28–39 mm (mean 33.2 mm) in Nova Scotia (Gilhen, 1984); 24–32 mm in Pennsylvania and New York (Meshaka, 2011a; Warny et al, 2012); 18–34.5 mm (mean 29.4 mm) in West ­Virginia (Meshaka et al., 2009d); 19–32.4 mm (mean 27.3 mm) in northern Louisiana (Meshaka et al., 2009a); 19.6–47.0 mm SUL (mean 28.3 mm) in southern Louisiana (Meshaka et al., 2009b); and 18.6–27.2 mm SUL (mean 22.2 mm) in Texas (Meshaka et al., 2011c). In laboratory experiments, montane Green Frogs from ­Virginia transformed at means of 19.8–36.1 mm, depending on constant temperature raised, whereas lowland Green Frogs transformed at means of 22.2–46.3 mm (Berven et al., 1979). As noted above, frogs occasionally metamorphose at smaller sizes (Walker, 1946; Martof, 1956a; Dodd, 2004). Larvae are eaten by a wide range of predators. Consequently, they have evolved both morphological (color) and behavioral means (rapid burst speeds) to avoid detection or escape predators. Traits such as burst speed, growth rate, and size are heritable, although swimming activity in the presence or absence of predator chemical cues is not (Watkins and McPeek, 2006). Burst speeds also are not correlated with predator avoidance, such as activity and the plasticity associated with it. Thus, ­there is a trade-­off between larval predation risk and foraging gains associated with increased activity that may have a ge­ne­tic basis, at least in part. Green Frog larvae respond to alarm cues given off by conspecifics and pheromones emanating from would-be predators, such as dragonfly larvae (Anax), by avoiding areas containing ­these mixed chemicals (T. Brown et al., 2019). Spatial avoidance did not occur in response to ­these cues when pre­sent singularly. Schiwitz et al. (2020), however, found no effect on larval activity in the presence of conspecific alarm cues. The percentage of individuals successfully completing metamorphosis varies considerably from year to year, depending upon pond and weather conditions. In Ontario, Shirose and Brooks (1997) recorded 32.5–85.3% of larvae transforming successfully over a 6 yr period. In Missouri,

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2 breeding ponds produced 3,206 metamorphs from 86 females (Hocking et al., 2008). Temporary breeding sites may occasionally dry before metamorphosis is completed, leading to mass mortality (Tinkle, 1959) and, although occasionally used, such temporary sites are not preferred for reproduction. DISPERSAL

Dispersal occurs rapidly a­ fter metamorphosis, and Schroeder (1976) found that nearly all frogs dispersed within 27 days ­after the first frogs transformed. However, Martof (1956b) suggested that nearly all frogs dispersed from a Michigan pond within a week of transformation. Dispersal from a West ­Virginia site occurred primarily from late June to early August (Rogers, 1999), from Missouri ponds as late as mid-­to late October (Hocking et al., 2008), and in Rhode Island from early July to early September, but particularly in August (Paton and Crouch, 2002). Unlike American Toads and Wood Frogs, Green Frogs show no special changes in activity metabolism or aerobic capacity immediately prior to dispersing from the natal pond (Pough and Kamel, 1984). Juveniles do not take any par­tic­u­lar route but disperse in all directions from the breeding pond (Schroeder, 1976). Dispersal pathways take advantage of topographical features, however, such as stream drainages and other wetlands interspersed throughout the landscape. Most of the dispersal at Martof’s (1953a) study site occurred upstream away from the natal site. A few frogs may not disperse at all but remain in close proximity to the pond in which they developed. Most frogs dispersed 183–448 m from Schroeder’s (1976) ­Virginia pond, but several frogs moved up to 4.8 km from the breeding pond. Other long-­distance movements are 457 m (Oldham, 1967), 600 m (Martof, 1953a), and 560 m (Lamoureux and Madison, 1999).

Displaced frogs do not have a tendency to home ­toward their original capture site if moved long distances, but they often move to streams and creeks where overwintering ­will occur. When dispersing, juveniles tend to avoid woody debris cover on the surface of the ground (Gravel et al., 2012). The abundance of dispersing juveniles also increases with minimum air temperature and precipitation. In Louisiana, juveniles are found in terrestrial situations throughout the late winter and early spring (Liner, 1954). Liner (1954) stated that “sometimes the woods seem to be covered with them.” At a 325 m2 pond in ­Virginia, Schroeder (1976) estimated ­there ­were 512 newly transformed Green Frogs dispersing at about the same time. In Michigan, Martof (1956b) estimated 10,500 newly transformed Green Frogs within a 6,090 m2 area. In contrast, 106 females produced only 2,214 metamorphs over a 16 yr period at a 1 ha Carolina Bay in South Carolina (Semlitsch et al., 1996); substantial numbers of metamorphs ­were produced in only 2 of the 16 years. This latter site did not fill with ­water in a number of years, and the numbers highlight the disparity in reproductive success of L. clamitans at temporary vs. permanent breeding sites. Green Frogs disperse randomly and take up residence at ponds, streams, and wetlands they encounter along the way. This means that frogs originating from 1 par­tic­u­lar pond may be found at virtually any other pond within an area, which ensures colonization of a wide variety of breeding sites and results in a fairly panmictic population. ­Because of this extensive dispersal, ­there is no reason to suggest Green Frog populations are structured as metapopulations ­either locally or regionally (Smith and Green, 2005). Instead, colonization and extinction estimates tend to be constant through time, except that wetlands with longer hydroperiods tend to have lower extinction probabilities than wetlands with short hydroperiods (Mattfeldt et al., 2009). Searcy et al. (2018) found that dispersal was positively correlated with body size but not with body condition or relative hindlimb length. Mortality of juveniles is quite high during the dispersal phase. DIET

Adult Lithobates clamitans, Northern Green phenotype. Photo: David Dennis

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Tadpoles feed on a variety of diatoms and filamentous algae, with blue-­green algae, protozoa, and microcrustaceans occasionally consumed (Munz, 1920; Jenssen, 1967). ­These items are grazed or picked up in muddy sediments from the pond or lake bottom. Larvae also have been observed eating the eggs of Wood Frogs (Jennette, 2010). Feeding occurs throughout the overwintering period. As a tadpole metamorphoses, epidermis is found within the gut, but feeding does not commence ­until the forelegs are well developed and the body mea­sures greater than approximately 32 mm. It is then

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pos­si­ble to find a wide variety of small invertebrates within a juvenile’s stomach, especially terrestrial crustaceans, land hemiptera (true bugs), spiders, flies, and beetles (Munz, 1920); it appears that juveniles ingest just about any small animal they can. Rogers (1999) found that insects comprised 93% of the 341 prey found in 65 metamorph stomachs; nearly all prey ­were of terrestrial origin. Green Frogs feed opportunistically year-­round when weather conditions permit. The greatest amount of feeding occurs in the spring, and adult females consume more food during the breeding season than adult males. Juveniles tend to forage more in the winter than adults. Aquatic organisms make up about one-­third of the diet of both adults and juveniles. The most impor­tant prey are lepidoptera (moths, including caterpillars), beetles, and snails and slugs, but spiders and flies also are eaten year-­round, depending on availability. Prey include both ground and ­water beetles, lymnaeid snails, crane flies, millipedes, katydids, grasshoppers, dragonflies and dragonfly nymphs, wasps, ants, walking sticks (McAllister and McAllister, 2013), and wolf spiders. Green Frogs are visually oriented predators that have been observed to stalk crayfish from a distance of 12 m (Hamilton, 1948). In winter, amphipods become the most impor­tant prey, at least in Illinois (Jenssen and Klimstra, 1966). ­These latter authors also found Blanchard’s Cricket Frogs (Acris blanchardi) and a small amount of plant material, perhaps inadvertently ingested, eaten by some frogs. Gilhen (1984) recorded Spring Peepers (Pseudacris crucifer) in the diet, and noted a Green Frog adult eating a conspecific tadpole. Cannibalism was also reported in Mary­land (Harris, 2013). DeGraaf and Nein (2010) recorded a hatchling spotted turtle (Clemmys guttata) eaten by an adult Green Frog, and Vergeer (1948) reported a Green Frog eating a jumping mouse (Zapus hudsonius). While the general prey items are similar in dif­fer­ent areas of North Amer­i­ca for both adults and juveniles (Surface, 1913; Hamilton, 1948; Whitaker, 1961; Jenssen and Klimstra, 1966; Stewart and Sandison, 1972; Gilhen, 1984; Werner et al., 1995; Forstner et al., 1998; Rogers, 1999), the frequency and volume of the prey taken change with availability and local invertebrate faunas. Regardless of frog size, frogs select the largest prey available that they can ingest, and ­there is a significant positive relationship between dietary niche breadth and among-­ individual diet variation (Cloyed and Eason, 2017). PREDATION AND DEFENSE

The best defensive attribute of Green Frogs is their protective coloration, which allows them to blend in with both terrestrial and emergent aquatic vegetation. Simply by sitting quietly they avoid detection. The most common active

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defense used by Green Frogs at the approach of a predator is to jump into the nearest body of ­water and give a short, high-­pitched cry that tends to startle or disorient the would-be attacker. As a result, the predator tends to lose track of the frog’s entry into ­water, and the frog quickly swims ­under ­water and buries into submerged debris and leaf litter. Alternatively, a frightened frog ­will swim sharply to one side and reemerge a few meters away, where it ­will sit motionless, blending in with pond vegetation (Bragg, 1945a). McKnight and Howell (2015) found that L. clamitans leaped away at a mean distance of 17.8 cm from an approaching model snake; ­there was no difference between males and females or any correlation with frog size regarding flight initiation distance. Juvenile Green Frogs ­will crouch and cease to move upon the approach of snakes, which makes them less likely to be detected and attacked (Marchisin and Anderson, 1978; Heinen and Hammond, 1997). They also retain the tendency to be immobile and crouch for a considerable time ­after the threat has left, thus making them less likely to be detected should the predator backtrack. When contacted, they ­will usually rapidly hop 2–5 times during the escape attempt. Body inflation also has been reported in both adults and juveniles; this allows the frogs to appear larger and perhaps more formidable than they ­really are (Marchisin and Anderson, 1978). Based on predator ­trials, it does not appear that postmetamorphic L. clamitans are distasteful or noxious to vertebrate predators (Formanowicz and Brodie, 1979). Larvae are unpalatable to some species, although the effectiveness of larval unpalatability is variable (Adams et al., 2011). As the tadpoles grow larger, they are more palatable, but their larger size makes them less vulnerable to attack by some predators. Tadpoles are avoided by vertebrate predators but not by invertebrates (Smith et al., 2019), and Shulse et al. (2013) found that mosquitofish had no effect on larval abundance in newly constructed ponds in Missouri. The eggs of L. clamitans are highly palatable to both fishes and invertebrates (Licht, 1969a; Smith et al., 2019). The mosquito Culex territans feeds on Green Frogs and even can use the frog’s call to locate its next blood meal (Bartlett-­Healy et al., 2008). Culex peccator also feeds on Green Frogs (Cupp et al., 2004). Other predators of adults include frogs (Lithobates catesbeianus), salamanders (Amphiuma means), turtles (Chelydra serpentina, Emydoidea blandingii), snakes (Agkistrodon piscivorus, Nerodia sipedon, Thamnophis sirtalis, T. sauritus), crows (Corvus), herons (Ardeidae), and hawks (Wright, 1932; Richmond, 1952; Martof, 1956b; Werner et al., 1995; Birchfield, 2002; Mitchell, 2013). Mammals such as mink, otters, and raccoons undoubtedly take a considerable toll (e.g., Lamoureux and

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Madison, 1999). Large L. clamitans also ­will consume small L. clamitans. Larvae are eaten by predaceous diving beetles (Dytiscus), ­water beetles (Hydrophylidae), whirly gigs (Gyrinidae), adults and nymphs of ­giant ­water bugs (Belostomatidae), waterscorpions (Nepidae), backswimmers (Notonectidae), dragonfly naiads (Aeschnidae, Libellulidae), salamanders (Amphiuma; Mitchell, 2013), and turtles (Glyptemys insculpta; Ernst, 2001). Catfish (Ictalurus) and bass (Micropterus, Ambloplites rupestris) prey upon both tadpoles and small frogs, although Green Frogs are often found in ponds and lakes with fish predators (Hecnar, 1997; Hecnar and M’Closkey, 1997b; Babbitt et al., 2003). The larvae may be somewhat unpalatable to certain predators, such as some species of sunfish (Lepomis) (Kats et al., 1988; Szuroczki and Richardson, 2011), although larvae are eaten by Rusty Crayfish, Bluegill Sunfish, and Grass Carp ­under experimental conditions (Ade et al., 2010). Eggs are eaten by 4 species of ostracods (Gray et al., 2010). POPULATION BIOLOGY

Juvenile Green Frogs grow rapidly following metamorphosis, regardless of ­whether it occurs early or late in the season. In Michigan, the highest growth rates occur in midsummer, when juveniles grow from 0.17 to 0.29 mm per day. Growth rates in spring and autumn are much less, from 0.03 to 0.09 mm per day (Martof, 1956a). Watkins and McPeek (2006) found that growth rates, swimming burst speeds, and size varied among the sites (populations) they monitored in Vermont, and that ­these ­were heritable traits. Growth rates ­were not correlated with any mea­sure of predator avoidance or escape ability. Some adult frogs appear to stop growing before ­others, so that 2 frogs of the same age may be of very dif­fer­ent sizes. As might be expected, growth rates are fastest for frogs in their

Adult Lithobates clamitans, Bronze phenotype. Alachua County, Florida. Photo: C.K. Dodd, Jr.

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Xanthic subadult Lithobates clamitans. Alachua County, Florida. Photo: C.K. Dodd, Jr.

first summer ­after metamorphosis (mean 33.6 mm/year), declining to 17.8 mm/year in the 60–70 mm adult size class. Growth still occurs at sizes greater than 70 mm SUL, but it decreases to only 2.1 mm/year in frogs greater than 90 mm SUL (all data for Michigan population; Martof, 1956a). Martof (1956a) concluded that it would take a Michigan Green Frog 4–5 yrs following metamorphosis to reach its maximum size (103 mm in males, 105 mm in females). Growth also occurs rapidly in New York Green Frogs. Examples include 40 to 78 mm in 13 months, 44 to 74 mm in 12 months, 40 to 70 mm in 5 months, and 40 to 57 mm in 3 months (Ryan, 1953). Growth patterns appear similar, despite the geographic differences. In Ontario, growth rates vary by age between males and females, with females at age 4 growing at significantly greater rates than males (Shirose and Brooks, 1995b). As above, the fastest growth rates occur 0–1 yr post-­metamorphosis. Growth rates ­were initially faster than sympatric L. septentrionalis, but t­ here ­were no differences among ­later year-­ classes between ­these species. ­There is no difference between the sexes in asymptotic size or growth constant. Shirose and Brooks (1995b) estimated that female Green Frogs reach maturity at 2 yrs post-­metamorphosis at a mean standard length of 69.4 mm SUL in Ontario, whereas males reach maturity at 3 yrs post-­metamorphosis at a mean

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standard length of 71.7 mm SUL. Most males in Michigan attained sexual maturity prior to 70 mm SUL (perhaps as small as 58.6 mm); the smallest female with eggs mea­sured 65.7 mm (Martof, 1956a). In New York, the smallest calling male was 63 mm, whereas the smallest female with eggs was 71 mm (Wells, 1977b). Pennsylvania males reached sexual maturity at 42.7 mm SUL 2 months ­after metamorphosis, and reached mean adult body size (62.8 mm) 10–11 months post-­metamorphosis; females reached sexual maturity at 57 mm SUL 11 months ­after metamorphosis, and reached mean adult body size (71 mm) 13–14 months post-­ metamorphosis (Meshaka, 2013b). Meshaka et al. (2011c) estimated that Texas females reached maturity at 50.6 mm SUL 5 months following transformation; the mean body size of sexually mature females (68.2 mm SUL) was reached 5–6 months following attainment of sexual maturity. Another estimate was 52 mm for males and 58 mm for females (Wright and Wright, 1949). Since females require a full year for the maturation of eggs, at least in Michigan (Martof, 1956b), females do not oviposit the first year of their maturity. In most other populations, maturation occurs the first year ­after transformation, but as the above indicates, ­there is much regional variation among populations and latitudes. In the Deep South, Bronze Frogs appear to have a rather dif­fer­ent life cycle. Meshaka et al. (2009a) recorded mature females at 56.7 mm 7 months ­after metamorphosis in northern Louisiana, and suggested sexual maturity was reached ­after only 4 months at a SUL of 45.2 mm. In southern Louisiana and Texas, Meshaka et al. (2009b, 2011c) estimated that maturity in males occurs ­after only 3–4 months at 39.9–44 mm SUL, and that they attain their mean adult body size (56.8–63 mm SUL) ­after only another 3–4 months. The smallest females in southern Louisiana reached maturity at 4 months at 43.1 mm SUL and attained their mean adult body size (59.8 mm SUL) ­after only 2 or 3 more months. The age structure of Green Frogs is not stable among years at Algonquin Park in Ontario. Maximum longevity was estimated at 4–6 yrs, with a mean life expectancy of 1.2–2.3 yrs post-­metamorphosis (Shirose and Brooks, 1995a). Mortality is very high for young individuals, then declines at an age of 2–3 yrs. COMMUNITY ECOLOGY

When calling, Lithobates virgatipes and L. clamitans form mixed-­species aggregations along a shoreline. ­There is a slight degree of habitat partitioning when calling, with L. clamitans calling from vegetation at the shoreline and L. virgatipes calling from locations 0.1–0.5 m into the wetland (Given, 1990). Male L. clamitans are capable of displacing male L. virgatipes from calling sites. When choruses are mixed, L. virgatipes tend to associate with conspecifics rather than

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the larger and heavier L. clamitans; L. clamitans shows no such preference. Florida L. clamitans and the closely related L. okaloosae also tend to be clumped in distribution, showing a positive degree of interaction. Both species selected similar calling positions and appeared to exclude neither conspecifics nor heterospecifics; interfrog distances ranged from a mean of 6.5–9.2 m (Gorman et al., 2009). Adult Green Frogs, Northern Leopard Frogs (L. pipiens), Mink Frogs (L. septentrionalis), and American Bullfrogs (L. catesbeianus) are often found occupying the same general nonbreeding habitat along the shorelines of larger lakes and ponds. ­These 4 species tend to select slightly dif­fer­ent microhabitats, presumably ­because of competition, overlap in prey size and composition, and the threat of predation by the large bullfrogs on the smaller ranids, particularly the subadults (Courtois et al., 1995). For example, Green Frog diets in New York overlapped by 63% with Mink Frogs and 24% with American Bullfrogs (Stewart and Sandison, 1972). Green Frogs likely compete with Northern Leopard Frogs and to a lesser extent with American Bullfrogs for prey. In turn, Green Frogs are preyed upon by the larger ranid. For example, Northern Green Frogs increased fourfold in an Ontario park ­after bullfrogs became extinct, suggesting that ­either predation by or competition with bullfrogs had been keeping populations of L. clamitans in check (Hecnar and M’Closkey, 1997c). In aquatic situations where they overlap spatially, Green Frogs occupy less dense vegetation than Northern Leopard Frogs, and they occupy microhabitats closer to shore, of lower ­water temperature, and in areas with a higher vegetation canopy than American Bullfrogs. American Bullfrogs breed in the more open ­water, whereas Green Frogs breed closer to the shoreline. Green Frogs also select areas that are closer to the shoreline (allowing rapid escape) and less vegetated than Northern Leopard Frogs (McAlpine and Dilworth, 1989), when on land. Mink Frogs, however, are usually far off in deeper ­waters associated with floating lily pads. Thus, the 4 species partition open ­water and lake shorelines allowing for coexistence despite overlap in diet and habitat occupancy. Since adults of ­these species occupy the same lakes or ponds, their larvae often overlap spatially within aquatic habitats. Both Green Frog and bullfrog tadpoles may overwinter and, as a result, vari­ous size classes of both species often are pre­sent at the same time. In experimental ­trials, competition is strong between the larvae of ­these species, except that neither small nor large Green Frog tadpoles have any effect on small American Bullfrog tadpoles (Werner, 1994). In general, American Bullfrogs are better competitors ­because of their greater activity levels; they have significant effects on resources as large individuals, as well as

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significant impacts as smaller larvae ­because of their ­great overall biomass. Green Frog larvae have the ability to respond morphologically to predators, at least in experimental ­trials. When raised with mudminnows (Umbra), they have shorter and narrower tail muscles and wider bodies than they do in the absence of ­these fish; with dragonfly larvae (Anax), they also had shorter tails and wider bodies; with salamanders (Ambystoma), they had shorter and shallower tails and wider and longer bodies; and with predaceous ­water bugs (Belostoma), they had shorter and shallower tails and deeper and wider bodies (Relyea, 2001b). ­These phenotypic responses to dif­fer­ent predators suggest that tadpoles respond to perceived threats by altering locomotor and/or vulnerability traits (e.g., by altering the predator’s ability to ­handle the prey or the extent of tail fin exposed to attack). ­Whether ­these responses occur in nature, and ­whether they actually confer advantages in survivorship in mixed-­species assemblages, has yet to be demonstrated. In general, Green Frog tadpoles do not respond to overall predation risk, but to predator-­specific situations (Relyea, 2001a). Larval be­hav­ior also changes in the presence of aquatic predators (Anax, Belostoma, Orconectes, Notophthalmus) (e.g., Hanlon and Relyea, 2013). In laboratory ­trials, Green Frog larvae greatly reduced their activity and shifted the area occupied away from the caged predator (Werner, 1991; Relyea and Werner, 1999; Peacor and Werner, 2000; Fraker, 2010). This response reduced the growth rate of the tadpole and resulted in a much smaller animal than the American Bullfrog larvae, whose size and growth rate ­were comparable in the absence of the predator (Werner, 1991). A reduced-­ activity response is not equal among predators, however. Hanlon and Relyea (2013) found that the weakest response occurred to ­water bugs (Belostoma), whereas the strongest response occurred to dragonfly nymphs (Anax). Larval survivorship also varies depending on temperature and species of dragonfly naiad when they are raised together in mesocosm experiments (Eck et al., 2014). Other ­factors may influence tadpole activity—­not just the presence of a predator, but ­whether the predator-­larvae interaction occurs in vegetated or unvegetated habitats, and ­whether pesticides are pre­sent. In mesocosms, Green Frog larvae reduce activity (feeding and swimming) in the presence of crayfish as long as vegetation is pre­sent, but not when vegetation is absent (Davis et al., 2012). Instead, they spend more time at the ­water’s surface away from where crayfish are foraging. The presence of atrazine did not alter larval susceptibility to predation or change the impact of the presence of vegetation. Of course, predators with dif­fer­ent foraging strategies, such as swimming at midwater or near the surface, likely would have altered the results of the

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experiment, which suggests the effects of vegetative structure on larval survival is context driven. Tadpole size and density also make a difference in the reduced-­activity response. The presence of the caged predator Anax reduced the activity of small Green Frog larvae but not large larvae, at low densities but not at high densities (Peacor and Werner, 2000). Large larvae presumably could avoid or reduce predation chances by size alone, regardless of density. Small larvae, however, had better chances of surviving when they ­were in a dense group of conspecifics, and thus did not change their activity levels. Small larvae at low densities would be particularly vulnerable to predation, and thus reduced their activity when predator odors ­were detected. In response to bluegill sunfish (Lepomis macrochirus), no reduction in activity or spatial avoidance occurred in experimental ­trials, and only moderate changes in activity and avoidance occurred in the presence of mudminnows (Umbra) (Relyea and Werner, 1999). Taken together, ­these results suggested that predator avoidance and activity responses in Green Frog larvae are predator, stage, and density specific. As noted above, larvae frequently reduce their activity levels when predator cues are pre­sent in experimental mesocosms. For example, Haislip et al. (2012) reported reduced activity in the presence of odonate and ­giant ­water bug cues. Smith et al. (2010) also noted that larvae responded similarly (by reduced activity) ­whether predator cues (from odonate larvae, mosquitofish [Gambusia]) ­were pre­sent singly or in mixed assemblages. Based on sibship experiments, ­these latter authors suggested that variation in response to predators might be genet­ically based. In par­tic­u­ lar, variation in larval reaction occurred to nonnative Gambusia and suggested an evolutionary response to an invasive species. As regards many such experiments, the context of the predation experiment becomes impor­tant in interpreting the results. In addition to the interactions described above, larvae interact with parasites as well as predators in natu­ral situations, and both affect larval survival. In mesocosm experiments, caged and uncaged predators (Anax) and parasites reduced larval survivorship, especially resulting in negative synergistic effects in ­trials featuring parasites and uncaged predators. It seems larvae are more active when exposed to parasites, which in turn makes them more susceptible to predation (Marino and Werner, 2013). Parasite infection rates did not differ among predator treatments. In addition to larval responses to predators, ­there is some evidence that eggs respond to chemical cues of predators. In laboratory experiments, eggs exposed in­de­pen­dently to ­water from an egg predator (crayfish, Procambarus) and a tadpole predator (Anax naiad) had a decreased time to

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hatching compared with eggs reared in predator-­free ­water. ­ hese results suggest that the eggs respond to a generalized T predator cue, rather than to a life stage–­specific predator cue (Anderson and Brown, 2009). Hatching success also decreases in the presence of crayfish, even though ­there is no direct contact. DISEASES, PARASITES, AND MALFORMATIONS

The bacterial disease “red-­leg” (Aeromonas hydrophila) has been reported to kill Green Frogs in Michigan (Martof, 1956b) and Québec (Bonin et al., 1997b). Another bacterium found on this species is Janthinobacterium lividum, which produces a potent antifungal metabolite that might contribute to an animal’s immune response to Bd (Standish et al., 2018). Frog Virus 3 (FV3) has been found in Green Frogs in Pennsylvania (Julian et al., 2019), and in ponds with access by ­cattle elsewhere (Gray et al., 2007a; Schmutzer et al., 2008). Ranavirus has been recorded from L. clamitans in Florida (Karwacki et al., 2021), Indiana (Kimble et al., 2015; Hoverman et al., 2019), Maine, Minnesota, New Jersey (Monsen-­ Collar et al., 2013), New York (Green et al., 2002; Gahl, 2007; Brunner et al., 2011; Wolff et al., 2012; Titus and Green, 2013; Youker-­Smith et al., 2016), North Carolina (Lentz et al., 2021), Ohio (Homan et al., 2013; Krynak and Dennis, 2014; Combs et al., 2015), Oklahoma (Davis et al., 2019), Prince Edward Island (Forzán and Wood, 2013), and Tennessee (O’Bryan et al., 2012), but Winzeler et al. (2016) did not detect it at 5 sites in Indiana. ­Water temperature appears to play a substantial role in the pathogenicity of FV3-­like viruses in Green Frogs, with greater mortality at higher temperatures (25°C) than at lower temperatures (10°C) (Brand et al., 2016). Although ranavirus ­causes substantial mortality in larvae in experimental ­trials, it does not act synergistically with predation to affect survivorship (Haislip et al., 2012). Larval Green Frogs are far more sensitive to ranavirus than FV3, and experience significant mortality from both ranavirus and FV3 infection (Hoverman et al., 2011). An Ichthyophonus-­like fungus was reported from a Green Frog in ­Virginia (Gibson and Sattler, 2020). Fungal Ichthyophonus-­like spores ­were found on 17 of 83 Green Frogs in Québec; ­these spores caused mild to severe myositis (Mikaelian et al., 2000). The significance and prevalence of this disease and its long-­term effects are unknown. A Dermocystidium-­like fungus is reported from Green Frogs in Minnesota (Green et al., 2002). Perkinsea, an alveolate pathogen, also has been found on museum specimens from Florida (Karwacki et al., 2021). Green Frogs are infected frequently by Batrachochytrium dendrobatidis (Bd). In Maine, for example, 17.2% of the frogs examined (34 of 197) ­were infected in the toe webbing, 8.3% (3 of 36) on the skin of the tibia, and 12.8% (22 of

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172) on the skin of the pelvis (Longcore et al., 2007; see also Gahl, 2007), whereas in Québec prevalence was 20% for 718 individuals collected from 1900 to 1999 (Ouellet et al., 2005a). In New York, 33% of Green Frogs ­were infected (Windstam and Olori, 2014). The earliest recorded case was from a Green Frog collected in 1961 at Saint-­Pierre-­de-­ Wakefield. Bd also occurs on Green Frogs from the Upper Midwest, Alabama, Connecticut, Florida, Georgia, Illinois, Indiana, Maine, Mary­land, Mas­sa­chu­setts, Michigan, Minnesota, New Brunswick, New Hampshire, New Jersey, Adirondack Park in New York, North Carolina, Ohio, Oklahoma, Pennsylvania, Tennessee, ­Virginia, and Wisconsin (Timpe et al., 2008; Sadinski et al., 2010; Hill et al., 2011; Krynak et al., 2012; Wolff et al., 2012; Hanlon et al., 2014b; Hughey et al., 2014; Igleski and Nicholson, 2014; Phillips et al., 2014; Wolff et al., 2014; Wilson et al., 2015; Sacerdote-­ Velat et al., 2016; Chiari et al., 2017; Tupper et al., 2011, 2017; Klemish et al., 2012; Marhanka et al., 2017; Fuchs et al., 2018; Robinson et al., 2018; Watters et al., 2018; Jongsma et al., 2019; Julian et al., 2019; Siddons et al., 2020; Karwacki et al., 2021; Lentz et al., 2021). Gahl et al. (2011) showed that the northeastern strain of Bd did not kill metamorphic Green Frogs, although other strains did. It has also been found on Green Frogs from Vermont (Green et al., 2002), Maine (Gahl, 2007), and New Jersey (Monsen-­Collar et al., 2010). Bd-­infected tadpoles show a decreased ability to obtain food, and decreased feeding per­for­mance is correlated with the intensity of Bd infection (DeMarchi et al., 2015). Green Frogs are parasitized by >13 species of nematodes, 36 species of trematodes (of which 12 affect larvae), 3 cestodes, a copepod on tadpoles, mites, 3 species of flies, and 32 species of protozoans (Walton, 1947; McAlpine, 1997b; McAlpine and Burt, 1998; Raffel et al., 2011). The coccidian parasite Eimeria menaensis was described from Green Frogs in Arkansas (McAllister et al., 2014a). Intraerythrocytic parasites include Aegyptianella ranarum, Hepatozoon sp., H. clamatae, Lankestterella minima, Trypanosoma rotatorium, and T. ranarum (Bollinger et al., 1968; Bonin et al., 1997b; Trites et al., 2013), and viruses (Desser and Barta, 1984a; Gruia-­Gray et al., 1989). A pathogenic protozoan (Brugerolleia algonquinensis) was described from Green Frogs in Canada (Desser and Jones, 1985; Desser et al., 1993); the ciliate protozoan Nyctotherus cordiformis was reported from Ohio and ­Virginia Green Frogs (Odlaug, 1954; Campbell, 1968); other protozoans include Entamoeba ranarum, Haematogregarina sp., Hexamita intestinalis, and Tritrichomonas augusta (Campbell, 1968). The myxozoan Myxidium serotinum also parasitizes L. clamitans (McAllister et al., 2008). A virulent alveolate pathogen affecting larval Green Frogs was reported from Maine (Gahl, 2007).

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Trematode cysts are common in the kidneys (Martin and Conn, 1990). Species include Alaria arisaemoides, Apharyngostrigea pipientis, Cephalogonimus americanus, C. vesicaudus, Clinostomum attenuatum, Echinostoma trivolvis, Glypthelmins quieta, Gorgodera amplicava, Gorgoderina attenuata, G. minima, G. simplex, G. subtropica, G. tanneri, G. translucida, Gyrodactylus sp. (on larvae), Haematoloechus breviplexus, H. longiplexus, H. parviplexus, H. similiplexus, H. varioplexus, Haematoloechus sp., Halipegus eccentricus, H. occidualis, Hichinastomia trivolvis, Loxogenes arcanum, Loxogenoides bicolor, Megalodiscus temperatus, Pneumobitis parviplexus, and Pneumonoeces parviplexus (Irwin, 1929; Bouchard, 1951; Odlaug, 1954; Najarian, 1955; Campbell, 1968; Brooks, 1979; Goater et al., 1990; Dyer, 1991; Wetzel and Esch, 1996; McAlpine, 1997b; Bursey and DeWolf, 1998; McAlpine and Burt, 1998; Zelmer et al., 1999; Green and Dodd, 2007). Tadpoles appear most susceptible to trematode infections in mid-­ season as opposed to early and ­later in the season (Raffel et al., 2011). The cestode Ophiotaenia saphena is common in Green Frogs (Sutherland, 2005), as are Cylindrotaenia americana, Mesocestoides sp., Ophiotaenia gracilis, O. saphenus, and Proteocephalus saphena (Williams and Taft, 1980; Dyer, 1991; McAlpine, 1997b; Bursey and DeWolf, 1998; Mc­ Alpine and Burt, 1998). Nematodes include Cosmocercoides dukae, C. variabilis, Falcaustra inglisi, Foleyella americana, Gyrinicola batrachiensis, Oxysomatium variabilis, Ozwaldocruzia pipiens, O. waltoni, Physaloptera sp., P. ranae, Rhabdias ranae, and Spiroxys contortus (Odlaug, 1954; Campbell, 1968; Adamson, 1981c; Dyer, 1991; McAlpine, 1997b; Bursey and DeWolf, 1998; McAlpine and Burt, 1998; Pryor and Greiner, 2004). Gilhen (1984) noted the presence of unidentified nematodes in the stomach. Acanthocephalan larvae are reported from the mesenteries of the body cavity (Odlaug, 1954), and Campbell (1968) recorded the species Centrorhynchus sp. and Centrorhynchus wardae from ­Virginia. The mite Hannemania penetrans is known from this species in ­Virginia (Campbell, 1968). Martof (1956b) observed that 80% of the male Green Frogs at Michigan ponds ­were infested by leeches, especially on the webs of the hind feet, and leeches (Placobdella [=Desserobdella] picta) are found commonly on Canadian and Arkansan Green Frogs (Barta and Desser, 1984; McCallum et al., 2011b). The parasitic copepod Lernaea cyprinacea has been reported widely from Green Frogs (Watermolen, 2019). Myiasis (dermal fly infestation) has also been reported in this species (Hughes, 2014). Green Frogs with malformations, including hind feet of abnormal shape or proportion, missing feet or partial limbs

Dodd_Canada_int_5pgs_B3.indd 501

(ectromelia), missing front legs, missing digits (ectrodactyly), missing eyes, multiple limbs, jaw deformities, eye discoloration, and even a missing tympanum, have been found in many locations (Martof, 1956b; Bonin et al., 1997b; Ouellet et al., 1997; Converse et al., 2000; Helgen et al., 2000; Gillilland et al., 2001; Hoppe, 2005; Lannoo, 2008; Terrell et al., 2014a). A photo­graph of a L. clamitans with multiple forelimbs is in Meteyer (2000). In Minnesota, 80 of 219 frogs examined had some type of malformation (Hoppe, 2005). Most malformations affected the hind limbs at Martof’s study site in Michigan, and 23 of 428 (5.3%) frogs had missing body parts or exhibited abnormal development. Most injuries, however, affected digits or limbs and appeared to represent unsuccessful predation attempts. The Rock Bass (Ambloplites rupestris) prob­ably was responsible for many of the predation attempts (Martof, 1956b). In Québec, malformations and abnormal DNA profiles ­were associated with agricultural activity, and agricultural contaminants (especially carbofuran) ­were suspected as the cause (Bonin et al., 1997b; Lowcock et al., 1997; Ouellet et al., 1997). The levels and types of abnormalities ­were consistent with acute or cumulative pesticide toxicity. Exposure to coal ash contaminated wastewater also results in larval malformations (Snodgrass et al., 2004). In addition to malformations, Green Frogs infected with ranavirus show higher levels of fluctuating asymmetry than uninfected frogs, suggesting that diseases can have sublethal effects on development (St.-­Amour et al., 2010). Frogs infected by Bd ­were not asymmetrical. In addition to external malformations, developmental anomalies are found in Green Frogs from urban and residential suburban areas in higher frequencies than in natu­ral populations. Such malformations involve the development of intersexes in males, with males from the Connecticut River Valley having gonads containing testicular oocytes. Up to 21% of male Green Frogs from suburban landscapes had abnormal testicular development; frogs from nearby agricultural areas had low levels of abnormalities (Skelly et al., 2010). In a ­later study at 16 ponds, abnormality frequencies (in males with oocytes) ranged from 6 to 58%, with no significant differences in neighborhoods served by septic or sewer systems, or neither (Smits et al., 2014). It seems reasonable to hypothesize that gonadal abnormalities are associated with the high levels of pesticides used in residential areas and the presence of other endocrine disrupting compounds. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. In a study of lead in tadpoles and their habitats near highways, lead concentrations ranged between 0.90 and 240 mg kg-1. Lead concentrations in tadpoles ­were positively

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502  Ranidae

correlated with the amount of lead in sediments and mean daily traffic volume. It is not known ­whether ­these concentrations adversely affect Green Frog larvae, although similar concentrations have been associated with adverse physiological and reproductive effects in other vertebrates (Birdsall et al., 1986). In addition, cadmium (0.1–0.19 ppm wet weight), copper (0.93–1.2 ppm), lead (1.4–1.5 ppm), mercury (0.04–0.1 ppm), zinc (3.7–6 ppm), magnesium (14–29 ppm), and manganese (1.1 ppm) have been reported from Green Frog tadpoles (Hall and Mulhern, 1984). Mercury has been found in Green Frog tadpoles in natu­ral ponds in Maine, with a mean of 25.1 ng/g; methyl mercury comprised 7.6–40% of the total (Bank et al., 2007). In a series of mine-­restored and seminatural areas in Ohio, metal concentrations (means in ppm) ranged from 41 to 113 for manganese, 289–423 for iron, 59–90 for aluminum, and 91–112 for zinc (Lacki et al., 1992). The presence of Green Frogs in Ontario is negatively correlated with aluminum concentrations (Glooschenko et al., 1992), and frog abundance was negatively associated with mining activities with regard to metal contamination (Sasaki et al., 2015). Although pre­sent at mine tailing sites in Sudbury, Ontario, biomass and abundance was decreased in comparison with unaffected sites (Leduc et al., 2012). Sublethal exposure to lead inhibits both acquisition and retention of discriminate avoidance learning in Green Frog tadpoles (Strickler-­Shaw and Taylor, 1990; Steele et at., 1999). In experimental ­trials, tadpoles do not avoid lead and lead has no effects on gross tadpole locomotor per­for­mance (Taylor et al., 1990). Likewise, pre-­exposure to lead did not increase sensitivity to subsequent encounters with lead. Instead, tadpole spontaneous activity increased in variation as a function of lead exposure. Lead accumulates in tadpole brains and produces neural damage (Strickler-­Shaw, 1988). ­Under laboratory conditions, Green Frog larvae accumulate heavy metals (arsenic, cadmium, iron, selenium, strontium, and vanadium) when raised in ­water contaminated with coal combustion wastes. Exposure to polluted ­water decreased growth and developmental rates, reduced survivorship by 26%, and decreased metamorphic success by 45%. Furthermore, the duration of the larval period increased, and the size at metamorphosis decreased by 10% (Snodgrass et al., 2004). ­These results suggest severe effects on amphibian larval development in ­water polluted by coal ash power plants. pH. Higher pH values are positively associated with Green Frog abundance (Mazerolle, 2003). The lethal pH for Green Frog embryos is 3.7–3.8, with a critical pH of 4.1 (Gosner and Black, 1957a; Freda and Dunson, 1986; Freda et al., 1991). Larval Green Frogs are generally unable to acclimate to low pH (≤5.0). Tadpoles are unable to initially regulate

Dodd_Canada_int_5pgs_B3.indd 502

ionic (Na+, Cl-­, Ca2+, K+) losses at low pH (Freda and Dunson, 1984), but they slowly recover ­after 7 hrs; juveniles do not, however, and sodium uptake is reduced by 60%. Chronic ion losses occur primarily in the gills of developing larvae. Thus, chronic acid exposure does not allow larvae to adapt to low pH (McDonald et al., 1984). ­There may be some regional variation in pH tolerance, as adults have been found in Nova Scotia in ­waters of pH 3.5, and larvae at pH 3.9 (Dale et al., 1985); most frogs ­were found in ­waters with higher pH content (3.88–7.26, mean 4.95). Freda and Dunson (1984) stated that Green Frog larvae can live for “several weeks” in ­water at pH 3.5. In experimental ­trials, Green Frog larvae are able to detect and move away from ­water that is very acidic (Freda and Taylor, 1992; Vatnick et al., 1999). Green Frogs do not appear to be affected by soil type in experimental situations, although the chemical properties of soils may affect acidity within breeding ponds (Sparling et al., 1995). The presence of Northern Green Frogs increases in ponds with a high buffering status; buffering status is inversely correlated with acidity (Glooschenko et al., 1992). Alkalinity. The presence of Northern Green Frogs is positively correlated with alkalinity concentrations in Ontario (Glooschenko et al., 1992). Ammonia and Nitrates. As might be expected, larvae metamorphosing from wetlands with high nitrate content have higher concentrations of nitrates than larvae metamorphosing from wetlands with low nitrate content (Jefferson and Russell, 2008). Green Frog larvae are sensitive to concentrations of >30 mg/L ammonium nitrate in acute exposure ­trials; the LC50 (96 hr) is 32.4 mg/L. Tadpoles exposed to higher concentrations of nitrate generally become sluggish, and they may forgo feeding. They do not appear to be sensitive to chronic exposure at exposures 0.5 m in dia­meter that possess a soil or gravel-­lined substrate with ambient light penetration. They prefer short tunnels over long tunnels. Fences 0.6– 0.9 m in height also prevented trespass (Woltz et al., 2008). Traffic noise does not inhibit calling by this species. When Green Frogs call in the vicinity of roads with low volumes of traffic noise, their call rates are significantly lower than when they are in the presence of high volumes of traffic noise. Call amplitudes are lower and dominant frequencies are higher when noise volumes are high (Cunnington and Fahrig, 2010). Traffic noise also elicits an immediate response in calling frogs by decreasing call rates and amplitude and by increasing call frequencies compared with calls immediately before exposure to traffic noise. However, traffic noise does not interfere with mate attraction in this species (Cunnington and Fahrig, 2013). This plasticity in call characteristics allows the frogs to communicate acoustically despite the potential interference from traffic noise. Roads are used extensively to monitor Green Frog populations based on call surveys over prescribed routes at regular intervals (Bishop et al., 1997; Bonin et al., 1997a; Lepage et al., 1997; Mossman et al., 1998; Sargent, 2000; Nelson and Graves, 2004). The probability of hearing Green Frogs during an aural survey is usually good, assuming individuals are calling that night, but this might be inversely proportional to traffic volume since Vargas-­Salinas et al. (2014) found that Green Frogs, a species with a low call peak frequency, called more often when traffic noise was low. Also in Ontario, de Solla et al. (2005) estimated that one could detect Green Frogs using aural surveys on only 3 sampling nights and still be 80% certain of detecting the species’ presence within the area of interest. Still, ­there is some degree of annual variation in the frequency with which Green Frogs are heard along standardized calling survey routes (Bishop et al., 1997). Green Frog abundance is correlated with the number of calls per minute, and abundance is positively correlated with calling index; ­these results suggest that calling index values are useful mea­sures of abundance during road survey transects (Nelson and Graves, 2004). Green Frogs are able to use wetlands found ­under power-­line rights-­of-­way, as long as forested habitat abuts the transmission corridor (Fortin et al., 2004b). They may colonize created or restored wetlands, as long as ­there are source populations nearby (Lacki et al., 1992; Briggler, 1998;

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Stevens et al., 2002; Touré and Middendorf, 2002; Weyrauch and Amon, 2002; Brodman et al., 2006; Terrell et al., 2014a; Walls et al., 2014a; Drayer and Richter, 2016; Kross and Richter, 2016; Mitchell, 2016; Ecrement and Richter, 2017; Stiles et al., 2017a). For example, Bronze Frogs breed in shallow-­sloped fishless tank defilades on military bases (Ecrement and Richter, 2017). Unfortunately, amphibian pathogens may be much more prevalent in restored wetlands compared with natu­ral wetlands and adversely impact this species (Julian et al., 2019). In contrast, Lehtinen and Galatowitsch (2001) did not find L. clamitans in ponds restored up to 20 months previously, although the restored ponds appeared to be in highly disturbed areas, including wetlands that had not held ­water for 50 yrs. Palis (2007) also recorded only small numbers of Green Frogs colonizing three restored wetlands in southern Illinois, as did Pechmann et al. (2001) at four ponds in South Carolina. Merovich and Howard (2000) found that it took Green Frogs at least 4–5 yrs to colonize small constructed ponds, and even then, frogs ­were not abundant. Taken together, ­these results suggest mixed effects of wetland restoration on this species. It seems likely that Green Frogs are more likely to become established at larger and more permanent created wetlands than at small ponds with varying hydroperiods. Large size, deep ­water, and permanent hydroperiod are critical ­factors in ­whether this species ­will use created wetlands (Denton and Richter, 2013). The success or failure of Green Frogs to colonize small restored wetlands also undoubtedly depends on location within the landscape in relation to source wetlands and nearby breeding populations, as well as on the viability of populations that had been displaced. In some cases, it may be necessary to repatriate adults or larvae to establish a population. Repatriation has proven unsuccessful at the Gateway National Recreation Area in New York (Cook, 2008), despite initial indications of success (Cook, 1989). Playing conspecific chorus sounds does not increase the likelihood of colonizing artificial ponds (Buxton et al., 2018). Green Frog larvae may be moved throughout a landscape through purposeful or inadvertent anthropogenic mediation in connection with sport fish stocking. The Ohio Division of Conservation stocked Green Frog tadpoles obtained in fish hatcheries throughout the state (Walker, 1946), and this undoubtedly has occurred elsewhere. In terms of economic importance, Green Frogs have been a minor source of commercial frog’s legs, and they have been harvested by biological supply ­houses for use in laboratories. A total of 32 L. clamitans ­were reported commercially collected for the pet trade in Florida from 1990 to 1994 (Enge, 2005a). This species is considered Threatened in Kansas.

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508  Ranidae

Lithobates fisheri (Stejneger, 1893) Mogollon Rim Leopard Frog ETYMOLOGY

fisheri: a patronym honoring Dr. A.K. Fisher (1856–1948), “in recognition of his share in the herpetological success of the Death Valley Expedition.” Fisher was a medical doctor and ornithologist on the expedition, and likely captured many herpetological specimens. NOMENCLATURE

Stebbins (2003): Rana onca Dubois (2006): Lithobates (Lithobates) fisheri Fouquette and Dubois (2014): Rana (Lithobates) fisheri Synonyms: Lithobates chiricahuensis (in part), Rana onca fisheri, Rana pipiens fisheri The common name of this species was the Vegas Valley Leopard Frog when the species was known only from that vicinity. Since the species may occur beyond the Vegas Valley, the name Mogollon Rim Leopard Frog, suggested by Jim Rorabaugh, is used as the common name for this frog. Information on L. fisheri populations along Arizona’s central Mogollon Rim and the San Francisco Mountains on the Arizona–­New Mexico border may be included in the lit­er­a­ture on L. chiricahuensis (e.g., Rorabaugh and Sredl, 2014).

Additional specimens in collections are described by Wright and Wright (1949). The only known photo­graphs of living individuals from the Vegas Valley are in Tanner (1931), Wright and Wright (1949), and Jennings and Hayes (1994a). Larvae. Wright (1929) and Wright and Wright (1949) described the colorful tadpole as buffy olive to “dull citrine” with pale, greenish-­yellow clusters over it and a greenish-­ yellow tail that was heavi­ly mottled. The venter is semitransparent, pure white to a faint cinnamon color. The tail is elongate with a rounded tip, “oil yellow” in color, and the TL reaches ca. 85 mm. Tadpoles have small black spots or mottling on the tail. Wright (1929) figured the mouthparts and has a lateral black-­and-­white photo­graph of a live tadpole. Eggs. No information is available, but presumably they are similar to other members of the leopard frog complex. DISTRIBUTION

­ ntil recent ge­ne­tic analy­sis of museum collections (Hekkala U et al., 2011), this species was known only from the Vegas Valley in southern Nevada. Collections ­were made west of the then small town (Wright and Wright, 1949), about 1.6 km north of the town (Slevin, 1928), and in Tule Springs, 25.7 km north of Las Vegas (A. Vanderhorst collections; Jennings, 2005). Stejneger’s and Slevin’s collections ­were made at Las Vegas Ranch. This population is extinct. Specimens are in the collections of the Smithsonian Institution (USNM), the California Acad­emy of Science, and the Museum of Vertebrate Zoology at the University of California–­Berkeley. Hekkala

IDENTIFICATION

Adults. The dorsal coloration is grayish to olive green with numerous small, distinct, dark greenish-­olive spots (in 3–4 rows) surrounded by lighter rings. Anteriorly, males tend to lose their dorsal spots and assume a bright green coloration, and females are more spotted than males. Dark olive spots extend along the sides of the body, which are grayish olive. The tympanum is large (greater than the distance between the nostrils and eyes) with no black patch. A dorsolateral fold bordered by light stripes is pre­sent but not strongly developed; it extends only about halfway down the back, making this and L. onca distinct from other Southwestern leopard frogs. The hind legs are relatively short, and the hind toes are two-­thirds webbed. Along the front and rear of the hind limbs are many reticulations of deep olive to pale gray. The throat is light green and spotted, at least in smaller individuals, and the venter is pinkish cinnamon. Ventral coloration of the hind limbs is honey yellow to buff. No external vocal sac is apparent. Males have an enlarged thumb. Males range from 44 to 64 mm SUL and females from 46 to 74 mm SUL (Wright and Wright, 1949), suggesting sexual size dimorphism. Slevin (1928) recorded 6 individuals from 45 to 60 mm SUL (as L. onca).

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Distribution of Lithobates fisheri. The population in the Vegas Valley is extirpated.

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Lithobates fisheri 509

et al. (2011) extended the range to include Arizona’s central Mogollon Rim and the San Francisco Mountains on the Arizona–­New Mexico border. Many of ­these populations have been extirpated. FOSSIL REC­O RD

No fossils are known. Holman (2003) noted Miocene (Hemphillian) fossils from Navajo County in northern Arizona that belonged to the L. pipiens complex. The relationship between ­these fossils and L. fisheri is unknown. SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates fisheri was described by Stejneger (1893) from the Vegas Valley of southern Nevada. In the description, he noted that other leopard frogs ­were found (“tolerably common”) along Beaverdam Creek near its junction with the Virgin River, but he could not determine ­whether they ­were L. fisheri. This taxon has been variously considered as a species, as a subspecies, or as conspecific with L. onca, a taxon to which it is closely related (Hekkala et al., 2011). Hekkala et al. (2011) extracted DNA from preserved frogs collected from the Vegas Valley and concluded that L. fisheri was conspecific with populations of leopard frogs from the central Mogollon Rim and the San Francisco Mountains on the Arizona–­New Mexico border, heretofore considered L. chiricahuensis. Additional molecular research is ­under way to elucidate the relationships of the Mogollon Rim frogs with L. chiricahuensis populations in southern Arizona and México and to verify the conclusions of Hekkala et al. (2011). Lithobates fisheri is a member of the Novirana clade of North American ranid frogs. ADULT HABITAT

According to Linsdale (1940), this species inhabited artesian springs and open, sedge-­bordered short streams in the vicinity of Las Vegas at an elevation of 610 m. Wright and Wright (1949) recount a collecting trip to the Vegas Valley in 1925. Lithobates fisheri was found in a small spring outlet that was about 1.0–1.2 m across, covered in algae, and bordered by sedges. Other L. fisheri ­were found along small spring holes in areas that looked “marly and alkali,” and in a number of springheads or well holes. Most locations appeared to be rather small, based on available descriptions. The largest populations ­were located at 3 large springs at the headwaters of Las Vegas Creek, which had a flow of 9 million m3 annually. Prior to 1938, riparian vegetation along Las Vegas Creek and surrounding springs included cottonwoods, willows, bulrushes, sedges, and cattails. The creek itself was reported to be 1–6 m in width with a sand, gravel, and mud bottom. Jennings and Hayes (1994a) provided historical photos of Las Vegas Creek, Kiel’s Spring run, and another small spring in Las Vegas Valley, all taken in 1903.

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Plate XCVI. Rana (= Lithobates) fisheri. 1. Male, 2–4 Females. From Handbook of Frogs and Toads by A.D. and A.A. Wright, Third Edition, Cornell University Press, Ithaca, NY. Reprinted with permission.

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510  Ranidae

Tadpole of Lithobates fisheri. Photo: A.D. Wright collection, Kroch Library, Cornell University.

Along the Mogollon Rim, the species is found in Madrean evergreen woodland, pinyon-­juniper conifer forest, and montane conifer forest habitats. Streams and associated plunge pools in rocky canyons constitute the preferred habitat for this species. The species occurs to 2,709 m in the White Mountains (Rorabaugh and Sredl, 2014). AQUATIC AND TERRESTRIAL ECOLOGY

­ ittle is known concerning the ecol­ogy of this species. Vegas L Valley frogs ­were said to be most active in spring and early summer, with capture dates from January to mid-­August (Stebbins, 1962). It likely does not venture far from ­water, and activity may be curtailed from November to February depending on weather. The ecol­ogy of this species on the Mogollon Rim has not been reported but is prob­ably similar to populations of L. chiricahuensis in other high mountain regions to the east.

Las Vegas River. Historic habitat of Lithobates fisheri, ca. 1903. Courtesy of Mark Jennings.

CALLING ACTIVITY AND MATE SE­L ECTION

No information is available, but the be­hav­ior of L. fisheri may be similar to that of L. chiricahuensis. When handled, L. fisheri in Las Vegas made a “semicroak” or a “very low croak” (Wright and Wright, 1949). BREEDING SITES

Presumably the species bred in the marshes, meadows, springs, and short streams that once ­were pre­sent in the Vegas Valley. ­These areas may have resembled the marsh and spring systems in Ash Meadows to the west. Photo­graphs of spring and creek habitats are in Jennings and Hayes (1994a), but even by the time the photos ­were made, the habitats had already been highly modified. Breeding habitats along the Mogollon Rim include stock tanks, ponds, and along slow-­moving streams and pools.

1949) found transforming froglets of 30–35 mm, some with remaining tail nubs, on 1 May 1913. Tadpoles also ­were observed on 20 August 1925 (Wright and Wright, 1949). If the species bred in late March and April, transformation likely occurred in summer. However, the observation of small frogs (ca. 30 mm SUL) in April and early May (reviewed by Wright and Wright, 1949) suggests a long larval period (perhaps even overwintering as larvae), ­little growth by metamorphs from summer transformation ­until the following spring (unlikely), or a rather extended breeding and larval period. Jennings (2005) suggested that metamorphosis could have occurred throughout much of the year. DIET

REPRODUCTION

Presumably, this species eats a variety of invertebrates found in the vicinity of springs, ponds, stock tanks, and marshes.

Lithobates fisheri prob­ably breeds in spring (February–­April), but no other information is available.

PREDATION AND DEFENSE

LARVAL ECOLOGY

Linsdale (in Wright and Wright, 1949) recorded transformation sizes of 28–30.5 mm. Slevin (in Wright and Wright,

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Wright and Wright (1949) noted that L. fisheri jumped into the ­water when approached, but rested on the bottom and made no attempt to bury into the substrate. The species was easily captured.

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Lithobates fisheri 511

Adult Lithobates fisheri. Coconino County, Arizona. Photo: Jim Rorabaugh

POPULATION AND COMMUNITY ECOLOGY

No information is available. DISEASES, PARASITES, AND MALFORMATIONS

No information is available, but parasites are prob­ably similar to ­those of L. chiricahuensis. SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available.

Spring in the Las Vegas Valley, ca. 1903. Historic habitat of Lithobates fisheri. Courtesy of Mark Jennings.

STATUS AND CONSERVATION

This species is presumed locally extinct in the Vegas Valley due to the destruction of its habitat during the expansion of Las Vegas in the 1940s. The ­water ­table was lowered due to groundwater pumping, and springs and wells ­were capped or destroyed. J.R. Slevin in 1913, C.L. Camp in 1923, A.H. Wright in 1925, J. Linsdale in 1934 and 1936, and Stanford biologists in 1934 and 1938 ­were able to locate springs and waterholes with L. fisheri and make collections (Wright and Wright, 1949). A. Vanderhorst also collected 13 frogs from Tule Springs in January 1942 (Jennings, 2005). On a return trip in May 1942, A.H. Wright had difficulty locating sites and only heard a few frogs splash into the ­water. He noted severe habitat modification as well as the presence of American Bullfrogs (L. catesbeianus) and crayfish. By this time, 13 km long Las Vegas Creek had been heavi­ly polluted. He expressed grave concern for the survival of this species and, unfortunately, his concern proved prophetic. If it was L. fisheri that jumped into the ­water during his 1942 collecting trip, it was the last recorded observation of this species in the Vegas Valley. Robert Stebbins attempted to find this species in 1949 without success and noted the loss of virtually all

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Mogollon Rim habitat of Lithobates fisheri. Apache County, Arizona. Photo: Jim Rorabaugh

riparian vegetation by 1938. Efforts to re-­establish leopard frogs in the Vegas Valley have focused on L. onca rather than populations of “L. fisheri” (sensu Hekkala et al., 2011) from the Mogollon Rim (see L. onca account). Populations along the Mogollon Rim have apparently declined dramatically over the last de­cades (see L. chiricahuensis account).

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512  Ranidae

Lithobates grylio (Stejneger, 1901) Pig Frog ETYMOLOGY

grylio: from the Greek grylos meaning ‘pig.’ The name refers to the species’ call. NOMENCLATURE

Conant and Collins (1998): Rana grylio Dubois (2006): Lithobates (Aquarana) grylio Fouquette and Dubois (2014): Rana (Lithobates) grylio Synonyms: Rana grylio In the older lit­er­a­ture, this species is referred to as the Southern Bullfrog or as “Joe Browns.” IDENTIFICATION

Adults. This is a large, olive to dark green frog with few black spots, at least in the east. Juveniles have a light tan dorsolateral band. This band is absent in eastern adults, but it tends to persist, albeit reduced, in western adults. The Pig Frog is very similar to L. catesbeianus and often has been confused with that species. Snouts tend to be more pointed and narrower than in L. catesbeianus, but this character is not always reliable. Throats are cream to yellowish, and bellies and the undersides of legs are white or yellowish white with black mottling ­toward the groin. The toes of L. grylio are longer than ­those of L. catesbeianus, but the fourth toe is the longest. When the fourth and fifth toes are adpressed, the tip of the fifth toe should reach beyond the base of the next to the last joint of the fourth toe. If it does not, the species is L. catesbeianus (Dundee, 1974). Males often have a bright yellow throat and a tympanum larger than the eye. Males are smaller than females. In the Everglades, the largest female captured was 157 mm SUL (431 g), whereas the largest male was 131 mm SUL (228 g) (Ligas, 1960). In Ugarte’s (2004) Everglades study, the maximum female size was 130 mm SUL. Differences prob­ably reflect harvest pressure. Duellman and Schwartz (1958) recorded males 97–117 mm SUL (mean 106 mm) and females 98–136 mm SUL (mean 115 mm). Jensen et al. (2008) gave an adult range of 83–162 mm SUL, but Mount (1975) gave a maximum size of 165 mm SUL. Larvae. Mature tadpoles are olive green with dark spots throughout the body and tail. About 9.5 days ­after hatching, the initially black larvae develop prominent, brassy transverse bands dorsally. One is located anterior to the eyes, whereas the other is posterior to the eyes; the bands dis­ appear by 35 mm TL. A gold spot may be vis­i­ble at the base

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of the tail. Snout stripes are poorly developed if pre­sent at all. Tadpoles are rounded in a dorsal view, have a more pointed snout, and are less depressed than other large ranids (Altig, 1972a). Intestines are partially vis­i­ble through a heavi­ly pigmented venter. The throat is light colored. The maximum size is 100 mm TL. Eggs. Eggs are black above and white below and oviposited in a surface film about 30 cm × 30 cm or larger. The vitellus is 1.4–1.8 mm in dia­meter, the inner envelope is 2.8–4 mm in dia­meter, and the outer envelope is 3.8–7 mm in dia­meter (Wright, 1932; Livezey and Wright, 1947). DISTRIBUTION

Lithobates grylio occurs on the Atlantic Coastal Plain from southern South Carolina throughout the Florida peninsula and west to southeastern Texas. The species has also been introduced into the Bahamas (Neill, 1964), China (in the early 1980s), and Puerto Rico (Kraus, 2009). Pig Frogs have been recorded on Ossabaw, Sapelo, and Cumberland islands, Georgia (Martof, 1963; Williamson and Moulis, 1994; Shoop and Ruckdeschel, 2006). Impor­tant distributional references include: range-­wide (Wright, 1932), Alabama (Mount, 1975), Florida (Dodd et al., 2017; Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), South Carolina (Beane et al., 2010; Dodd and Barichivich, 2017), and Texas (Livezey and Johnson, 1948; Dixon, 2000, 2013; Tipton et al., 2012). FOSSIL REC­O RD

Pleistocene fossils of this species are known from deposits in Florida (Holman, 2003). The species is distinguished from

Distribution of Lithobates grylio

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Lithobates grylio 513

L. catesbeianus and L. heckscheri by differences in the maxilla and ilium. SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates grylio is a member of the American Bullfrog species group (Aquarana clade) that consists of 7 closely related species (L. catesbeianus, L. clamitans, L. grylio, L. heckscheri, L. okaloosae, L. septentrionalis, L. virgatipes) that evolved rapidly from a common ancestor during the late Miocene to early Pliocene (Austin et al., 2003a; Hillis and Wilcox, 2005). It is closely related to L. heckscheri. Duellman and Schwartz (1958) noted that south Florida Pig Frogs ­were less brown in coloration and had more ventral pigmentation than Pig Frogs from north Florida and Mississippi. However, ­there is considerable variation in ­these characters even within the same population. ADULT HABITAT

This is a species of large ­water bodies (ponds, lakes, wet prairies, freshwater marshes, Carolina Bays) throughout the southern Coastal Plain and peninsular Florida. Other habitats include sloughs, canals, sinkhole ponds, permanent ditches, ponds in pine flatwoods, and sawgrass marshes (Meshaka et al., 2000; Chandler et al., 2015a). Preferred habitats contain extensive amounts of emergent and floating vegetation in open-­canopied wetlands of varying depths. Wetlands frequently have dif­fer­ent vegetation zones, such as an emergent herb zone, a grass-­herb zone, and a cypress-­hardwood zone. Vegetation provides cover and support for foraging and basking frogs. Surrounding habitats consist of bayheads, mesic hammock, mixed deciduous forest, and upland longleaf pine sandhills. Pig Frogs may be pre­sent in flatwoods ponds, but numbers are low (Vickers et al., 1985). Pig Frogs frequently occur with many species of fish (Holbrook and Dorn, 2016). This species is the most common big frog of large wetlands such as Okefenokee Swamp and the Everglades. AQUATIC AND TERRESTRIAL ECOLOGY

Lithobates grylio is a highly aquatic species that rarely leaves the ­water. Adults can lose no more than ca. 27% of their body ­water before succumbing (Thorson and Svihla, 1943). However, individual Pig Frogs, mostly juveniles, have been found dispersing through sandhills habitats at some distance from the nearest large wetlands (Dodd, 1992; Branch and Hokit, 2000; Enge and Wood, 2001; Langford et al., 2007). ­These frogs may occupy seasonally temporary wetlands, presumably to feed rather than to breed (Babbitt and Tanner, 2000; Greenberg and Tanner, 2005b). During the nonbreeding season, male and female

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Pig Frogs inhabit ­waters along the fringes of ponds and lake margins, such as in the cypress and hardwood zones that fringe many coastal plain lakes. ­There is no relationship between winter activity and canopy cover or bank slope, but winter occurrence is negatively associated with plant species richness (Borski and Bickford, 2019). As the breeding season commences, males move out into the grass-­herb zones 40–50 m from the shoreline; females remain in the shallow ­waters (see Walkowski, 2020). The only time females move away from the shore is to mate with a male calling from the more open emergent vegetation. ­After oviposition, they return to the pond margins. Pig Frogs are active year-­round, weather permitting, especially in Florida. They ­will move around within large aquatic systems. For example, Ligas (1960) recorded a tagged frog moving 411 m in 52 days between recaptures. Pig Frogs are generally photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond or lake (Hailman and Jaeger, 1974). Pig Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling may occur at any time during the warm season. In Georgia, this extends from late March to early September (Wright, 1932; Lamb, 1980, 1984b), but in central Florida calls are heard from early March to November (Bancroft et al., 1983). In south Florida, calling can occur all year (Duellman and Schwartz, 1958; Ligas, 1960), although breeding likely takes place only from April to July. Calls in Louisiana are heard from February to August (Dundee and Rossman, 1989). The majority of chorusing occurs at night, but calls can be heard at most any time of the day. The call is a guttural series of grunts reminiscent of a pig grunting, hence its common name. Males also produce a loud single grunt which may serve a territorial function. Unreceptive females produce a grunt that is similar to a male’s call. If a male approaches a calling male too closely, agonistic be­hav­ior consisting of the assumption of an agonistic posture, chasing, and wrestling ensues. Although unstudied, the vocal repertoire of this species and its territorial be­hav­ior are prob­ably as complex as in L. catesbeianus. Calling occurs while floating in the ­water or while perched on floating vegetation, such as lily pads or lotus (Walkowski, 2020). Males also tend to form loose choruses in thick vegetation, which provides concealment. Males tend to remain at 1 location. Chorusing occurs at temperatures >21°C.

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514  Ranidae

BREEDING SITES

DIET

Breeding occurs in a variety of wetlands, from small to large ponds and lakes to large wet prairies. Pig Frogs ­favor quiet, open ­waters with an abundance of emergent vegetation (see Adult Habitat). In the Everglades, Pig Frogs are more abundant in sloughs and sawgrass prairies than they are in large wet prairies (Baber et al., 2005).

The diet consists of many types of invertebrates, principally beetles, larval and adult dragonflies, and crayfish (Procambarus) (Carr, 1940a; Duellman and Schwartz, 1958; Ligas, 1960; Lamb, 1980, 1984b; Ugarte, 2004). Other items include grass shrimp, snails, millipedes, centipedes, spiders, crickets, true bugs, wasps, grasshoppers, cicadas, alligator fleas, larval and adult lepidopterans, grasshoppers, leeches, and flies. Pig Frogs eat vertebrates, including lizards (Plestiodon laticeps), snakes (Nerodia fasciata), frogs (Acris gryllus, Dryophytes cinereus, L. grylio, L. sphenocephalus), salamanders (Eurycea quadridigitata), and fish (mosquito fish, least killifish, sailfin molly, marsh killifish) (Duellman and Schwartz, 1958; Lamb, 1980, 1984b; Ugarte, 2004). They likely consume any animal they can cram into their large mouths, much as American Bullfrogs do, but in the Everglades, crayfish comprised 75% by volume of the prey consumed (Ligas, 1960). During the nonbreeding season, males and females have the same diet, consisting mostly of aquatic species. Fish assume more importance during droughts and the dry season; males tend to eat more crayfish in the wet season, whereas females eat more frogs, at least in the Everglades (Ugarte, 2004). However, ­there also are sex-­related differences in the relative types and abundance of prey eaten during the breeding season, reflecting the differences in the habitats of males and females at this time (Lamb, 1980). Males eat infrequently during the breeding season, and most frogs lack fat bodies during the wet summer season.

REPRODUCTION

In southwest Georgia, mature spermatozoa are pre­sent in male testes year-­round, whereas mature ova are pre­sent in females from April to July (Lamb, 1980, 1984b); thus, the breeding season is likely about 4 months in the north. Oogenesis begins in August. Even in south Florida, most breeding takes place from March to September with a peak in June (Ligas, 1960), although breeding ­there can take place year-­round (Babbitt and Tanner, 2000; Ugarte, 2004; Baber et al., 2005). Clutch size ranges from 6,000 to 34,000, with clutch size proportional to female body size (Wright, 1932; Livezey and Wright, 1947; Ligas, 1960). The mean clutch size in Ugarte’s (2004) Everglades study was 9,149 eggs. Egg dia­meter is not correlated with female SUL. Hatching occurs in 2–4 days. LARVAL ECOLOGY

Larvae begin feeding 6.5 days ­after hatching (Ligas, 1960) and feed extensively on unicellular algae. Tadpoles spend most of their time in shallow ­water and are abundant among stands of Juncus (Lamb, 1980). The larval period is 3–15 months and varies considerably among individuals and with latitude. In southwest Georgia, for example, larvae overwinter and transform ­after 10–15 months (Lamb, 1980); in the Everglades, transformation occurs as soon as 3 months (Ligas, 1960). Newly metamorphosed froglets are 30–49 mm SUL (Wright, 1932; Ligas, 1960).

PREDATION AND DEFENSE

The skin secretions of L. grylio contain antimicrobial peptides, which may assist in defense against microorganisms (Kim et al., 2000). Pig Frogs are wary and difficult to approach, but Carr (1940a) indicated this might be the result of hunting pressure. Their coloration makes them difficult to see in emergent vegetation. Allen (1932) noted that Pig Frogs give off a musty odor when handled and that their “slime” is ­bitter to the taste. Still, many vertebrates likely eat Pig Frogs, especially alligators, snakes (Nerodia fasciata, N. erythrogaster), wading birds, and mammals (raccoons) (Wright, 1932). The tadpoles appear to be fully palatable and are eaten by snakes and predaceous invertebrates. POPULATION BIOLOGY

Tadpole of Lithobates grylio. Photo: David Dennis

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Females must be 94–96 mm SUL to begin breeding in the Everglades, with males reaching maturity at 70–75 mm SUL (Ligas, 1960; Ugarte, 2004). Ovaries may have developing eggs and oocytes at 74 and 86 mm SUL, respectively. In the Everglades, adult sizes are reached in 1 yr following meta-

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COMMUNITY ECOLOGY

The Pig Frog is the common large ranid throughout much of its range. Lithobates catesbeianus often occurs within the area, but it is not nearly as abundant and often inhabits smaller ponds. Occasionally the species are found together, however, such as at Kingfisher Pond on Savannah National Wildlife Refuge in South Carolina. The interaction between the Florida Bullfrog and the Pig Frog deserves study. DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus has been found on this species in Florida, based on museum specimens (Karwacki et al., 2021). Perkinsea and Ranavirus (as early as 1922) also ­were found on specimens from the state. This species is parasitized by the trematodes Clinostomum marginatum, Gyrodactylus sp., and Haematoloechus longiplexus (Manter, 1938; Paetow et al., 2009). Ranavirus has been found in Pig Frogs introduced into Chinese aquaculture (Zhang et al., 2001). As of 2007, amphibian chytrid fungus had not been detected on Pig Frogs (Rothermel et al., 2008). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Adult Lithobates grylio. Photo: Alan Cressler

morphosis in males and ­after 1.5 yrs in females. This suggests that males are capable of breeding the first summer ­after metamorphosis, but that females first breed the second summer following transformation. In the north, age at maturity is prob­ably ­later ­because of the more extended larval period and cold winter season, although Mount (1975) recorded 2 calling males in Alabama at only 52 and 59 mm SUL. Maximum longevity is 5 yrs based on growth curves (Ugarte, 2004). In north central Florida, female Pig Frogs have higher rates of survivorship than ­either adult males or juveniles (Wood et al., 1998). Survivorship rates do not change monthly, although low capture probabilities vary among sampling periods. Survivorship varies among sampling sites, with frogs at some locations having much higher rates of monthly survival than at other locations. Abundance also varies considerably among sampling periods based on mark-­recapture studies. At 1 pond in Gainesville, Wood et al. (1998) estimated ­there ­were as many as 715 juveniles, 219 males, and 499 females, but at a smaller pond the maximum estimates ­were 39 juveniles, 16 males, and 19 females.

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Metals. Mercury is common in Pig Frogs from south Florida (Ugarte et al., 2005). Ugarte (2004) found a mean of 121.4 ng/g total mercury (maximum 2.3 mg/kg wet mass), with liver levels 2–5 times the levels in leg muscle. The amount of mercury is not correlated with SUL. High levels are recorded in both protected and unprotected sites, and Ugarte et al. (2005) recommended no harvesting in areas with high mercury content. STATUS AND CONSERVATION

This species is hunted heavi­ly for food in Florida and Georgia. Ligas (1960) noted that literally “hundreds of thousands” ­were taken by commercial hunters in the Everglades throughout

Habitat of Lithobates grylio. Jasper County, South Carolina. Photo: C.K. Dodd, Jr.

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516  Ranidae

the year. Frogs ­were spotlighted and gigged, dip-­netted, or captured by hand, much as they are ­today. Catches ­were as high as 34–45 kg/night in 1959 (Ligas, 1960). The effect of long-­term un­regu­la­ted harvest is seen in the maximum size decrease from the 1950s to the 2000s. Harvest selectively removes the largest frogs (Ugarte, 2004). In north Florida, Beck (1948) indicated that Pig Frog populations in unprotected areas had been decimated by frog hunters. Ligas (1960) recommended a series of habitat and administrative mea­sures to ensure and monitor the harvest, including the implementation of a licensing program and closure to harvest during the breeding season, at least in the Everglades. Although a commercial license is now required, ­there are still no season, size, or quantity limits on any large ranid species in Florida, and no recreational license is required. Ugarte (2004) noted that both harvest and periodic ­water draw-­downs affected Pig Frog demography. Frogs become less common ­after draw-­downs and survivorship decreases, with highest catches in areas with the longest hydroperiod. This is particularly evident among the juvenile population, where recruitment is low following draw-­down. The species is relatively tolerant of urbanization and environmental perturbation as long as wetlands are main-

tained (Delis et al., 1996; Davis et al., 2017). Since it does not venture far from the shoreline, development of uplands may not affect the species as long as pollution and urban runoff is controlled. Still, populations may not be as extensive or abundant in urban, silvicultural, or agricultural settings as they are in native ecosystems (Surdick, 2005). ­Because the diet consists largely of crayfish, and crayfish populations may be reduced in disturbed habitats, prey availability rather than surrounding habitat use per se may be responsible for reduced abundance in some areas. The species quickly recovered from saltwater overwash ­after Hurricane Dennis hit the Gulf Coast of Florida (Gunzburger et al., 2010), and Pig Frogs ­were pre­sent following hurricanes Ivan and Katrina in Louisiana, although abundance estimates ­were not available (Schriever et al., 2009). Viosca (1923) noted they could tolerate moderate salinity. Roads likely take a considerable toll on Pig Frogs, especially when they cross expansive marshes and wet prairies; kills ­were especially prevalent from April to August at Paunes Prairie south of Gainesville (Smith and Dodd, 2003). Culverts and underpasses helped reduce mortality considerably (Dodd et al., 2004). From 1990 to 1994, 613 Pig Frogs ­were collected in Florida for the pet trade (Enge, 2005a).

Lithobates heckscheri (Wright, 1924) River Frog

dark gray. River Frogs are often heavi­ly mottled ventrally on the throat and between the front legs. A pale, crescent-­ shaped line may girdle the groin. ­There are narrow dark bars on the upper sides of the legs. The ultimate phalange of the fourth hind toe is ­free of webbing. Males are smaller than females but have wider tympanums. Adults are 83–155 mm SUL. Larvae. Tadpoles are initially small and black with gold to white transverse bands on the snout and body. ­These bands dis­appear when the tadpole is about half grown. Tails are transparent. Mature tadpoles are dark greenish olive to olive with fine, pale greenish-­yellow spots or flecks over the dorsum. Venters are pigmented and the gut is not vis­i­ble. The lateral line system is easily vis­i­ble. Tails are elongate, and the top fin is not as deep as the musculature. The tails have a prominent black band that extends along the upper tail musculature about two-­thirds of its length. The entire tail is rimmed with black pigment. The black band and tail border pre­sent a striking contrast to the nearly clear tail fins. Tadpole lengths reach 160 mm TL (Mount, 1975). Older tadpoles and metamorphs have brick-­red eyes; the iris eventually turns golden in adults. Altig (1972a) described tadpole development. Albino tadpoles ­were observed in Georgia (Wright, 1924). Eggs. The eggs have not been described in detail, but presumably they are similar to L. grylio. The egg mass is

ETYMOLOGY

heckscheri: named for the Heckscher Foundation for the Advancement of Research, established at Cornell University by August Heckscher. The Heckscher Foundation supported A.H. Wright’s work in the Okefenokee Swamp. NOMENCLATURE

Conant and Collins (1998): Rana heckscheri Dubois (2006): Lithobates (Aquarana) heckscheri Fouquette and Dubois (2014): Rana (Lithobates) heckscheri Synonyms: Rana heckscheri IDENTIFICATION

Adults. This is a large, brown to grayish-­olive frog with evenly spaced tubercles occurring throughout its dorsum which gives it a textured appearance. On the sides, the color blends to a more olive or even cinnamon-­brown color. The dorsum is mottled in black. White spots may be pre­sent on the dark lower jaw. Dorsolateral folds are absent. Venters are gray with white spots or they may be a uniform light to

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prob­ably oviposited in a sheet-­like surface film as in other members of the Aquarana clade (Wright and Wright, 1949), although Allen (1938) stated that wild-­caught females deposited eggs in a “mass” in captivity. The vitellus is 1.5–2 mm in dia­meter (Wright, 1932). DISTRIBUTION

The species historically occurred from the Lumber and Cape Fear river systems in North Carolina (not seen since 1975; Beane, 1998) south along the Atlantic Coastal Plain below the Fall Line, throughout the northern and central Florida peninsula, and westward to the Wolf River in southern Mississippi. Krysko et al. (2019) gave the southernmost locations as Marion and Volusia counties, Florida, but Punzo (1991b, 1992b) supposedly used tadpoles from the Hillsborough River in the Tampa Bay area in his experiments. The identity of t­ hese individuals is questionable. The species also has been introduced into China (Kraus, 2009). Impor­tant distributional references include: Alabama (Mount, 1975), Florida (Krysko et al., 2019), Georgia (Williamson and Moulis, 1994; Jensen et al., 2008; Stevenson and Chandler, 2017), Mississippi (Allen, 1932), North Carolina (Simmons and Hardy, 1959; Beane, 1998; Dorcas et al., 2007), and South Carolina (Leiden et al., 1999; Beane et al., 2010; Dodd and Barichivich, 2017). FOSSIL REC­O RD

No fossils are known.

SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates heckscheri is a member of the American Bullfrog species group (Aquarana clade), consisting of 7 closely related species (L. catesbeianus, L. clamitans, L. grylio, L. heckscheri, L. okaloosae, L. septentrionalis, L. virgatipes) that evolved rapidly from a common ancestor during the late Miocene to early Pliocene (Austin et al., 2003a; Hillis and Wilcox, 2005). The species is closely related to L. grylio. Laboratory crosses of L. heckscheri with L. clamitans ­were unsuccessful (Moore, 1949a, 1955). ADULT HABITAT

The River Frog is a species of flowing streams, rivers, swampy backwaters, fluvial swamps, and medium-­to large-­sized ponds and lakes; the species also occurs in small streams flowing between larger ­water bodies, oxbows, sinkhole ponds, and permanently flooded borrow pits. Shaded banks with good cover are preferred. Surrounding forest often consists of mesic hammock and riparian cypress and hardwoods (Harima, 1969). AQUATIC AND TERRESTRIAL ECOLOGY

River Frogs are active at temperatures of 18–35°C, with an optimum of 25°C (Hansen, 1957). They frequent banks and shorelines with high moisture content, and often are found in sphagnum. They usually sit within 23 cm of ­water and face the open ­water. Frogs ­will change position along the shore, but usually stay within a par­tic­u­lar area. For example, Hansen (1957) recorded movements of about 10 m between recaptures, with no differences between males and females; juveniles tended to move farther than adults. Activity areas averaged 17 m × 1.5 m. Allen (1938) found a single juvenile dispersing 183 m from ­water a month ­after transformation; few juveniles remained around the larval lake. Adults also may be observed sitting on logs or debris in the ­water. River Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities (Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a river or pond (Hailman and Jaeger, 1974). River Frogs likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling occurs from April to August. The call is a loud, distinctive snore or snort. Males call from both the shoreline and from shallow ­water. BREEDING SITES Distribution of Lithobates heckscheri

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Breeding occurs along the shorelines of rivers, swampy backwaters, and ponds (see Adult Habitat).

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518  Ranidae

REPRODUCTION

Breeding occurs from spring through late summer. Reports of clutch size include about 5,000 eggs per mass (Allen, 1938) and 14,000 eggs estimated from a single female (Wright, 1932). Hatching occurs in 10–15 days. According to Allen (1938), about 10% of the eggs fail to hatch. LARVAL ECOLOGY

Lithobates heckscheri tadpoles form large schools consisting of hundreds of individuals. For example, I captured 1,132 tadpoles in 8 trap nights at a pond on the Savannah National Wildlife Refuge; 3 traps accounted for >300 tadpoles each on a single night. ­These schools move parallel to the shoreline and remain in shallow ­water during the day. At night, tadpoles move into deeper ­water, although the tadpoles are photonegative in laboratory experiments (Altig and Christensen, 1981). Allen (1938) noted that tadpole schools tended to remain within a defined area of about 46 m in linear distance. Tadpoles respond positively to the presence of food and food odors by increasing feeding activity, even when food is abundant. Tadpoles in schools

Recent metamorph of Lithobates heckscheri. Photo: C.K. Dodd, Jr.

exhibit a degree of social facilitation whereby swimming speeds are increased and avoidance of unpleasant stimuli is enhanced in the presence of groups of conspecifics (Punzo, 1991b, 1992b). Tadpoles can be heard making a smacking sound as dozens of tadpoles si­mul­ta­neously gulp air at the surface as a school moves through shallow ­water. The larval period extends throughout the winter (Wright and Wright, 1949). For example, in Florida, eggs hatching on 8 June resulted in metamorphs by 10 April the following year (Allen, 1938). Allen (1938) estimated that as many as 20% died during transformation. Mass metamorphosis has been observed, with accounts of thousands of individuals transforming at once (Wright and Wright, 1933). Jensen and Wright (2014) counted 856 in a single photo­graph and estimated that the full aggregation of transforming frogs was >4,000 individuals. The transformed aggregation may remain together for a period of several days before dispersing. DIET

Lithobates heckscheri eats a variety of invertebrates, particularly crayfish and insects. Specific items include millipedes, centipedes, spiders, beetles, true bugs, tabanid flies, cockroaches, and caddisflies (Lamb, 1980). It seems likely that a River Frog would eat any prey item it could fit into its mouth. PREDATION AND DEFENSE

Schooling tadpoles of Lithobates heckscheri. Photo: Dirk Stevenson

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Adult skin secretions of L. heckscheri contain antimicrobial peptides which may assist in defense against microorganisms (Conlon et al., 2007a). However, the larval skin does not appear to contain toxic or noxious properties (Altig and Christensen, 1981). This species is easily approached by ­human observers and may go limp when handled.

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Lithobates heckscheri 519

Predators of tadpoles and recent metamorphs include ­ ater snakes (Nerodia fasciata), turtles (Sternotherus minor), w and Boat-­tailed Grackles (Wright, 1932; Allen, 1938). Tadpoles respond to predator odors and the odors of injured tadpoles by swimming wildly and erratically away from the source of the odor (Altig and Christensen, 1981). They also attempt to dive and hide. POPULATION BIOLOGY

Juvenile growth rates apparently are rapid. For example, Hansen (1957) recorded growth in juveniles of 11 mm in 30 days, 6 mm in 21 days, and 4 mm in 27 days. Juveniles continue to grow through October, and growth slows appreciably in winter. The age at maturity is unknown but presumably similar to other large Southeastern ranids. DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus has been found on this species in Florida from individuals collected as early as 1928, based on

Habitat of Lithobates heckscheri. Jasper County, South Carolina. Acris gryllus, Anaxyrus fowleri, Anaxyrus terrestris, Lithobates catesbeianus, Lithobates clamitans, Lithobates grylio, Lithobates sphenocephalus, and Dryophytes cinereus also bred in this pond. Photo: C.K. Dodd, Jr.

museum specimens (Karwacki et al., 2021). Perkinsea (as early as 1928) and Ranavirus also ­were found on specimens from the state. Larval River Frogs are parasitized by the crustaceans (“fish lice”) Argulus americanus (Goin and Ogren, 1956; Watermolen, 2019) and A. diversus (Clark, 2001). The trematode Gorgodera amplicava has been reported from L. heckscheri (Parker, 1941), as has the nematode Gyrinicola batrachiensis (Pryor and Greiner, 2004). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Recent metamorph of Lithobates heckscheri. Photo: Dirk Stevenson

Metals. Mercuric chloride inhibits fertilization at >1mg/L, with total prevention at 5 mg/L (Punzo, 1993a). The LC50 (3 hr) is 1.43 mg/L for eggs and the LC50 (96 hr) for tadpoles is 0.68 mg/L. Mercury also impairs the development of oocytes in adult L. heckscheri (Punzo, 1993b). Developmental abnormalities (kinked and curved tails) are common in L. heckscheri exposed to mercuric chloride concentrations >1 mg/L (Punzo, 1993a). STATUS AND CONSERVATION

Adult Lithobates heckscheri. Photo: C.K. Dodd, Jr.

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­ here are no published studies on the status of this species T throughout much of its range. As a large ranid, it presumably has been hunted for food in the rural South. Undoubtedly, populations have been lost as habitats ­were destroyed and river floodplains altered. According to Jensen et al. (2008), the species is relatively common in Georgia. Aresco (2004) noted only 6 breeding localities in Alabama and that ­there have been no rec­ords for 25 yrs. Beane (1998) could find no extant populations in North Carolina despite extensive surveys. From 1990 to 1994, 32 River Frogs ­were collected in Florida for the pet trade (Enge, 2005a). This species is considered a Protected Nongame Species in Alabama

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520  Ranidae

Lithobates kauffeldi (Feinberg, Newman, Watkins-­Colwell, Schlesinger, Zarate, Curry, Shaffer, and Burger, 2014) Atlantic Coast Leopard Frog ETYMOLOGY

kauffeldi: named for herpetologist Carl F. Kauffeld (1911– 1974), who first recognized that leopard frogs in the vicinity of New York City might represent an undescribed species. NOMENCLATURE

Feinberg et al. (2014): Rana kauffeldi Synonyms: Lithobates pipiens [in part], Lithobates sphenocephalus [in part], Rana brachycephala, R. halecina, R. pipiens [in part], R. sphenocephala [in part], R. utricularius [in part], R. virescens Although Kauffeld recognized the distinctiveness of leopard frogs in the vicinity of New York City as early as the 1930s (Kauffeld, 1937), this species was not formally described ­until 2014 (Feinberg et al., 2014). As such, most prior lit­er­a­ture referring to leopard frogs on the northern Atlantic Coastal Plain refer to ­either L. pipiens or L. sphenocephalus. In addition, this species is sympatric with both species at numerous locations—­with L. pipiens in northern New Jersey and New York and with L. sphenocephalus from central and southern New Jersey southward to eastern North Carolina. Thus, it may be impossible to associate published lit­er­a­ture and rec­ords of “leopard frogs” within ­these regions with a par­tic­u­lar species. The nomenclatural history of this taxon is reviewed by Feinberg et al. (2014). IDENTIFICATION

Adults. The base color is dark brown to green with dorsal oval and irregular black spots. Overton (1914) noted that some individuals ­were nearly black dorsally. The spots may be bordered by a lighter whitish ring, and are generally fewer in number than ­those found on other leopard frog species in the region. Most of the spots are smaller than the eye when compared with L. pipiens and, unlike this latter species, the spots are usually absent from the snout. The dorsum and lateral sides are rugose with slight ridges paralleling a distinct dorsolateral fold. This fold runs from the back of the eye to the frog’s posterior. ­There is a dark stripe ­running from the pointed snout through the eye and extending dorsally around the tympanum. A white horizontal stripe runs across the top of the iris. The tympanum has a faint white spot in its center. A white line is pre­sent along the upper lip. Black parallel bars are evenly spaced on the upper surfaces of the tibiofibula of

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the hind limbs, and the rear of the femur is dark and heavi­ly reticulated (see Schlesinger et al., 2017, Fig. 6 for a comparison of the femoral patterns of the 3 eastern leopard frogs). Black bars are pre­sent on the forelimbs, but they are not as regularly spaced. Venters are white. Males are easily distinguished from females by their very large vocal sacs that are pre­sent year-­round. An albino adult was reported from Long Island, New York (Mulertt, 1896). Ditmars (1905) gives a length of ca. 65–80 mm SUL, whereas Mathewson (1955) gives a maximum length of 110 mm SUL. The Atlantic Coast Leopard Frog looks very similar to the Southern and Northern Leopard Frogs, and ­there is no single set of characters that differentiate L. kauffeldi from L. sphenocephalus. When compared with L. sphenocephalus, L. kauffeldi is usually duller with a blunter snout. The femoral reticulum (dorsal pattern on the thighs) in the Mid-­Atlantic offers a means to separate L. kauffeldi (dark reticulum) from L. sphenocephalus (light reticulum) in the field. Farther south beyond the known range of L. kauffeldi (e.g., south of North Carolina), however, L. sphenocephalus also has a predominantly dark reticulum (J. Feinberg, personal communication). The vocal sacs of L. kauffeldi also are larger than ­those of L. sphenocephalus. Larvae. No formal descriptions of the tadpoles are available, but they appear very similar to L. sphenocephalus. Eggs. No descriptions of the eggs are available, although presumably they are similar to other leopard frog eggs. Moore (1949b) reported egg dia­meters of 1.63–1.97 mm. DISTRIBUTION

Atlantic Coast Leopard Frogs occur from central Connecticut southward along the coast to Washington County, North Carolina. The current distribution is divided into 4 geographic regions, although the northern 3 likely ­were contiguous at one time. The northern 3 populations include central Connecticut (two populations in Middlesex County; Klemens et al., 2021), southeastern New York, and from southwestern New York through the Delmarva Peninsula. The southernmost part of the range includes southeastern ­Virginia through northeastern North Carolina. The species ranges from 208 m in northern New Jersey to sea level along the coast (Schlesinger et al., 2017). Kiviat (2011) provided survey data from the Hackensack Meadowlands, New Jersey, in 2006. Schlesinger et al. (2017) reviewed the distribution of this species. Atlantic Coast Leopard Frogs ­were found historically on islands, including Staten Island and Long Island, New York (Overton, 1914, possibly extirpated on Long Island; Schlesinger et al., 2017; Mathewson, 1955; Nicholls et al., 2017, tiny population on Staten Island). Reports of Southern Leopard Frogs from Kent Island, Mary­land (Grogan and Bystrak, 1973b), Assateague Island in Mary­land and ­Virginia (Lee, 1972;

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Lithobates kauffeldi 521

ADULT HABITAT

Lithobates kauffeldi inhabits lowland areas along a relatively narrow band bordering the Atlantic Coast. The farthest inland population is approximately 40 km from the coastline. ­These frogs live in freshwater wetlands, interior riparian floodplains, and adjacent to tidally influenced backwaters (Feinberg et al., 2014). Overton (1914) gives the habitat as marshes, especially salt marshes. Lithobates kauffeldi prefers large, open wetlands, marshes, cypress-­gum swamps, and slow-­flowing streams bordered by open upland and early successional habitats. ­Water should be clear and shallow with emergent shrubs and vegetation such as cattail (Typha sp.) and the invasive common reed Phragmites australis. Temporary wetlands are sometimes used. AQUATIC AND TERRESTRIAL ECOLOGY

Distribution of Lithobates kauffeldi. Dark gray indicates extant populations; light gray indicates extirpated populations.

Mitchell and Anderson, 1994), Chincoteague and Wallops islands, ­Virginia (Conant et al., 1990; Delis and Meshaka, 2019), and Shackleford Banks, Smith, Bodie, Hatteras, and Roanoke islands in North Carolina (Brimley, 1944; Lewis, 1946; Engels, 1952; Braswell, 1988; Gaul and Mitchell, 2007; Parlin et al., 2019) may actually refer to L. kauffeldi. FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Kauffeld (1937) noted phenotypic differences among the leopard frogs of the northern Atlantic Coast and suspected that dif­fer­ent taxa ­were pre­sent. Newman et al. (2012) used nuclear and mitochondrial DNA analyses to identify a unique ge­ne­tic lineage and confirm a cryptic leopard frog subsequently named as L. kauffeldi (Feinberg et al., 2014). Three evolutionary lineages ­were involved in the tri-­state analyses, with no detectable gene flow with adjacent leopard frog species. Feinberg et al. (2014) expanded ge­ne­tic sampling and added bioacoustic analyses, thus extending the known range southward and clarifying relationships with other leopard frog populations northward and inland. Although similar in appearance to L. pipiens and L. sphenocephalus, this species maintains a number of unique morphological characteristics. Previous suggestions of morphological variation among ­these geo­graph­i­cally proximate or sympatric species may actually refer to characteristics of L. kauffeldi.

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­ ittle is known concerning the life history of L. kauffeldi L during the nonbreeding season or about its dispersal away from breeding sites. The species is reported to be terrestrial on Staten Island, New York (Overton, 1914; Mathewson, 1955). Occasional individuals have been observed terrestrially late in the season (Feinberg et al., 2014). CALLING ACTIVITY AND MATE SE­L ECTION

Breeding occurs from late winter through spring. In New York, it may begin in February depending on temperature; most breeding ­there occurs from late March to April and into May. On Long Island, calling extends from early March to June (Overton, 1914; Gibbs et al., 2007). Calling in other areas begins mostly in mid-­March, but calls in North Carolina have been heard as early as 3 February (Schlesinger et al., 2017). Chorusing begins ­after several days of above average temperatures and continues when nighttime air temperatures are 10–18°C. Calling is largely nocturnal for most of the season, but diurnal calling occurs early in the season and for the first 2–3 weeks of the breeding season. As the season progresses through early June in New York, nocturnal chorusing becomes less frequent and more episodic. Calls have been heard as late as 12 June in New Jersey (Kiviat, 2011). ­There may be a second late-­season calling period from late August to November with the advent of cooler weather (Feinberg et al., 2014). Males congregate in small groups of 5 of more frogs to call. Individuals may be as close as 30 cm from one another in shallow ­water among the emergent vegetation. Overton (1914) stated that males ­will call together in a chorus lasting 2–3 minutes followed by a period of silence. Calls are low pitched and do not carry far (Mathewson, 1955). Feinberg et al. (2014) described the call as a “single-­noted unpulsed ‘chuck’ that is distinct from the pulsed ‘ak-­ak-ak’ of R. ­sphenocephala and the snore-­like calls of R. pipiens and

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522  Ranidae

R. palustris.” Calls are produced from large, lateral, paired external vocal sacs. Call characteristics include a call length of 33–71 ms, a call rate of 0.7–2.35 call/sec, and a dominant frequency of 1,211–1,593 Hz at 10°C (Feinberg et al., 2014). ­These authors provided a sonogram of the call in comparison with other Lithobates species. BREEDING SITES

Breeding occurs in shallow ­water along the shorelines of large wetlands.

Tadpole of Lithobates kauffeldi. Photo: Jeremy Feinberg

REPRODUCTION

­ ittle information is available. Egg masses are globular, L oviposited ­under ­water, and clustered near one another (Moore, 1949b). From 2,000 to 3,000 eggs are oviposited and take 2–3 weeks to hatch (Mathewson, 1955). In laboratory crosses between L. kauffeldi and L. pipiens, development occurs through metamorphosis (Porter, 1941). Eggs are tolerant of higher temperatures than eggs of L. pipiens during development, with complete development at 28–29°C and some development at higher temperatures (Porter, 1941; Moore, 1949b).

PREDATION AND DEFENSE

Nothing is known, but it is prob­ably similar to other leopard frogs; that is, is has an alertness to predators followed by a warning scream and rapid jump to deeper ­water. Predators undoubtedly include raccoons and other mammals, predatory birds, snapping turtles, and ­water snakes. The larvae are likely consumed by predaceous diving beetles, dragonfly nymphs, and a variety of small vertebrates. POPULATION BIOLOGY

LARVAL ECOLOGY

­ ittle information is available. Mathewson (1955) stated L that the larval period is ca. 2 months. Gaul and Mitchell (2007) noted the presence of larvae (as L. sphenocephalus) at all times of the year in eastern North Carolina, DIET

Specific items have not been reported but are prob­ably similar to the mostly insectivorous diet of other leopard frogs.

Nothing is known of this species’ population dynamics. The species seems to be uncommon in North Carolina (Schlesinger et al., 2017), but more data are needed. DISEASES, PARASITES, AND MALFORMATIONS

It is pos­si­ble that some of the endoparasites reported from leopard frogs (L. sphenocephalus) in North Carolina by Brandt (1936) might have come from this species. Both Bd and ranavirus have been reported from this species in North Carolina (Lentz et al., 2021). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Nothing is known, but prob­ably similar to stressors affecting other leopard frog species. STATUS AND CONSERVATION

Egg mass of Lithobates kauffeldi. Photo: Jeremy Feinberg

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Undoubtedly many f­ actors may affect the current status of L. kauffeldi, particularly habitat loss and emerging infectious disease such as amphibian chytrid fungus. Populations in the northern portion of the species’ range have been extirpated, particularly in Connecticut (in New Haven and Fairfield counties; Klemens et al., 2021) and adjacent areas of New York and Pennsylvania. Overton (1914) stated that it was very common in the salt marshes of ­Great South Beach, Long Island, but it may now be extirpated from the entire island. Habitat alteration also may adversely affect this species by allowing vegetation encroachment thus closing off the preferred open canopy. Climate change

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Lithobates okaloosae 523

Adult Lithobates kauffeldi. Red phase. Male. Note the thigh reticulations and the very large vocal sac. Photo: Erik Kiviat/Hudsonia Type locality of Lithobates kauffeldi, Richmond County, New York as it appeared in 2007. Much of this site was destroyed by the construction of Amazon and Ikea ware­houses. A small portion of the wetland remains to the right of the photo­graph. Photo: Jeremy Feinberg

leading to sea level rise could also affect coastal populations in adjacent freshwater marshes. Coastal impoundment populations in Delaware dis­appeared following storm impacts, but long-­term effects of storms are unknown, as are the effects of (temporary) saltwater intrusion. More information on threats to existing populations is necessary in order to assess the status of this cryptic species. The species is proposed as Endangered by the state of Connecticut (Klemens et al., 2021). Adult Lithobates kauffeldi. Green phase. Female. Photo: Jeremy Feinberg

Lithobates okaloosae (Moler, 1985) Florida Bog Frog ETYMOLOGY

okaloosae: named for Okaloosa County, Florida, where the species was first discovered. NOMENCLATURE

Conant and Collins (1998): Rana okaloosae Dubois (2006): Lithobates (Aquarana) okaloosae Fouquette and Dubois (2014): Rana (Lithobates) okaloosae Synonyms: Rana okaloosae IDENTIFICATION

Adults. The Florida Bog Frog is a small, yellowish-­greenish to yellowish-­brown frog that is superficially similar to L. clamitans.

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Light-­colored dorsolateral folds are pre­sent, but they do not extend to the groin. The dorsum is uniformly colored and covered with small, white tubercles, but the white venter is smooth with dark vermiculation. The tympanum is flat in both sexes. Throats may be infused with yellow in both sexes, and ­there are light spots on the lower jaw. The toe webbing is distinctive, with 3 phalanges of the fourth toe and 2 phalanges of all other toes ­free of webbing. In sympatric ranids, toe webbing is more extensive. Males have internal vocal sacs, a slightly larger tympanum, and swollen thumbs. Males are only slightly smaller than females. In the type series, males ­were 34.8–45.8 mm SUL (mean 40.6 mm) and females ­were 38.2–48.8 mm SUL (mean 44.6 mm) (Moler, 1985). Bishop (2005) recorded a mean male size of 40.2 mm SUL (range 34.4–56.9 mm) and a female mean size of 41.5 mm SUL (range 33.5–48.8 mm). Gorman (2009) recorded L. okaloosae with a mean of 39.7 mm SUL (range 26.6–49.8 mm). Body mass increases with SUL.

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524  Ranidae

Larvae. The tadpole is olive brown with numerous dusky spots on the tail; tadpoles of L. clamitans lack ­these spots. The black venter is marked by well-­defined white spots, and the coiled intestines are not vis­i­ble. The white ventral spotting is distinctive and separates them from sympatric L. clamitans larvae, whose spots are more gold on a deep brown background. Tadpoles reach a maximum length of 56 mm TL (Moler, 1985). Moler (1985) illustrated the tadpole and its mouthparts, and Priestley et al. (2010) illustrated differences between sympatric L. okaloosae and L. clamitans larvae. Eggs. The eggs of this species have not been described. Presumably they are similar to ­those of L. clamitans. Eggs are oviposited in a single-­layer surface film. DISTRIBUTION

This species is known only from Walton, Okaloosa, and Santa Rosa counties in the western Florida Panhandle. Sites (>60 known) are within the East Bay, Yellow, and Shoal river drainages. Most of its distribution occurs on Eglin Air Force Base. Impor­tant distributional references include Bishop (2005) and Krysko et al. (2019). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates okaloosae is a member of the American Bullfrog species group (Aquarana clade), which consists of 7 closely related species (L. catesbeianus, L. clamitans, L. grylio, L. heckscheri, L. okaloosae, L. septentrionalis, L. virgatipes) that evolved rapidly from a common ancestor during the late

Miocene to early Pliocene (Austin et al., 2003a; Hillis and Wilcox, 2005). Lithobates okaloosae is actually phyloge­ne­ tically embedded within L. clamitans, which suggests a recent origin for the species (Austin et al., 2003). Polymorphism is low in L. okaloosae. Non-­panmictic populations are maintained by continuous ge­ne­tic structuring along streambeds, with weak isolation-­by-­distance (Austin et al., 2011). Lithobates okaloosae may be the result of hybrid origin, incomplete lineage sorting, or recent hybridization. Hybrids with L. clamitans have been reported (Moler, 1992). ADULT HABITAT

The Florida Bog Frog is found in isolated acid seeps (pH 4.1–5.5), along shallow boggy overflows of larger seepage streams, in headwater streams, stream sections below small impoundments, and rarely along the edges of ponds. ­Water should be clear, shallow, and not stagnant. For example, Gorman et al. (2009) studied the species’ biology along a shallow (1–20 cm) headwater stream that varied from 7 to 22 m in width. The species is frequently associated with lush beds of sphagnum moss (Sphagnum). Habitats consist of small streams that usually are bordered by black titi (Cliftonia monophylla) and sweetbay (Magnolia virginiana), and may be associated with Atlantic white cedar (Chamaecyparis thyoides). The habitat of clear, low-­flowing streams in excessively drained sandy soils is ­limited within the area, thus restricting the distribution of the species. Florida Bog Frogs are found most often in drainages with nearby mixed-­forest wetlands. Surrounding habitats include longleaf pine sandhills and flatwoods (Chandler et al., 2015a). Prescribed burning helps limit encroaching vegetation. In areas where hardwoods have encroached on streams, the species may be confined to areas such as power-­line rights-­of-­way. AQUATIC AND TERRESTRIAL ECOLOGY

This species apparently rarely ventures far from streams and bogs, and adults and juveniles occupy the same habitats. Activity occurs year-­round. Daily activity occurs within a relatively small area, and Gorman et al. (2009) recorded a mean daily movement of only 1.8 m; the maximum daily movement was 8.9 m. Bishop (2005) estimated the home range to be 37–188 m2 for L. okaloosae depending upon estimator used. Florida Bog Frogs likely bury into sphagnum moss or other available aquatic streamside vegetation to avoid cold temperatures. CALLING ACTIVITY AND MATE SE­L ECTION

Distribution of Lithobates okaloosae

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Calls occur throughout the warm months of March–­August. The advertisement call is distinctive from that of all other frogs within its range. The call is described as a “series of 3–21 guttural chucks [pulses] repeated at about 5 notes per

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Lithobates okaloosae 525

second” (Moler, 1985) that slows audibly ­toward the end. Bishop (2005) identified 3 basic call types (advertisement, single chuck, response) plus a release call that is given when handled. Advertisement calls average 1.4 sec in length, 7.5 pulses, a pulse rate of 0.2, and have a dominant frequency of 651–2,466 Hz (mean 1,505 Hz) (Bishop, 2005), but ­there is a ­great degree of variation among individual males. Intervals between calls range from 23 to 600 sec (mean 136 sec), and 16–21 notes constitute a call. Advertisement calls do not carry well. Males also have a single, quieter call (response) that is issued in response to calls from nearby males. The rates of this call increase in response to playback experiments of conspecifics, but not as strongly to calls of L. ­clamitans (Bishop, 2005). Females also have a soft “chuck” call, the function of which is unknown. Call dominant frequencies are much higher in L. okaloosae than in sympatric L. clamitans. Males call from along streamsides and bogs, and their distribution tends to be clumped. The mean number of males calling in Gorman et al.’s (2009) study was 7.3 (range 5–10) in a 60 m stretch of headwater stream. Males spaced themselves at a mean distance of 9.2 m, which was actually farther than their nearest neighbor distance with L. clamitans at 6.5 m. Satellite males may locate themselves near calling males; ­these satellites tend to be slightly smaller than calling males. Males usually call from 1 location for several nights before moving to a dif­fer­ent location. ­There is no correlation between male body condition and the number of nights a male attends a chorus (Neto et al., 2014). BREEDING SITES

Breeding occurs in shallow, sandy-­bottomed streams with low but steady surface flow. Calling sites are not randomly positioned but are close to the bank (mean 21 cm) and to cover (mean 7 cm), adjacent to flowing ­water, near woody debris, and have higher ­water temperatures (mean 26°C) when compared with random available sites. Most calling sites have nearby emergent and submerged vegetation (Bishop, 2005; Gorman, 2009). Canopy cover does not influence occupancy, although L. okaloosae is generally found in more open habitats. REPRODUCTION

Eggs are deposited in a single-­layer surface mass that is ­free floating. Occasional masses may be folded or become attached to vegetation. Egg masses average 12.5 cm in length (range 7–21) by 9 cm in width (range 6–13). Egg masses are deposited in shallow ­water (2–7 cm) and are 0–1,300 cm from the bank (Bishop, 2005). Clutch size is 152–345 eggs/ mass (mean 235). Bishop (2005) suggested that females might produce more than 1 clutch per year.

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Eggs of Lithobates okaloosae. Photo: Ronn Altig

Males are capable of fertilizing more than 1 clutch. For example, Bishop (2005) found that 15 males fertilized 35 egg masses, with a mean rate of 1.8 masses per male (range 0–9). The number of egg masses fertilized was directly correlated with the number of nights spent calling. The smallest successful male was 37 mm SUL, although males as small as 34.4 mm gave advertisement calls. The smallest gravid female was 37.6 mm SUL. Oviposition sites are located near calling sites, with a mean distance of only 14 cm. LARVAL ECOLOGY

Larvae have been collected throughout the year, and at least some overwinter prior to transformation. Laboratory observations suggest that tadpoles do not swim very much. Recent metamorphs are 18–20 mm SUL (Bishop, 2005). DIET

Nothing has been reported on the diet of this species. Presumably it eats any available invertebrate it can catch, and prey consumption is likely in proportion to availability. PREDATION AND DEFENSE

The coloration of this frog makes it difficult to see, thus affording crypsis from visually oriented predators. Upon approach of an intruder, the frog quickly moves to ­water. The skin secretions of L. okaloosae contain antimicrobial peptides, which may assist in defense against microorganisms (Conlon et al., 2007a).

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526  Ranidae

Adult Lithobates okaloosae. Photo: C.K. Dodd, Jr.

Comparison of the venters of tadpoles of Lithobates okaloosae (left) and Lithobates clamitans (right). Photo: David Bishop

Bishop (2003) reported a ­water snake (Nerodia fasciata) eating 2 tadpoles and attempting to catch a juvenile. Florida Bog Frogs are likely eaten by a variety of snakes, turtles, birds, and mammals. Larvae are certainly prey of predaceous invertebrates and vertebrates. POPULATION BIOLOGY

Over a 3 yr period, Bishop (2005) recorded sex ratios of 1.7:1, 1.1:1, and 2.3:1. Only 1 frog was captured in all 3 yrs. He estimated survivorship as 90–96.5% over a 2 month period during a single yr. Male L. okaloosae have high site fidelity and survival rates during the breeding season, but mark-­ recapture data suggest high annual mortality or that dispersal rates are high. Neto et al. (2014) found few frogs between years (7–14%), but more within a year (47–78%). Densities vary among populations. Neto et al. (2014) estimated densities of 10.1–19.8 frogs/ 1000 m2 at 1 site, 0.8–5.4 at a second, 5.5–9.2 at a third, and 0.8–1.1 at a fourth site. COMMUNITY ECOLOGY

This species is sympatric with L. clamitans, and the 2 have similar calling sites and breeding biology. ­There is no evidence of interspecific competition, and indeed interactions between the 2 tend to be positive (Gorman et al., 2009). Call structure and body size are very dif­fer­ent between the

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Habitat of Lithobates okaloosae. Photo: David Bishop

2 species. Lithobates okaloosae prefers sites with a greater percentage of submerged vegetation, slow-­flowing ­water, shorter distances to cover, greater amounts of emergent vegetation and woody debris, and shallower ­water than does L. clamitans (Gorman, 2009; Gorman and Haas, 2011), although ­there is considerable microhabitat overlap. Gorman

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Lithobates onca 527

(2009) could not demonstrate strong evidence for competition between the larvae of ­these closely related species. DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus has been found on this species in Florida, based on museum specimens (Karwacki et al., 2021). Perkinsea and Ranavirus also ­were found on specimens from the state, primarily from the 1980s. SUSCEPTIBILITY TO POTENTIAL STRESSORS

STATUS AND CONSERVATION

Despite its ­limited range, most known sites are on Eglin Air Force Base and are thus protected to some extent. A management plan to conserve the species has been implemented. Potential threats include fire suppression, feral pigs, stream siltation, stream impoundment, invasive plants, accidental pollution, and disease. Moler (1992) and Bishop (2005) provided a series of management recommendations to address ­these threats. This species is considered Threatened in Florida.

No information is available.

Lithobates onca (Cope in Yarrow, 1875) Relict Leopard Frog ETYMOLOGY

onca: a Latin noun meaning ‘spotted,’ as in a spotted frog. NOMENCLATURE

Stebbins (2003): Rana onca Dubois (2006): Lithobates (Lithobates) onca Fouquette and Dubois (2014): Rana (Lithobates) onca Synonyms: Rana draytoni onca, Rana pipiens onca IDENTIFICATION

Adults. This is a relatively small, brown to olive-­green leopard frog with generally smooth skin, although ­there may be warts or tubercles on the head and neck. Small black spots are bordered by gray. ­These spots are fewer, and the gray border is less distinct than in L. pipiens or L. fisheri. Brown dorsolateral folds are pre­sent, but ­there are no longitudinal folds between the dorsolateral folds. The dorsolateral folds only extend about halfway down the back, making this and L. fisheri distinct from other Southwestern leopard frogs. The head is broad and long, and spots do not occur on the snout. The stripe on the upper lip is faint or absent in front of the eye. The tympanic membrane is large and smooth. Bellies are white, although ­there may be a yellowish coloration ­toward the legs. The limbs are short, and the hind toes are fully webbed. Males are smaller than females. At Blue Point Spring, the median size was 53.5 mm SUL for males and 61.5 mm SUL for females (Bradford et al., 2004). Tanner (1931) gave mea­sure­ ments of 2 individuals as 53 and 55 mm SUL. Brennan and Holycross (2006) stated a maximum size of 89 mm SUL. Larvae. Tadpoles are large, light to olive green or grayish, with black mottling dorsally. Heads are wider than bodies.

Dodd_Canada_int_5pgs_B3.indd 527

­ here is a light line in front of the eye that extends about T halfway to the snout, and the lateral line may be vis­i­ble as tiny white dots on the body. Irises are coppery. In some larvae, when viewed laterally, the tail musculature is dark with numerous white spots extending about halfway down the tail. The rest of the tail musculature is lighter with melanophores giving a stippled appearance. The tail fins are broad and light with numerous dark flecks. In other larvae, the tail musculature is more uniform and lighter green in coloration with light tail fins; both the musculature and the tail fins have much lighter melanophore flecking. Undersides are light gray to whitish covering the intestines, but dark anteriorly around the downward-­oriented mouth with a coppery area between the oral region and the intestines. Eggs. The eggs have not been described, but are black dorsally and light gray ventrally. Eggs are deposited in a jelly mass, and clutch size is ca. 250 eggs (Brennan and Holycross, 2006). Masses may be 40–60 mm in dia­meter. DISTRIBUTION

Lithobates onca was known historically from at least 24 localities in extreme northwestern Arizona and adjacent Utah and Nevada along the Colorado, Virgin, and Muddy rivers. The historic range likely included about 190 km of river miles between Hurricane, Utah, and Black canyons below Lake Mead. It now occurs at only 5 localities in or near Lake Mead National Recreation Area. The species may have occurred in the Pahranagat Valley (Stejneger, 1893), but the identity of ­these frogs is uncertain. Eaton (1935) reported a single individual of L. onca from Rainbow Bridge Canyon, well to the east of known populations; the individual was likely L. pipiens. Lithobates onca once was much more widespread within its historic range; populations along the Virgin and Muddy rivers have been extirpated, as have ­those in Corral and Reber springs along the Colorado River within the last 15 years. Populations likely ­were extirpated with the building of Boulder Dam and the subsequent reservoir filling and the

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528  Ranidae

Microsatellite markers ­were characterized by Savage and Jaeger (2009). Oláh-­Hemmings et al. (2009) affirmed the close relationship but distinctiveness between L. onca and L. yavapaiensis. Lithobates onca and L. yavapaiensis diverged during the early Pleistocene. A population of leopard frogs found in Surprise Canyon in the western ­Grand Canyon is more closely related to L. yavapaiensis than to L. onca, despite the close proximity of populations of L. onca. Lithobates onca and L. yavapaiensis have closely related but distinct antimicrobial skin peptides (Conlon et al., 2010). ADULT HABITAT

Distribution of Lithobates onca. Dark gray indicates extant populations; light gray indicates extirpated populations. This species has been introduced into the Vegas Valley.

introduction of nonindigenous predators. Impor­tant distributional references include: Tanner (1931), Cowles and Bogert (1936), Linsdale (1940), Bradford et al. (2004), Brennan and Holycross (2006), RLFCT (2016), and Murphy (2019). FOSSIL REC­O RD

No fossils are known. Holman (2003) noted Miocene (Hemphillian) fossils from Navajo County in northern Arizona that belonged to the L. pipiens complex. The relationship between ­these fossils and L. onca is unknown.

The species historically inhabited ponds, springs, and streams within Mohave Desert Scrub habitats. The 5 known localities are all undisturbed spring systems that lack the presence of American Bullfrogs (L. catesbeianus), crayfish, and predaceous game fish. Spring pools should be open and ­free of emergent vegetation such as Scirpus. The species prefers open shorelines without dense vegetation. Bradford et al. (2005b) described 3 areas where the frog occurs or once occurred, and noted differences among the habitats (marshes vs. geothermal spring runs). AQUATIC AND TERRESTRIAL ECOLOGY

Relict Leopard Frogs likely stay close to w ­ ater and do not travel ­great distances between their isolated spring habitats. They are usually observed within a few meters of the shoreline in low riparian vegetation or directly at the ­water’s edge. In a mark-­recapture study, Bradford et al. (2004) noted that most frogs ­were captured within 17.8 m of the previous capture, and that the maximum distance between captures was 120 m. Harris (2006) also recorded a maximum distance of 121 m for frogs tracked >10 times over an 8 month period. Unpublished data suggest movements >300 m (RLFCT, 2016).

SYSTEMATICS AND GEOGRAPHIC VARIATION

Lithobates onca is a member of the Novirana clade of North American ranid frogs. It is an associate of the mostly lowland and tropical leopard frog group (or Scurrilirana) (Hillis and Wilcox, 2005). The species’ closet relatives in the United States include L. berlandieri, L. blairi, L. sphenocephalus, and especially L. yavapaiensis. Although described in 1875, this species has variously been considered a species, a phenotypic variant, or a subspecies of L. pipiens. Long thought extinct, the species was rediscovered in 1991, with subsequent molecular and morphologic analy­sis confirming its phyloge­ ne­tic uniqueness (Jaeger et al., 2001). The species is prob­ably a Pleistocene-­Holocene isolate from ancestral populations to the south. Lithobates onca currently exhibits a rather low level of ge­ne­tic heterozygosity.

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Eggs of Lithobates onca. Photo: Dana Drake

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Lithobates onca 529

Movements occur along stream courses, and the frogs prefer less vegetated portions of the riparian corridor (Harris, 2006). No information is available on overwintering habits, but frogs living near geothermal springs could be active year-­round. CALLING ACTIVITY AND MATE SE­L ECTION

The advertisement call consists of a series of quiet soft clucks. Recordings of the call are at: http://­www​ .­californiaherps​.­com​/­sounds​/­roncajrice1008​.­mp3. BREEDING SITES

Breeding occurs in shallow springs, pools, riverside marshes, and spring runs. ­Water should be lentic or slow moving. Emergent vegetation may be pre­sent, but it should not clog the entire wetland. REPRODUCTION

Calling takes place from January to April, with oviposition occurring usually during February and March. Bradford et al. (2005b) reported breeding in November. As such, this species may have a bimodal breeding season, with most breeding occurring in the spring. Egg masses are attached to vegetation in shallow ­water just below the ­water’s surface. Bradford et al. (2005b) reported “many hundred eggs” per mass. Embryos hatch in 5–7 days (Drake, 2010). LARVAL ECOLOGY

Nothing is reported in the lit­er­a­ture on the larval ecol­ogy of this species in nature, except that tadpoles may occasionally overwinter (RLFCT, 2016). However, tadpoles reared in the laboratory metamorphose at 2–3 months when raised at 24–25°C (Goldstein, 2007). Survivorship was 82% at 20°C, 94% at 25°C, and 66% at 30°C, with tadpoles raised at 25°C taking 67 days to metamorphose (Goldstein et al., 2017). Optimal temperatures for tadpole be­hav­ior ­were 25–30°C. Tadpoles likely feed on alga, and opportunistic oophagy has been reported (Drake, 2010). This species does not have any of the morphological characteristics of oophagous tadpoles.

Adult Lithobates onca. Photo: Dana Drake

mellifera scutellata) that sometimes resulted in mortality to the frog. Like other leopard frogs, they prob­ably consume a variety of invertebrates in proportion to their availability. PREDATION AND DEFENSE

Relict Leopard Frogs ­will sit motionless, hidden in riparian vegetation, but take to ­water at the approach of a predator; ­there, they seek shelter in vegetation or ­under rocks. Nothing has been reported concerning predators, but ­these undoubtedly include snakes (Thamnophis elegans), birds, and mammals. Tadpoles have been observed feeding on the eggs of conspecifics (Drake, 2010). POPULATION BIOLOGY

At Blue Point Spring, Bradford et al. (2004) estimated a 90% monthly survivorship rate during the course of their observations, which extrapolated to a 27% annual survivorship rate. Most populations of this species only contain a small number of adults. ­Under laboratory conditions, maturity is attained in 1 year (Malfatti, 1998). Some individuals attain at least 4 yrs of age.

DIET

Tanner (1931) reported a damselfly, a beetle, and a wasp from a single individual collected in June. Bennet et al. (2020) reported juveniles feeding on Africanized Bees (Apis

Tadpoles of Lithobates onca. Photo: Dana Drake

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DISEASES, PARASITES, AND MALFORMATIONS

Lithobates onca possesses antimicrobial skin peptides that aid in re­sis­tance to some bacteria (Escherichia coli, Candida albicans) but not ­others (Staphylococcus aureus) (Conlon et al., 2010). Although presented as L. pipiens, Parry and Grundman (1965) reported the nematodes Cosmocercella haberi and Physaloptera sp. from leopard frogs along Santa Clara Creek and the Virgin River in extreme southwestern Utah that ­were possibly L. onca. Other parasites included the cestode Cysticercus sp.; the trematodes Glypthelmins quieta, Haematoloechus coloradensis, and an unidentified Plagiorchiinae; the protozoans Chilomastix sp., Hexamita

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530  Ranidae

intestinalis, Karatomorpha swazyi, Nyctotherus cordiformis, Opalina sp., Trichomonas sp., and Zelleriella sp.; and the mite Hannemania hegeneri (Parry and Grundman, 1965). Amphibian chytrid fungus has been detected in Relict Leopard Frogs at 1 site, but the high geothermal ­water temperatures at L. onca sites may prevent the disease from having serious effects on the frogs (Jaeger et al., 2017). The situation is complicated by the strain of Bd used to challenge frogs. Relict Leopard Frogs show re­sis­tance to some strains of Bd when challenged, but Waddle (2017) demonstrated that L. onca was susceptible to local Bd strains from other frog species within its range. Still, frogs from within a currently Bd-­infected area cleared infections and survived at higher proportions than ­those from Bd-­free areas, dependent on life stage; recently metamorphosed frogs showed low survivorship when challenged with Bd. SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Small population size combined with ­limited heterozygosity and a small, fragmented range make the status of the species very precarious. Urgent conservation mea­sures ­were recommended as early as 1980, when the continued presence of leopard frogs along the Virgin River was recognized (Bury et al., 1980). Bradford et al. (2004) estimated the entire population as only 1,100 frogs (range 693–1,833) at the then 5 known sites. Seven sites are now known to contain this species (RLFCT, 2016). Jaeger and Rivera (2013, cited by RLFCT, 2016) estimated the population size in 2012 as 1,381–2,082 (mean 1,584) or 1,442–2,326 (mean 1,682) depending on which estimator was used. Populations undoubtedly ­were lost during dam building along the Colorado River, and populations along the Muddy and Virgin rivers are apparently extinct due to agricultural and ­water development. Threats to its continued survival include the introduction of nonindigenous predators (game fish, American Bullfrogs, crayfish), drought, encroachment from emergent vegetation into its spring habitats, and flash flooding. In this regard, restricting livestock from some springs actually may hasten the decline of the frog population, as livestock access prevents vegetation encroachment. The aggressive spread of tamarisk is likely detrimental to this species, and this highly invasive tree should be removed whenever encountered. A conservation agreement and strategy has been developed ­under the auspices of a complex of state and federal agencies (RLFCT, 2016). Bradford et al. (2005b) and RLFCT (2016) have reviewed the conservation mea­sures needed to ensure the survival of this species. ­There have been several efforts to translocate this species to other

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Habitat of Lithobates onca. Photo: Dana Drake

suitable sites, some of which have been successful (RLFCT, 2016). Current information suggests an increasing population with the establishment of frogs at translocated sites. In 2018, Relict Leopard Frogs ­were introduced into constructed pools as part of the Las Vegas Creek restoration proj­ect (the Springs Preserve) in Las Vegas, Nevada, home formerly to the Vegas Valley Leopard Frog (L. fisheri), now extirpated in this region. A nocturnal visual encounter survey in July 2018 recorded only 6 Relict Leopard Frogs. In April 2019, in situ reproduction was confirmed when hundreds of small tadpoles ­were observed in the ponds. Tadpoles began to metamorphose in July 2019. In October–­November 2019, 214 Relict Leopard Frogs ­were captured and marked in the ponds. Results from a subsequent survey suggested that 424 frogs inhabited the ponds. Relict Leopard Frogs reproduced at the Springs Preserve again in 2020. In September–­October 2020, 286 Relict Leopard Frogs (40 adults and 246 juveniles) ­were captured during mark-­recapture surveys, and the population was estimated to be 71 adults and 539 juveniles. Thus, a self-­sustaining population appears to have become established. Details of this proj­ect are in Saumure (2020) and Saumure et al. (2021). This species is considered a Sensitive Species in Nevada.

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Lithobates palustris 531

Lithobates palustris (LeConte, 1825) Pickerel Frog Grenouille des marais ETYMOLOGY

palustris: from the Latin paluster meaning ‘of the marsh.’ The name literally means ‘a marsh frog.’ NOMENCLATURE

Conant and Collins (1998): Rana palustris Dubois (2006): Lithobates (Lithobates) palustris Fouquette and Dubois (2014): Rana (Lithobates) palustris Synonyms: Rana pardalis, Rana palustris mansuetii IDENTIFICATION

Frogs of the Leopard Frog complex most closely resemble the Pickerel Frog in shape, size, and coloration. In Pickerel Frogs, the dorsal spots are squared rather than rounded, occur in 2 well-­defined parallel rows down the back between the dorsolateral folds, and are paired rather than scattered about. ­These frogs also have distinct yellowish to orange color on the underside of the thighs and groin, unlike the leopard frogs. The American Bullfrog lacks the dorsolateral folds altogether, and the Green Frog has only partial dorsolateral folds, and is unspotted. Adults. This is a medium to large, olive-­green to gray, tan, or brownish frog with a light-­to cream-­colored dorsolateral fold that extends down the body from ­behind the eye to the groin. Two to more than 4 parallel folds occur between the main dorsolateral folds on the back. ­There are large, paired, square dorsal spots on the back and sides; ­these spots (9–15 normally, but range 7–21) normally extend in 2 rows down the back between the dorsolateral folds. The rear limbs have black bars, giving a banded pattern. ­There is no white spot in the center of the tympanum, but a white line is pre­sent on the posterior part of the upper lip. The snout is pointed, and the eyes are large. The belly and throat are white, but the underside of the hind limbs and groin are yellow to orange. In some populations, mottling may be pre­sent on the ventral surface. Males have paired vocal sacs and enlarged thumbs with thickened pads to clasp the female during amplexus. They also may be slightly smaller and lighter colored than females. Adults are mature by 44 mm TL; maximum size is 87 mm SUL (Gibbs et al., 2007). Specific size mea­sure­ments are few in the lit­er­a­ture. In Delaware, males ­were a mean SUL of 58.6 mm (range 47–66 mm SUL) and weighed a mean of 16.7 g (range 8.0–23.75 g) (Given, 2005). In Ohio, males ­were 45–58 mm and females ­were 60–76 mm SUL (Walker, 1946), whereas in

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Connecticut males ­were 41–58 mm SUL (mean 50.2 mm) and females ­were 51–79 mm SUL (mean 60.8 mm) (Klemens, 1993). Paton (in Raithel, 2019) found males averaging 50–55 mm SUL and females 50–65 mm SUL in Rhode Island. Pennsylvania males averaged 55.1–54.9 mm SUL (range 40.9–66.5 mm) and females 67–69.1 mm SUL (range 54.1–87.4 mm) (Meshaka et al., 2012b, 2017b; Meshaka and Morales, 2020). In Nova Scotia, males ­were 49.0– 63.5 mm SUL (mean 56.3 mm) and females ­were 50.8– 74.6 mm SUL (mean 61.8 mm) (Gilhen, 1984). Males in Louisiana ­were significantly smaller (mean 52.5 mm, range 42–65 mm SUL) than females (mean 64.1 mm, range 48–75 mm SUL) (Hardy and Raymond, 1991). Larvae. The tadpole is large, deep, and full bodied. The dorsal color is olive green grading to yellow on the sides. The belly is cream colored with white blotches, whereas the dorsum is marked with fine black and yellow spots. The belly is iridescent, and the viscera are vis­i­ble, but often only slightly. The tail is very dark with black blotches and tail fins that are variously patterned. When viewed from above, the body is very round or oval. Tadpoles of L. palustris may be large, to 76 mm TL (Wright, 1929). The tadpoles of L. palustris, L. pipiens, L. clamitans, and L. catesbeianus are difficult to differentiate. The throat of the Northern Leopard Frog tadpole is more extensive and translucent than that of L. palustris. Tadpoles of L. palustris usually contain a yellow wash on the sides of the body. Tadpoles ­were described by Wright (1929) and Altig (1970), and the larval mouthparts ­were illustrated by Hinckley (1881), Wright (1914, 1929), and Dodd (2004). Eggs. The eggs are deposited in a firm, ­spherical cluster 38–100 mm in dia­meter; each cluster contains 32 mg/L), NTA (134.6 mg/L), and phenol (9.87 mg/L) (Birge et al., 1980). In a study examining the effects of oil runoff on frogs in Kentucky, ­there ­were about the same number of Pickerel Frog egg masses deposited in ponds receiving runoff from pastureland and oil brine pits as in reference ponds. However, sample sizes ­were small, making conclusions about the effects of ­these substances on Pickerel Frogs impossible (Westerman et al., 2003b). Death occurs in about 21 hrs at 30 mg/L of carbaryl, a broad-­spectrum insecticide, and it significantly reduces larval activity levels at 2.5 mg/L (Bridges and Semlitsch, 2000). Pickerel Frogs do not avoid areas where herbicides have been sprayed, particularly diquat dibromide (Reward®), glyphosate (Roundup®), and chelated copper that acts as an algaecide (Picone, 2015). STATUS AND CONSERVATION

Throughout much of its range, the Pickerel Frog is considered uncommon or rare (e.g., Cox, 1899; Brown, 1992; Hecnar and M’Closkey, 1997a; Hemesath, 1998; Mierzwa, 1998; Mossman et al., 1998; Platt et al., 1999; Sargent, 2000; Rice et al., 2001; Brodman, 2003; Baber et al., 2004; Florey and Mullin, 2005; Herrmann et al., 2005). ­Whether

Breeding habitat of Lithobates palustris. G ­ reat Smoky Mountains National Park, Tennessee. Photo: C.K. Dodd, Jr.

Dodd_Canada_int_5pgs_B4.indd 539

this reflects a naturally spotty distribution or is the result of long-­term habitat changes is largely unknown. Weir et al. (2014) suggested population declines in the Northeast based on road call surveys over an 11 yr period. Redmer (1998a) considered the species not to have declined in Illinois, except in the urbanized Chicago metropolitan area. The species is considered to have declined in Iowa (Christiansen, 1981) and to have experienced localized declines in Ontario and Québec but not New Brunswick or Nova Scotia (Weller and Green, 1997). The species has not been seen in Kansas since reported by Smith (1934; see Collins, 1993). In several areas, populations of Pickerel Frogs are isolated in fragmented habitats, such as on the small hills and mountains of southern Québec that are surrounded by agriculture and urban habitats (Ouellet et al., 2005b). ­These Frogs may survive in areas with moderate to high housing densities, but abundance is reduced (Picone, 2015). The Pickerel Frog is a forest species when not at breeding ponds, and it ­will move along cool streambeds when dispersing. It is not prone to moving through open areas such as ­those created by transportation corridors. Thus, the presence of roads near breeding ponds influences dispersal by this species and act to filter movement across a landscape (Gibbs, 1998a). Still, Marsh et al. (2017) found a positive relationship between road density and population distribution at large spatial scales. Distribution was not affected by impervious surface cover. Mortality by highway traffic was reported by Glista et al. (2008). Unfortunately, the species does not readily colonize restored wetlands (Shulse et al., 2010). Mitchell (2016) found this species at only 8% of agriculturally restored ponds in the Mid-­Atlantic region. In habitats invaded by the nonnative shrub Lonicera maackii, overall amphibian species richness and evenness decreases. Microclimatic temperature is affected, and counts of Pickerel Frogs decrease in habitats invaded by this species (Watling et al., 2011b). High saponin (amphipathic glycosides) concentrations (>15 mg/L-1) from the introduced common reed (Phragmites australis) reduced survival, developmental rate, and size of L. palustris larvae in experimental outdoor mesocosms (Martin and Blossey, 2013). Purified tannins, however, had no effect on larval survival, final developmental stage, or final SUL. Also in contrast, Ritten­house (2011) found no effects on Pickerel Frog larval growth and development from the introduced reed canary grass (Phalaris arundinacea). ­Because Pickerel Frog populations are spottily distributed, it is impor­tant to identify and monitor them to ensure their long-­term per­sis­tence. Pickerel Frogs have been detected using call surveys in conjunction with regional monitoring programs. In Tennessee, for example, they ­were heard at 79 of 321 calling stations. Listening for 5 or 10

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540  RANIDAE

minutes made no difference in terms of an observer’s ability to detect this species (Burton et al., 2006). However, detection of L. palustris during call surveys was exceedingly uncommon in Québec and Ontario (Bishop et al., 1997; Lepage et al., 1997) and elsewhere, suggesting other protocols for monitoring might be necessary for this

species. Skelly et al. (2003) noted that the probability of determining occupancy for L. palustris was low (20%) for surveys conducted annually, and that long-­term resurveys of historical sites are necessary to determine the presence and per­sis­tence of this species. This species is considered of Special Concern in Michigan.

Lithobates pipiens (Schreber, 1782) Northern Leopard Frog Grenouille léopard du Nord

“Rana pipiens,” the applicability of the information to e­ very species within the complex is questionable. When consulting much of the scientific lit­er­a­ture prior to the early 1970s, readers are cautioned to remember that results and interpretations of numerous past studies involving “leopard frogs,” even when localities or sources of individuals are provided, are based on assumptions of clinal or geographic variation rather than presently recognized species-­specific differences.

ETYMOLOGY

pipiens: from the Latin pipiens, meaning ‘peeping.’ Apparently the first collector of this species heard Spring Peepers and thought that the leopard frogs he had just collected ­were responsible for the loud ‘peeps’ he heard. NOMENCLATURE

Conant and Collins (1998) and Stebbins (2003): Rana pipiens Dubois (2006): Lithobates (Lithobates) pipiens Hillis (2007): Rana (Pantherana) pipiens Fouquette and Dubois (2014): Rana (Lithobates) pipiens Synonyms: Rana brachycephala, Rana burnsi, Rana burnsorum, Rana halecina, Rana kandiyohi, Rana noblei, Rana pipiens brachycephala, Rana virescens brachycephala ­There are many scientific publications in which the identification of “leopard frogs” has been confused among the 9 currently recognized species within the United States and Canada. This is especially true in physiology and toxicology, where it may be impossible to determine which species was studied. For example, Bachmann (1969) discussed the influence of temperature on development in “Rana pipiens,” yet included frogs from New Jersey, Vermont, Texas, Florida, Costa Rica, and Mexico in his analy­sis. Other studies have relied on frogs from commercial suppliers (Bagnara and Frost, 1977), and it cannot be assumed that “leopard frogs” ­were collected in proximity to the supplier. As such, ­there is confusion about species identity in the following publications: Moore (1949b), Volpe (1957a), Bresler (1963, 1964), Kaplan and Overpeck (1964), Kaplan and Glaczenski (1965), McLaren (1965), Licht (1967), Levine and Nye (1977), Cole and Casida (1983), Wygoda (1984), Dial and Bauer (1984), and Dial and Bauer Dial (1987). Accounts of this species also may be confused with the recently described L. kauffeldi (Schlesinger et al., 2017). If the original lit­er­a­ture does not specify the origin of

Dodd_Canada_int_5pgs_B4.indd 540

IDENTIFICATION

Adults. This is a medium to large, olive-­green or brown frog with smooth skin. It possesses a yellow to cream-­colored continuous dorsolateral fold that extends the entire length of the body from the snout to the groin. ­There are scattered, large, dorsal, unpaired, oval or roundish spots on the head, back, and sides; ­these spots may extend in 2 or 3 irregular rows down the back between the dorsolateral folds. ­There are usually 0–1 spots between the posterior extent of the eyes and the snout when viewed dorsally (Bresler, 1963). The rear limbs, especially, have black bars giving a banded pattern. ­There is a white spot in the center of the tympanum, but the spot may be unclear or inconspicuous, and a white line (supralabial stripe) is pre­sent on the upper lip. The snout is pointed, and the eyes are large. The belly, throat, and underside of the hind limbs are white. Sexual dimorphism among Northern Leopard Frogs is not apparent, except during the breeding season when males have paired lateral vocal sacs (evidenced by loose skin between the ­angle of the jaw and forearm) and swollen thumbs to assist with amplexing females. Males also may be less patterned than females on the head and dorsal regions but more patterned on the lateral and ventral surfaces (Schueler, 1982a). Unlike some other members of the Leopard Frog complex, males possess oviducts. Females are usually swollen with eggs during the breeding season, but ­there is no other distinguishing external characteristic. Females are also usually larger than males, but differences are slight. Schueler (1979, 1982a) provided an extensive analy­sis of pattern variation in L. pipiens in Canada and the northeastern United States. He concluded that leopard frogs from warmer and moister climates tended to be darker than ­those

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Lithobates pipiens 541

from cooler and drier climates; variation in dermal secretory glands is in­de­pen­dent of pattern, but ­these glands are more extensive where ­water is widespread, as well as in the North; linear patterns in coloration and reduced leg spotting are associated with field, lake edge, and marsh habitats; variation in spot number varies considerably within populations, but not between populations; and among-­population variation in spot number is inversely correlated with spot area. In addition, the brown morph tended to be associated with marshes and lakes in the North, whereas the green morph was found most often in boreal forest. ­Whether ­these trends apply to other populations is unknown. Adults are 51–111 mm SUL. Specific size rec­ords include: a mean of 67 mm SUL for females ­after egg deposition (range 55–80 mm SUL) and 63.8 mm SUL for males (range 55–75 mm SUL) in Wisconsin, whereas body mass averaged 27.8 g (range 17–48 g) for females and 25.8 g for males (range 17–39 g) (Hine et al., 1981); females from 63–90 mm SUL and males 58–77 mm in Ohio (Walker, 1946); a mean of 60.3 mm SUL for females and 58.2 mm for males in West ­Virginia (Sutton, 2004); males 51–65 (mean 57.2) mm SUL and females 53–65 (mean 57) mm in Connecticut (Klemens, 1993); females 49–95.3 mm (mean 71.5) and males 47.1– 79.1 mm (mean 65.8) in Pennsylvania (Meshaka et al., 20111b); mean SUL in males of 68.3 mm and for females 74.2 mm in New Mexico (in Degenhardt et al., 1996); males 57–78 mm SUL and females 66–95.3 mm in Nova Scotia (Gilhen, 1984); and males 58–78 mm SUL (mean 69.4 mm) and females 60–91 mm (mean 80.2 mm) on Prince Edward Island (Cook, 1967). Matsuda et al. (2006) give maximum sizes of 111 mm SUL for females and 80 mm SUL for males. Adult albinos have been reported from New York and Wisconsin (Hensley, 1959) and South Dakota (Browder, 1972). Other unusual color variations include melanistic frogs of both the wild and Burnsi phenotypes (Richards et al., 1969; Richards and Nace, 1983) and blue (axanthic) leopard frogs (Berns, 1966; Berns and Uhler, 1966; Black, 1967). Larvae. The tadpole is large (normally 50 km apart may exhibit significant differentiation, primarily due to ge­ne­tic drift. This diversity is most evident in Ontario and Manitoba and tends to decrease among more

Dodd_Canada_int_5pgs_B4.indd 544

western populations (see above). However, the ge­ne­tic structure of populations in the northern Prairie Pothole Region of North Dakota over a 68 km linear study area was rather homogenous based on an examination of 6 microsatellite loci and accompanying Bayesian assignment tests (Mushet et al., 2013); heterozygosity and allelic richness ­were high. ­These results suggest that Northern Leopard Frog prairie populations are panmictic and maintain a high level of ge­ne­tic diversity. Wilson et al. (2008a) suggested that all populations in the far North share a close evolutionary history and belong to a western haplotype group. ­These authors found that ge­ne­tic diversity declined westward in contrast to Mushet et al. (2013), with the least diversity occurring at the periphery of the species’ range in the northwest. This low diversity could result from an increasingly dry habitat gradient that results in a scarcity of overwintering sites and drought refugia (Mushet et al., 2013). In any case, peripheral populations generally have lower diversity than more centralized populations. Taken together, ­these results have conservation implications, especially when man­ag­ers consider repatriating Northern Leopard Frogs to depleted Alberta localities, for example. Source populations with similar haplotypes may not be readily available. Lithobates pipiens hybridizes with a number of other members of the L. pipiens complex and other Lithobates in the laboratory, but most hybrid crosses are not ­viable (reviewed by Hillis, 1988, and references therein). Development may appear normal in crosses with L. areolatus, but not ­after metamorphosis; hybrid inferiority (increased mortality, delayed development, defects, reduced fertility) occurs in crosses with L. blairi, L. forreri, L. magnaocularis, L. megapoda, L. neovolcanica, L. palustris, L. sphenocepha­ lus, L. taylori, and L. yavapaiensis; severe hybrid inferiority (high mortality, sterility) occurs with some L. berlandieri, L. neovolcanica, and L. sphenocephalus; complete hybrid incompatibility (no appreciable development) occurs with other L. berlandieri populations. In contrast, Gillis (1975) could find no significant ge­ne­tic incompatibility between L. pipiens and L. blairi. Based on hybridization experiments, Ruibal (1962) considered the L. pipiens population at San Felipe Creek, California, to be more closely related to species farther south in Mexico and Central Amer­i­ca than to “cold races” farther north; this and other California desert populations, no longer extant, are now referred to L. yavapaiensis ( Jennings and Hayes, 1994b). Natu­ral hybrids may be found where members of the complex co-­occur. For example, L. pipi­ ens × L. chiricahuensis hybrids occur at a rate of about 9% in some Arizona populations (Frost and Platz, 1983). Lithobates pipiens × L. blairi hybrids occur at a rate of

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Lithobates pipiens 545

1–16% in overlapping Nebraska populations (Lynch, 1978), and at 1–4.4% in Iowa and South Dakota populations (Dunlap and Kruse, 1976). Throughout much of the area where contact occurs, ­there has been asymmetrical ge­ne­tic swamping of L. pipiens nuclear haplotypes by L. blairi haplotypes (Di Candia and Routman, 2007). Other hybrid populations have been reported from Illinois (Smith, 1961), Colorado (Post, 1972; Gillis, 1975), and New Mexico (Degenhardt et al., 1996). The percentage of populations in a region that contain hybrids may change through time, perhaps reflecting the extent of interaction between the parental species (Cousineau and Rogers, 1991). Two polymorphic phenotypes, reflected in both adults and larvae, are often observed in the same breeding ponds in some areas. In West ­Virginia, about 67% of the leopard frogs are green and 33% are brown (Sutton, 2004). Corn (1981) noted that the brown morph was pre­sent in about 24–68% of Northern Leopard Frog populations in northern Colorado, and that brown morph larvae had a shorter time to metamorphosis than sympatric green morph larvae (68 vs. 74 days and 83 vs. 86 days in 2 populations). The brown morph is more common than the green morph in early season metamorphs. Corn (1981) suggested that the brown morph was selectively favored over the green morph in ponds experiencing high rates of predation. The color pattern appears genet­ically stable, with the green morph a ­simple dominant to the brown morph (Fogleman et al., 1980). In the Sheyenne National Grasslands of North Dakota, the green morph was stable in frequency (38.6%) over the period of 2001–2010 (Gustafson et al., 2019). ­There are 2 prominent unusual phenotypes within L. pipiens, the unpatterned “Burnsi” (Brown and Funk, 1977) and the distinctively mottled “Kandiyohi” (both described as species by Weed, 1922). The Burnsi phenotype is found in western Wisconsin, central and north central Minnesota, eastern South Dakota, and southeastern North Dakota; it is also reported from Dickinson County, Iowa, and from Maine (Lindemann et al., 2019a). It occurs at variable frequencies among dif­fer­ent populations. At Block Lake, Minnesota, for example, the frequency of Burnsi frogs was 11.3% from 1967 to 1986, and 18.5% from 1985 to 1999 (McKinnell et al., 2005). At populations in Minnesota, North Dakota, and South Dakota, most frequencies are 40–60% of the sites in Ontario over a period of several years (Hecnar, 1997; Hecnar and M’Closkey, 1996a, 1997a, 1998), at 100% of 27 sites in an agricultural area of Iowa (Swanson et al., 2019), and at 50% of 210 ponds

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sampled over an 11 yr period in Missouri (Drake et al., 2015). ­There is considerable annual turnover rate in wetland occupancy from one year to the next. As might be expected ­because of its dispersal capability and generalist habitat requirements, it was one of the species least likely to show nested-­assemblage occupancy patterns (i.e., subsets of co-­occurring species) when compared with other species found in the same region (Hecnar and M’Closkey, 1997a). Lithobates pipiens is occasionally found in springs (Allen, 1963), along sandy beaches of large freshwater lakes (Stille, 1952), and at the entrances to caves (Garton et al., 1993). TERRESTRIAL ECOLOGY

Northern Leopard Frogs are commonly found long distances from ­water. During the summer, adult Northern Leopard Frogs often are found in distant terrestrial habitats, where they sit concealed in “forms” on the moist forest floor or in grassy, moist meadows. Older common names, the grass or meadow frog, refer to its terrestrial habitat preferences. They frequent shallow freshwater marshes, grassy-­sedge woods, old fields, wet meadows, unmowed pasture, and even hayfields (Merrell, 1970, 1977; Hine et al., 1981). Beauregard and Leclair (1988) found that preferred habitats in Québec included areas close to a marsh with tall herbaceous vegetation of high species richness and a low percentage of moss cover. The primary ­factors affecting terrestrial habitat se­lection are the levels of moisture in leaf litter and soil, a nearness to ­water, cover, and temperature (Blomquist and Hunter, 2009). Northern Leopard Frogs usually select the warmer habitats when provided a choice. Frogs may also use crevices or other cavities as retreat sites in drier situations or when moisture conditions change. Frogs remain in their secretive terrestrial locations for less than 24 hrs to more than 5 days (Dole, 1965a). Most shifts in terrestrial locations are from 5 to 10 m, with movement occurring usually at night or on overcast days and during favorable moisture conditions (rain, heavy dews) rather than during periods of dry weather. Northern Leopard Frogs may use the same form or retreat site multiple times, and they usually remain within a defined home range during this period. In Maine, mean home ranges varied between 348 and 1,347 m2 depending on location. Individual home ranges varied between 13 and 8,425 m2 (mean 1,096 m2), and did not vary by sex (Blomquist and Hunter, 2009). In Iowa, Swanson et al. (2018) reported that males had smaller total home ranges (median 4,740 m2) than females (median 6,423 m2), but that male frogs’ total movements (median 553 m vs. 389 m) and average daily movements (median 18 m vs. 16 m) ­were greater than female movements. Median core home ranges ­were 3,154 m2 for males and 1,347 m2 for females.

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The fidelity of frogs to a home range varies, depending on the amount of habitat available and its proximity to ­water bodies. If ­little habitat is available, frogs tend to return to terrestrial feeding areas; if much habitat is available, frogs may disperse more widely and not return to the same area as in the previous activity season (Dole, 1965b). Smaller frogs tend to occupy home ranges in wetter areas than larger frogs. Dole (1965b) estimated home ranges at 1 spatially confined site that ­were considerably smaller (males 78 m2, females 113 m2, subadults 68 m2) than ­those found in more extensive habitats (males 362 m2, females 503 m2, subadults 283 m2). ­There was considerable individual variation, however. Reports in the lit­er­a­ture stating that L. pipiens does not occupy home ranges (Fitch, 1958) likely refer to other members of the Leopard Frog complex. Occasionally, adult frogs may make extended journeys (100–240 m; Dole, 1965a) to adjacent habitats. Long-­distance movements usually occur during nocturnal rains and appear directed, that is, in a straight line without the wandering paths seen during short movements within the home range. Northern Leopard Frogs may move as much as 47 m/hr during long-­ distance dispersal events (Dole, 1965a). ­These frogs then remain at the distant locations for several days before returning to the original home range. Long-­distance movements usually occur on only 1 night, although Dole (1965a) recorded 1 frog that made long-­distance movements on 2 consecutive nights. Such periodic movements by anurans have been documented in a number of species and may facilitate familiarity with adjacent areas. Extended rainfall events and warm temperatures allow for wide dispersal in terrestrial habitats. Displaced Northern Leopard Frogs are capable of returning to near the place they ­were captured when displacement distances do not exceed 1 km. At greater distances, they are unable to move back ­toward the original site and disperse more randomly. Homing movements are fairly direct, as frogs immediately orient ­toward the home direction following release; they adjust their movements homeward the farther they travel. Blind frogs are also able to orient ­toward the home direction when displaced at much lesser distances, although their movements are not as direct. Blind frogs are able to navigate at night and upwind from their release point, as are anosmic animals released in heavy fog, suggesting that vision, brightness, and olfaction may not be required for homing be­hav­ior. In additional experiments, displaced anosmic or deaf leopard frogs ­were able to home (Dole, 1972b). Dole (1968) suggested that homing ability may result from prior familiarity with the habitat surrounding a wetland, or perhaps a combination of general olfactory cues, rather than specific cues from a par­tic­u­lar site. Lithobates pipiens is tolerant of wide variations in ambient temperature. The CTmax of adults is 38–39°C, depending

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upon acclimation temperature. This is not surprising, as Northern Leopard Frogs are heliotropic. Animals acclimated at longer photoperiods also tend to have a slightly higher CTmax than frogs acclimated at shorter or 12:12 photoperiods (Mahoney and Hutchison, 1969). Tolerance for a wide range of temperatures allows for an extended activity season; for example, activity extends from February to mid-­March to late October or even November in upstate New York, Pennsylva­ ngland (Wright, 1914; Klemens, 1993; nia, and New E Meshaka et al., 2011b; Raithel, 2019), and to mid-­October during warm weather in South Dakota (Bubac et al., 2017). Like many aquatic and semiaquatic amphibians, Northern Leopard Frogs are prone to lose body ­water while on land. Smaller frogs are more tolerant of ­water loss per se, but they lose ­water at a greater rate ­because of an increase in surface to volume ratio (Thorson, 1955). Rehydration occurs in about 3 hrs for a frog desiccated to a loss of 27–29% body weight, and in 48 hrs for a frog desiccated to 65–75% of its hydrated weight, as long as the soil moisture is at least 20%. Rehydration can occur at lower soil moisture contents, but not to maximum pre-­dehydration weight (Dole, 1967a). Adults resorb moisture from soil or from wet vegetation through the highly vascularized skin of the groin (termed the pelvic patch). Northern Leopard Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities ( Jaeger and Hailman, 1971, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”), which apparently helps them orient ­toward areas of increasing illumination, such as the open area above a pond (Hailman and Jaeger, 1974). Northern Leopard Frogs likely have true color vision. AQUATIC ECOLOGY

Lithobates pipiens is frequently found in or near ­water, even during the nonbreeding season. Indeed, Hahn (1968) recorded leopard frogs active in January and February around warm springs in Colorado, and Olson (2010) found a Northern Leopard Frog active in December in northern Wisconsin on ice along the shore. Whereas many frogs disperse, ­others choose to remain in or around ponds and wetlands. They are heliothermic, basking in the direct sun while sitting on floating vegetation or along the wetland margins. Temperature is controlled by evaporative cooling. Brattstrom (1963) recorded body temperatures ranging from 18 to 33.5°C, and stated that on cool nights, Northern Leopard Frogs moved to the warmer parts of a pond when ­water temperatures exceeded air temperatures or sheltered in small depressions along the shore to reduce the effects of air flow.

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Northern Leopard Frogs overwinter in lakes, near the outlets of lakes or other bodies of ­water, and along stream beds. Overwintering occurs in lakes and streams deep enough not to freeze solid (but see Blais, 2017). Dormancy occurs openly on the lake bottom, or in vegetation, detritus, mud, or other debris on lake or stream substrates. In Ontario, they ­were found beneath 13–40 cm dia­meter rock rubble in a stream >85 cm in ­water depth with a velocity of 22.5 cm/sec (Cunjak, 1986); winter ­water temperatures ­were between 0.5 and 2.1°C. ­There are reports of leopard frogs resting in shallow pits on the muddy bottoms of lakes, with a light coating of silt over them (Emery et al., 1972), or wedged into crevices or even ­under turtles (Ultsch et al., 2000). Emery et al. (1972) recorded them as deep as 3.1 m below the ice. In neither of the latter instances ­were the frogs concealed, as access to well-­oxygenated ­water took pre­ce­ dence over midwinter predator avoidance. Terrestrial dormancy may not be common, and Northern Leopard Frogs have been shown to choose ­water over leaf litter on land when laboratory tested at a temperature of 1.5°C (Licht, 1991). However, Waye (2001) reported 2 radio-­ tracked leopard frogs in British Columbia that moved to small mammal tunnels and apparently overwintered ­there, Wright (1914) reported dormancy ­under stones in ravines, and Badje et al. (2016) recorded a single individual in a crayfish burrow. When dormant, leopard frogs lie motionless on the substrate with legs and arms extended, and make no effort to move in the deep cold ­water; they are capable of sluggish movement if disturbed, and may attempt to burrow farther into the substrate (McAdam and Nagel-­Hisey, 1998). ­There also are reports of leopard frogs being active ­under the ice in shallow rivers with cobble substrates, where they are capable of movement at 0°C (Kreitals et al., 2014). Leopard frogs prefer dormancy sites that are well oxygenated. For this reason, they often overwinter near lake outlets or spillways where flowing ­water supplies fresh oxygen, and ­there is a ­limited chance of oxygen depletion. Ultsch et al. (2004) demonstrated that Northern Leopard Frogs are tolerant of a large decrease in oxygen tension (the critical PO2 at which anaerobic respiration occurs), which permits them to remain in hypoxic icebound ponds for extended periods. Mass winterkills due to oxygen depletion (anoxia) are not uncommon (Manion and Cory, 1952), although for some reason, the Burnsi phenotype of this species is associated with an increased ability to survive low oxygen levels during winter dormancy (Merrell and Rodell, 1968). Frogs may remain dormant for long periods. For example, in Minnesota they remain underwater from October ­until March. Unlike other northern frog species, Northern Leopard Frogs are unable to survive freezing temperatures at the

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ground surface (Schmid, 1982), and significant mortality may occur if Northern Leopard Frogs are exposed to unusually cold weather. Cold fronts may kill frogs if the fronts occur ­earlier than normal and catch individuals moving to overwintering locations (Merrell, 1977), or if they occur in the spring as frogs are beginning to leave protected sites and move to breeding ponds. Mortality from oxygen depletion also occurs if snowpack completely blocks sunlight, preventing aquatic photosynthesis. Prolonged exposure to sublethal levels of cold weather reduces the immune response in L. pipiens (Maniero and Carey, 1997), which could make them more susceptible to pathogens or toxic chemicals. The immune response is rapidly restored as frogs warm with increasing ambient temperatures. CALLING ACTIVITY AND MATE SE­L ECTION

Northern Leopard Frogs may emerge from winter habitats as long as a month before calling begins. In West ­Virginia, for example, they have been observed to emerge as early as the first week in February, even though calling did not begin ­until mid-­March (Sutton, 2004). Ohio individuals also have been found in February (Walker, 1946), and they have been observed in January in Indiana during an exceptional warm spell (Minton, 2001). In Ohio, Zenisek (1963) noted emergence in late March and amplexus by 7 April, but no egg masses ­until 15 April. Wright (1914) reported emergences from 3 to 18 (mean 7) days prior to calling. Breeding in Northern Leopard Frogs occurs during the cool spring months of the year, a time that varies by latitude and elevation. The frogs may be active for several weeks prior to breeding. In the southern portion of the species’ range, breeding commences in late winter, whereas breeding usually occurs in spring and into the early summer in more northern localities. In the Chihuahuan Desert, eggs or very small tadpoles have been found from April to July, and even in September and October (Scott and Jennings, 1985). Some calling months include: beginning in March in Ontario (Seburn, 2013), mid-­March to mid-­April, depending on temperature, in Wisconsin (Hine et al., 1981), Michigan (Cummins, 1920), New ­England (Klemens, 1993), and West ­Virginia (Sutton, 2004); March–­May in Kentucky (Barbour, 1971); March–­early June in Colorado, depending on elevation and year (Post, 1972; Hammerson, 1999); mid-­ April to mid-­May in Nova Scotia (Gilhen, 1984), Québec (Lepage et al., 1997), and South Dakota (Kiesow, 2006); mid-­April to late May in South Dakota (Ernst, 2001); late April–­early May in Michigan (Dole, 1967b); April–­June in New Brunswick (Gorham, 1964) and Prince Edward Island (Cook, 1967); early May–­early July in North Dakota (Bowers et al., 1998); and mid-­May to June in northern Ontario (Schueler, 1973). Wright (1914) reported that males

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would occasionally call well ­after the breeding season, such as during rainfall or on cloudy days. The latest he heard a call was on 14 September; September calling is also reported by Walker (1946). Climate change appears to be affecting the initiation of emergence and calling at the northern portions of the species’ range in Ontario. For example, Klaus (2012) recorded ­earlier emergence, by 22 days, based on a 40 yr historical data set. With increasing temperatures, Northern Leopard Frogs are calling ­earlier than they used to (by 37.5 days; Klaus, 2012), a significant trend that has occurred over a very short period of time as spring air temperatures have increased by 2.8°C in southeastern Ontario (Klaus, 2012; Walpole et al., 2012). Increasing air temperature (13–14°C in Minnesota; Merrell, 1977) stimulates adult Northern Leopard Frogs to move from overwintering sites to breeding ponds. Males are first to arrive at breeding locations, with females arriving 3–14 (normally 5–7) days ­after calling begins. In the upper Midwest, males begin to call when ­water and air temperatures reach about 10°C. When air temperatures drop below 10°C, calling activity decreases even if ­water temperature is above 10°C (Hine et al., 1981), although Ernst (2001) reported calling near 0°C. In Alberta, calling is associated with ­water temperature (>7.5–8°C, but as low as 5°C) and the timing of the ice melt, but not with photoperiod (Sommers et al., 2018). Calling is negatively associated with wind speed and humidity. Even within close proximity, calling does not begin at all ponds at the same time. Sommers et al. (2018) found that initiation of calling varied by as much as 13 days even among nearby sites. Peak daily periods of calling (i.e., morning, midday, eve­ning) also vary among sites. ­Because daytime temperatures may remain much more favorable than cool nocturnal temperatures, calling and reproduction are often diurnal in northern populations in the early spring, although nocturnal calling occurs throughout the season (Wright, 1914; Cummins, 1920; Sommers et al., 2018). Frogs generally occupy the same calling position, and movements while at the breeding site are minimal. Short movements position adults in warmer locations from which to call, especially on cool or partially overcast days. ­There is no evidence of territoriality among males. Leopard frogs are quite vocal and make a variety of sounds ­under dif­fer­ent circumstances (Schmidt, 1968). The mating call of the Northern Leopard Frog, using its paired vocal sacs, has been described as a “staccato snore” (Frost and Platz, 1983), “ir-­a-­a-­a—­a-­a-­h” (Noble and Aronson, 1942), a “long low guttural croak” (Wright and Wright, 1949), a “guttural snore lasting approximately 3 sec, followed by grunts, squeals, and several clucking notes” (Ernst, 2001), and a chuckle lasting several seconds. Schmidt (1968) reviewed the vari­ous

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calls of the Northern Leopard Frog, and restricted the “chuckle” to a series of sounds following the mating call. The dominant frequency of the mating call is 1.5 kHz, with a duration of 2–4 secs. The pulse rate is temperature dependent. For example, it is 21 pulses/sec at 17–19°C in Arizona, 14 pulses/sec at 12–16°C in Colorado, 30 pulses/sec at 21°C in South Dakota, and 19 pulses/sec at 18°C in Illinois (Littlejohn and Oldham, 1968; Brown and Brown, 1972a; Dunlap and Kruse, 1976; Frost and Platz, 1983). Frost and Platz (1983) provide a sonogram comparing this species’ advertisement call with ­those of other Southwest leopard frogs. In contrast, Schlesinger et al. (2017) compared the sonogram of this species with other Northeastern species. They found call lengths of 1604–2409 ms, call rates of 0.06–0.08 calls/s, and dominant frequencies of 1,098–1,328 Hz at 18°C. Males call while floating on the ­water’s surface with limbs outstretched, and they vigorously pursue any approaching frog (Merrell, 1977). Occasionally, frogs may call out of ­water or from the bottom of a pond. When temperatures are high (20°C), males are unwary during courtship, but if temperatures decrease, they become more secretive in their calling be­hav­ior. Both males and females are approached by a reproductive male; if amplexed, males give a release croak that ­causes the primary male to release the amplexed male. Females likely choose a male by moving ­toward or near him, but they are concealed by vegetation so as not to attract other nearby males. Males readily grasp the rotund females, a body form that indicates a full complement of eggs. Amplexus is pectoral. The male arches his body convexly to fit tightly to the female; the hind limbs are flexed; and the front limbs encircle the female so that the digits are firmly holding the female’s venter. Gravid females remain ­silent when amplexed, as a distended body apparently inhibits the release croak (Diakow, 1977). Females are usually unable to dislodge an amorous male ­because of the male’s tenacity and his ability to grasp her with his swollen thumb pads. Spent females also have a release croak, which appears to be initiated through stimulation of the skin on the body’s trunk (Diakow, 1977). ­There are subtle differences in the release croaks of males and females (e.g., in pulse rate, amplitude, call rate) despite their similar functions, and males tend to call more often than females when employing the release croak (McClelland and Wilczynski, 1989). Amplexus lasts from several minutes to as long as a day, at least ­under laboratory conditions (Wright, 1914; Noble and Aronson, 1942). ­After some time following initial amplexus, the female begins to move backward in position in relation to the position of the male (“backward shuffling”), and extends her thighs backward and laterally in a 45° ­angle. The female then shifts her hind legs and feet into an oviposition

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posture forming a diamond-­shaped enclosure. The male rotates his hind limbs downward and outward, at which time the pair is in the egg-­laying posture. The female begins oviposition through a series of contractions of her abdominal walls, followed by arching her back. The male extends his legs and flexes his body convexly, thus bringing the ­couple’s cloacae into close proximity. As a cluster of eggs is extruded from the female’s cloaca, the male fertilizes them (termed ejaculatory pumps). Oviposition usually involves 10–23 distinct cycles of ejaculatory pumps. Total oviposition takes 2–8 minutes. Males usually release females within 6 minutes ­after oviposition is completed. Noble and Aronson (1942) provide a detailed description of the entire sequence. Males sometimes amplex inanimate objects (Merrell, 1977; Livo, 1981a), other frog species (Wright, 1914; Brown and Pierce, 1965), or even fish. Males amplexing spent females release them rapidly, presumably ­because the slender bodies of the females indicate previous oviposition and ­because of the female’s release croak. Noble and Aronson (1942) describe the contexts of the vari­ous types of release croaks issued by this species. BREEDING SITES

Northern Leopard Frogs choose clear ­water breeding sites that range from small (2.8 ha; Fischer, 1998) and man-­made ­water bodies such as gravel pits and golf course ponds (Mifsud and Mifsud, 2008); some individuals may breed in the quiet backwater of streams and rivers, or in clear, sand-­bottomed streams (Lynch, 1978). Frogs appear selective in their choice of available sites, avoiding both large and small wetlands, and they prefer ponds that warm rapidly in spring when compared to deeper and thus cooler ponds. ­Water depths are usually from 1.5 to 3 m, with bottom substrates of silt, muck, and decaying vegetation. Northern Leopard Frogs prefer breeding sites with >50% cover of the ­water’s surface by submerged and emergent vegetation and with extensive vegetation (cattails, grasses, willows) along the shoreline. The types of plants include Phalaris arundinacaea, Cornus stolonifera, Lythrium salicaria, Scirpus fluviatilis, Ambrosia artemisiifolia, and Poa sp. (Gilbert et al., 1994). Vegetation provides both protection and a place to attach the egg mass. Hine et al. (1981) suggested that open ­water allowed the pond to warm up faster than sites ­under canopy cover, and they noted that most reproduction takes place in the warmer sections of a breeding pond. However, Fischer (1998) reported nearly equal ­percent occupancy in ponds that ­were open canopied (70% cover). Fishless ponds are preferred,

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although leopard frogs are found with both predatory and nonpredatory fishes (Hecnar, 1997; Hecnar and M’Closkey, 1997b; Herwig et al., 2013; Lannoo and Stiles, 2020b). Data on breeding site preference are in Merrell (1968, 1977), Collins and Wilbur (1979), and Fischer (1998). As with many species of frogs, Northern Leopard Frogs do not inhabit all sites that appear suitable to a ­human observer. In Wisconsin, for example, occupancy ranged from 16% in 1975 to 32% in 1978 of 83 ponds surveyed, with breeding occurring in only 6% of ponds in 1975 and 23% in 1978 (Hine et al., 1981). Skelly et al. (2003) reported only a 20% probability of detecting this species on the E.S. George Reserve in Michigan over a 5 yr period at 32 sampled ponds. ­These low numbers may represent a declining regional population, however. Brodman (2009) also noted considerable variation in annual occupancy over a 14 yr study in Indiana. Lithobates pipiens do not necessarily use the same pond from one year to the next, but instead choose from among a suite of nearby ponds. The pond actually used in any one year reflects the weather conditions or other environmental variables within the region; a pond suitable one year may not be suitable the next, for example, ­because of drought conditions. During moderate to wet years in Iowa, Northern Leopard Frogs breed in temporary or semipermanent wetlands, but shift to permanent wetlands during periods of drought (Lannoo and Stiles, 2020b). They also may change breeding sites annually, presumably in response to weather conditions. For example, Lannoo and Stiles (2020b) captured tadpoles at 50 of 99 sampled sites in 2014 (a drought year in Iowa), and at 80 of 118 wetlands in 2012 (a wet year). Tadpoles ­were found at the same sites at only 35 wetlands, with 16 of 38 wetlands colonized in 2014 that ­were not occupied in 2012; frogs abandoned 45 of 80 wetlands occupied in 2012. Other ­factors also may play a role in breeding site se­lection. In Ontario for example, Hecnar and M’Closkey (1996b) found that Northern Leopard Frogs preferred sites with lower nutrient contents. In prairie regions, ponds in grasslands are preferred over ­those in parklands (Herwig et al., 2013). REPRODUCTION

The timing of reproduction varies geo­graph­i­cally, as might be expected for such a wide-­ranging species occupying varying elevations. In Minnesota, the breeding season extends from mid-­March to mid-­May, but occurs mostly in April (Merrell, 1977). In Arizona, eggs are observed from mid-­April to early June (Frost and Platz, 1983), but ­these populations are at higher elevations than ­those of other members of the L. pipiens complex directly to the west in California. Other

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dates for breeding are mid-­March to June (Illinois: P.W. Smith, 1947); mid-­March–­late April (Ohio: Walker, 1946); late March–­April (Colorado: Gillis, 1975; Indiana: Minton, 2001); March–­May (Indiana: Brodman and Kilmurry, 1998; New York: Wright, 1914; Ohio: Zenisek, 1963); April–­May (Alberta: Randall et al., 2014; Ontario: Piersol, 1913; Québec: Desroches and Rodrigue, 2004; New Hampshire: Oliver and Bailey, 1939; South Dakota: Fischer, 1998; Montana: Werner et al., 2004); late April–­late May (Manitoba: Eddy, 1976; Nova Scotia: Gilhen, 1984); April–­June, depending on elevation (Idaho: Linder and Fichter, 1977); April–­August (Montana: Black, 1970); early–­late May, depending on elevation (Colorado: Corn and Livo, 1989); and mid-­May–­June (New Hampshire: Taylor, 1993). Frogs in breeding condition have even been observed into mid-­ October during warm weather in South Dakota (Bubac et al., 2017), and although gravid females are most often observed in March–­April in Pennsylvania, gravid females also have been found in July and September (Meshaka et al., 2011b). Females deposit eggs from 2 to 7 days ­after arrival, which is about 5–14 days ­after the males start calling. Most reproduction occurs over a period of 10–14 days (Frost and Platz, 1983; Gilbert et al., 1994), although that can be extended by cold weather. In Québec, oviposition begins when ­water temperatures are >8°C (Gilbert et al., 1994). When cold weather interrupts the breeding activity, breeding ceases and resumes once warmer weather appears. Reproduction occurs in shallow ­water (11.8 mg/L), nitrilotriacetic acid (NTA) (>48.8 mg/L), and phenol (>0.074 mg/L). The LC50 for chloroform is 4.16 mg/L, for NTA is 39.3 mg/L, for carbon tetrachloride is 1.76 mg/L, for methylene chloride is >85.0 mg/L, and for phenol is 0.04 mg/L (Birge et al., 1980, 2000). Birge et al. (2000) also provide LC50s for aniline, benzene, dichlorobenzene, nitrobenzene, toluene, and m-­xylene. DDT ­causes a variety of larval deformities, respiratory paralysis, and growth inhibition at 1 ppm (Schreiman and Rugh, 1949). Exposure to sublethal doses of pesticides (DDT, malathion, dieldrin) ­causes an array of effects related to immunosuppression, such as a decreased antibody response. If exposure occurs ­after the frogs have already been exposed to pathogen antigens, immunosuppression does not occur. In wild populations, significant differences can occur in immune function between populations exposed to pesticides and ­those that are pesticide ­free (Gilbertson et al., 2003). The insect growth regulator methoprene did not cause limb malformations in this species in laboratory ­trials (Ankley et al., 1998). Retinoic acid (impor­tant for limb development, but developmentally toxic at certain stages) is in­effec­tive at inducing hind limb malformations at certain stages of larval development of L. pipiens, but is toxic to varying degrees at other stages (Degitz et al., 2000).

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PCBs, polychlorinated dibenzo-­p-­dioxins, and polychlorinated dibenzofurans have been found in the tissues of Northern Leopard Frogs from Wisconsin. Tissue concentrations of PCBs ­were correlated with sediment concentrations. However, the amount of PCBs in the tissues was considered low (Huang et al., 1999). Larvae exposed to the polycyclic aromatic hydrocarbon fluoranthene showed no mortality during a 48 hr experimental uptake period at vari­ous concentrations, and this chemical was rapidly depurated within 48 hrs of exposure to clean ­water. However, exposure to UVA light following fluoranthene exposure significantly increased toxicity. Since this chemical is bioaccumulated in larval tissues, even a residue of 2–10 µg/L could be lethal when exposed to UVA in summertime at northern latitudes (Monson et al., 1999). The chemical perfluorooctanesulfonate (PFOS) is used in a wide variety of products, from protective coatings to adhesives, grease, and insecticides. At 10 mg/L, larvae die within 2 weeks of exposure. At 3 mg/L, time to metamorphosis is delayed and growth is reduced. Tadpoles accumulate the chemical directly from ­water. Ankley et al. (2004) concluded that L. pipiens was not particularly sensitive to PFOS in terms of toxicity or bioaccumulation. In contrast, Foguth et al. (2020) showed that PFOS accumulates in tadpole brains through time and that the extent of concentration was dose dependent Per-­and polyfluoroalkyl substances (PFAS), of which PFOS, PFOA, and PFHxS are examples, are widely used in a variety of ­house­hold applications. PFAS rapidly accumulate in larval Northern Leopard Frogs, reaching a steady state often in 48–96 hrs, but usually by 0.5 m in dia­meter, not too long, and with some light permeability. When Northern Leopard Frogs call in the vicinity of roads with low volumes of traffic noise, their call rates are significantly lower than when they are in the presence of high volumes of traffic noise. Call amplitudes do not change with noise volume, but the dominant frequency of the call is greater when traffic noise levels are high (Cunnington and Fahrig, 2010). Traffic noise also elicits an immediate response by calling frogs lowering the call rate, decreasing the call frequency, and decreasing the call amplitude compared with calls made immediately before exposure to traffic noise. This plasticity in call characteristics allows the frogs to communicate acoustically despite the potential interference of traffic noise. Leopard frogs also have been widely used as biological teaching specimens, as bait, and for food (Chamberlain, 1897; Anonymous, 1907). Harvest for ­these uses has been so heavy at times that biologists have voiced concern for the long-­term survival of the species over extensive regions (Gibbs et al., 1971). Lannoo et al. (1994) suspected at least a 3 fold decline in Northern Leopard Frog populations from the beginning to the end of the twentieth ­century in Dickin­ ecause of overharvest. Early in the son County, Iowa, b ­century, as many as 20 million frogs ­were collected annually

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as food from this one county alone, an extraordinary harvest. Such figures are prob­ably not that uncommon. Evermann and Clark (1916) noted that this species was almost extinct in the Chicago region where hunters took them “in ­great numbers,” and that they ­were sold in fish markets at $0.15 per pound. In 1981, 85,000 Northern Leopard Frogs ­were collected from Lac St. Pierre and Missisquoi, Québec, for use in school laboratories, for food, and as bait (Bider and Matte, 1996). In 1989, >50,000 frogs ­were collected in Minnesota for biological supply ­houses (Oldfield and Moriarty, 1994). Changes in the population size-­class structure may signal long-­term effects from harvest, such as the general decline of large frogs observed (Hoppe and McKinnell, 1997). Northern Leopard Frogs usually breed in semipermanent ponds and dis­appear if fish are stocked into a formerly fishless site (Bovbjerg, 1965). This may account for much of their disappearance in the Midwest, as state fisheries departments stocked predaceous fishes into prairie potholes and farm ponds for sport fishing. In addition, a number of authors have suggested that the introduction of American Bullfrogs into wetlands inhabited by L. pipiens has led to the decline or extirpation of the species (Hammerson, 1982; Panik and Barrett, 1994). Livo (1984) noted declines in a leopard frog population ­after the introduction of bullfrogs. The bullfrogs ate the tadpoles and metamorphic Northern Leopard Frogs and, as a result, ­there ­were fewer egg masses deposited, although adults persisted. Although the bullfrogs did not eat the adults to extinction, they effectively eliminated recruitment. Embryonic and larval L. pipiens have poor hatching success and survivorship in apple orchards affected by a variety of pesticides, although identifying the specific cause or ­causes of effects was not pos­si­ble in the study by Harris et al. (1998b). ­These authors concluded that the influx of chemicals from orchards into breeding sites, in addition to ­factors such as temperature, could act as environmental stressors affecting premetamorphic development. Northern Leopard Frogs readily colonize restored wetlands (Lehtinen and Galatowitsch, 2001; Brodman et al., 2006; Bartelt and Klaver, 2017). For example, they ­were found in greater numbers in small wetlands dredged from 30 to 95% of their area than in undredged reference wetlands on Prince Edward Island (Stevens et al., 2002). Bartelt and Klaver (2017) noted colonization within 3 yrs. Northern Leopard Frogs moved from 31 to 857 m to reach the restored sites, and depended upon roadside ditches to do so. In a similar study, Pouliot and Frenette (2010) found that Northern Leopard Frog tadpoles grew faster in large, artificially constructed waterfowl management basins than in natu­ral wetlands, and that they ­were longer and weighed more as metamorphs. In the Prairie Pothole Region, they may use restored conservation grasslands, but they are more common in

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native habitats (Ba­las et al., 2012). Captive rearing and re­introduction have been attempted in British Columbia (Adama et al., 2004), but results have not been published. In any case, source populations of similar ge­ne­tic history may not be available for repatriation (Wilson et al., 2008a). In Washington State, for example, the last known population actually consists of 3 subpopulations based on analyses of 7 microsatellites (Seaborn and Goldberg, 2020). One of the best methods to ensure survival is habitat protection, such as the establishment of reserves within agriculturally dominated areas of the Richelieu Plain in Québec (Galois and Ouellet, 2005). Another way is to combine ge­ne­tic analyses with population viability analyses to determine optimal approaches when undertaking management and translocation (Seaborn and Goldberg, 2020). Based on such an assessment, ­these latter authors suggested that adults may actually be the optimal life history stage for successful translocations, although large numbers are necessary and ­there is frequently a prob­lem with adults leaving a release site ­because of their high site fidelity. ­Because only metamorphs are usually available, however, success may rely primarily on active management programs, such as releasing frogs within an existing metapopulation structure of multiple ponds. The models used by Seaborn and Goldberg (2020) further suggested the potential impacts of stochastic events on transloca-

tion success and noted that released frogs should come from a variety of subpopulations to maximize ge­ne­tic diversity. Northern Leopard Frogs are frequently surveyed via road transects, as researchers or volunteers make periodic stops to listen for frog calls (Bishop et al., 1997; Bonin et al., 1997a; Lepage et al., 1997; Bowers et al., 1998; Mossman et al., 1998; D. Smith et al., 2014). Northern Leopard Frogs ­were detected during 60% of such nocturnal call surveys in Ontario, making them the least detected species. In Alberta, detection probabilities varied annually and seasonally (better in summer than spring) from 30 to 79% (D. Smith et al., 2014). Some variation in detection probability may be due to methodology, rather than mirror population trends or serve as an indication of rarity. Call surveys for this species likely underestimate occupancy, as frogs do not call continuously, and their calls often are not heard across distances (de Solla et al., 2005). In terms of research techniques, toe clipping does not appear to have adverse effects on survivorship (Ginnan et al., 2014). This species is considered Endangered in Washington, of Special Concern in Connecticut (Klemens et al., 2021) and Colorado, a Sensitive Species in Nevada, and a Species at Risk in Alberta. The species is considered Endangered (Rocky Mountain population) or of Special Concern (Boreal and Prairie populations) by COSEWIC.

Lithobates septentrionalis (Baird, 1854) Mink Frog Grenouille du Nord

becomes more purplish ­toward the rear and lateral sides of the frog. Kramek and Stewart (1980) identified 3 main patterns: light spotted, dark spotted, and dark reticulated. Large, irregular, light or dark brownish-­black, round spots are pre­sent that are broken up by coalescing, greenish-­ yellow lines of varying width in no par­tic­u­lar pattern. Very dark reticulated patterns are only seen in large females (55–72 mm SUL) (Kramek and Stewart, 1980), but in general, frogs become darker with age and size. This occurs as the result of an increase in spot size, but not number. Dorsolateral folds may be pre­sent, but they are often faint or absent altogether. The tympanum is large, especially in males. The iris is hazelnut brown. Bellies are grayish white with dusky spots and reticulations on the tibiae, but pale yellow may be pre­sent on the lower sides and chins of some animals. The legs are short and without white tuberculations on the back and hind leg skin. The rear toes are fully webbed. Most juveniles have a uniform pattern of dark spots and are similar to light-­spotted adults. This species may be very difficult to distinguish from L. clamitans, especially among the smaller size classes. In sympatric populations, L. clamitans is usually the larger frog.

ETYMOLOGY

septentrionalis: Latin for ‘northern,’ referring to the northern distribution of this species. NOMENCLATURE

Conant and Collins (1998): Rana septentrionalis Dubois (2006): Lithobates (Aquarana) septentrionalis Fouquette and Dubois (2014): Rana (Lithobates) septentrionalis Synonyms: Rana sinuata This species has frequently been confused with Lithobates catesbeianus and L. clamitans, and the lit­er­a­ture is replete with identification errors (Hedeen, 1986). IDENTIFICATION

Adults. Adults are brownish to olive dorsally with slightly rough skin. The blotched or boldly mottled dorsal coloration

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Lithobates septentrionalis 569

Males are smaller than females, and frogs from northern populations are larger than ­those from southern populations (Leclair and Laurin, 1996). The maximum size rarely exceeds 70 mm SUL, but Shirose and Brooks (1997) give the maximum size as 76 mm SUL. Schueler (1975) provides mean SULs for adult males and females and large juvenile females from throughout the range (see Systematics and Geographic Variation). In Maine, males ­were 30.2–57.2 mm SUL over a 2 yr period (means 51.3 and 51.7 mm) (Bevier et al., 2006). In Minnesota, males ­were 45–72 mm SUL and females ­were 54–72 mm SUL (Hedeen, 1972a) in 1 study, with a mean of 52–56 mm SUL and 54–63 mm SUL, respectively, in another study (Tenneson, 1983). Ontario females averaged 58–60 mm SUL over a 6 yr period, with the largest individuals at 66–76 mm SUL (Shirose and Brooks, 1997); Logier (1928) reported adults 42–57 mm SUL, also in Ontario. In Nova Scotia, males ­were 49–68 mm SUL (mean 56.3 mm) and females ­were 53.5–65 mm SUL (mean 60.1 mm) (Gilhen, 1984). In New York, males averaged 53.1 mm SUL and females 46.9 mm (Patrick et al., 2012). Larvae. Most accounts describe the tadpole’s dorsum as dark brown to green with small, dark markings or spots, although Vogt (1981) reported tadpoles being emerald green or yellowish green with black spots. Wright (1932) indicated that the bright colors tend to become more subdued as the tadpole approaches metamorphosis. Bellies are opaque straw yellow, and the sides are mottled. Tails have pinkish to buff spotting or blotches and the tail musculature is not bicolored; tails terminate to the posterior of the spiracle and are acute. The iris is black and pinkish cinnamon. Larvae can reach 100 mm TL and weigh nearly 16 g (Hedeen, 1971). Tadpoles ­were described by Altig (1970) and Vogt (1981). Eggs. The eggs are brown to black dorsally and buff to creamy yellow ventrally. Eggs have 2 envelopes surrounding the vitellus (Livezey and Wright, 1947). The outer envelope is 5.6–7.0 mm in dia­meter (mean 6.3 mm) but swells to 8–9 mm; the inner envelope is 2.4–3.0 mm (mean 2.7 mm). The vitellus is 1.3–1.7 mm (mean 1.4 mm) (Wright, 1932; Livezey and Wright, 1947; Moore, 1952). Bishop (in Wright, 1932) mentions the possibility of a third envelope. The eggs are deposited in a jelly mass mea­sur­ing 5 cm × 5 cm × 2.5 cm to 15 cm × 7.5 cm × 5 cm. DISTRIBUTION

Mink Frogs occur from southern Labrador westward around James Bay, southern Ontario, and into southeastern Manitoba (Whiteshell and Mopining Provincial Parks). ­There are several expanding breeding populations on Newfoundland, but ­these represent introductions (Warkentin et al., 2003; Campbell et al., 2004; Kelly et al., 2017). ­These frogs are found around Lake Superior in northeastern Minnesota,

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Distribution of Lithobates septentrionalis

northern Wisconsin, and the Upper Peninsula of Michigan. Isolated populations occur in west central Ontario and near the coast in southern Maine. Populations are found from Nova Scotia and New Brunswick west across the northern New ­England states to upstate New York (Adirondack and Tug Hill plateaus). The species is absent from extreme southern Ontario. A rec­ord from Okak in eastern Labrador (Packard, 1866) is in error (Harper, 1956; Maunder, 1983). Mink Frogs may be found on islands in the Apostle Archipelago of Lake Superior (Hecnar et al., 2002; not observed by Bowen and Beever, 2010), Isle Royale (Ruthven, 1908), An­ti­cos­ti Island at the mouth of the St. Lawrence River (Bleakney, 1954), and on northern Cape Breton Island (Gilhen, 1984). Impor­tant distributional references include: range-­wide (Hedeen, 1986), eastern Canada (Bleakney, 1958a; Logier and Toner, 1961), the ­Great Lakes Region (Harding and Mifsud, 2017), Labrador (Maunder, 1983), Maine (Hunter et al., 1999; Persons and DeMaynadier, 2016), Manitoba (Cook, 1963; Preston, 1982), Michigan (Ruthven, 1912; Harding and Holman, 1992), Minnesota (Oldfield and Moriarty, 1994; Moriarty and Hall, 2014), New Brunswick (Cox, 1899; Rowe, 1899), New Hampshire (Oliver and Bailey, 1939; Taylor, 1993), New York (Gibbs et al., 2007), Ontario (Logier, 1928; Schueler, 1973; MacCulloch, 2002), Québec (Bleakney, 1954; Harper, 1956; MacCulloch and Bider, 1975; Bider and Matte, 1996; Desroches and Rodrigue, 2004), Vermont (Andrews, 2001, 2019), and

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570  RANIDAE

Wisconsin (Suzuki, 1951; Vogt, 1981). Moore (1952) and Hedeen (1986) provided a comprehensive summary of the early distributional lit­er­a­ture on this species and noted many misidentifications. FOSSIL REC­O RD

­There are no fossils reported for this species. SYSTEMATICS AND GEOGRAPHIC VARIATION

The Mink Frog is a member of the Lithobates catesbeianus species group (Aquarana) of North American (Novirana) ranid frogs (Hillis and Wilcox, 2005). The species appears to have diverged from the ancestral lineage about 11 mya (Austin et al., 2003a). The species undoubtedly assumed its pre­sent distribution through a combination of vicariance and dispersal events, but the exact way this occurred is not known. In one scenario, the ancestral lineage may have been pre­sent on the Atlantic Coastal Plain, with subsequent isolation and dispersal by L. septentrionalis northward ­after the last Ice Ages. Alternatively, an ancestral species may have been very widespread, with subsequent isolation and dispersal in the Appalachian (L. catesbeianus-­L. clamitans) and the northern Laurentian (L. septentrionalis) regions (Austin et al., 2003a). In this regard, a previous suggestion that the catesbeianus species group is paraphyletic (Pytel, 1986) is not currently supported. Morphological and phenotypic variation is pre­sent among Mink Frog populations. For example, Cox (1899) noted that individuals from New Brunswick ­were larger than more southern forms, and Mink Frogs in northern portions of Québec and Ontario and eastern Manitoba ­were much larger (female means 60–67 mm SUL; male means 52– 64 mm SUL) than ­those from the southern portions of Ontario and Québec (female means 49–55 mm SUL; male means 44–55 mm SUL) (Schueler, 1975). In northern Québec, males average 61.1 mm SUL and females 67.9 mm SUL, whereas in southern Québec males average 52.6 mm SUL and females 58.6 mm SUL (Leclair and Laurin, 1996); in general, the northern frogs ­were 17% larger than ­those in the south. New Brunswick L. septentrionalis have much blotching on the throat that is not observed in other populations, and Gilhen (1984) noted individuals in Nova Scotia that ­were strikingly green, bronze, and black. ADULT HABITAT

Mink Frogs prefer larger wetlands (ponds and lakes >1.5 ha) with medium to high densities of emergent vegetation (such as Carex sp., Muhlenbergia sp., Typha sp.), a large amount of floating vegetation (such as Nuphar sp.), and substantial amounts of mud and silt as substrates (Marshall and Buell, 1955; Courtois et al., 1995; Popescu and Gibbs, 2009).

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Presence is often associated with beaver ponds. Mink Frogs do not generally occur in areas where the mean temperature in July is >19.5°C (Popescu and Gibbs, 2009), and they are not associated with rocky substrates. This species prefers littoral habitats, although frogs may be found along the shoreline in tamarack (Larix sp.) vegetation (Marshall and Buell, 1955). Some authors have classified the Mink Frog as a boreal species or northern hardwood–­white pine associate, but the type of surrounding forest is not impor­tant. Instead, L. septentrionalis is an inhabitant of what has been termed the humid cold climatic region (Hedeen, 1986). Mink Frogs may be restricted to cold-­water lakes ­because the globular egg mass may not allow sufficient oxygen diffusion for development at warmer temperatures. Although Mink Frog aquatic habitats are surrounded by densely wooded terrestrial habitats (Knutson et al., 2000), this is truly a large wetland species that lives in the forest. In addition to large wetlands, L. septentrionalis occurs in bogs, in quiet streams ­running through bogs, and even in ditches, pools, and puddles (Hedeen, 1971). For example, Mink Frogs have been reported from a small creek (0.5 m deep × 1.5 m wide) ­running through a forest of willows and alders (Schueler, 1973), and from along riverbanks (Cox, 1899). Mink Frogs inhabit a variety of riparian habitat types including herbaceous, shrubby, and wooded areas (Maisonneuve and Rioux, 2001). Dickerson’s (1906) assertion that the Mink Frog is a river frog that is not found in lakes is incorrect and prob­ably was obtained from Garnier’s (1883) observations. Mink Frogs occur to 680 m in New Hampshire (Oliver and Bailey, 1939), 872 m in New York (Moore, 1952), and 896 m (actually 917 m) in Vermont (Hoopes, 1938; Persons et al., 2021). The highest known elevation is from 1,036 m in the Mahoosuc Range, Oxford County, Maine (Persons et al., 2021). Hoopes (1938) states that even in northern Maine along the coast, the species is not found at elevations 7.5. Abundant emergent vegetation should be pre­sent for the frogs to sit upon and to attach their egg masses. Ponds may contain fish, as might be expected of a species that occupies large bodies of ­water (Hecnar and M’Closkey, 1997b). Nothing has been reported concerning the characteristics of bog breeding sites, or ­whether breeding occurs in rivers (unlikely), although

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Egg mass of Lithobates septentrionalis. Photo: David Patrick

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Lithobates septentrionalis 573

1984). Shirose and Brooks (1995a) reported 2 clutches of 610 and 525 in Ontario. Clutch size is positively correlated with female body size (Gilhen, 1984). Hatching occurs in 4–5 days in the laboratory (Moore, 1952; Patrick et al., 2012) to 11–13 days in the field (Wright, 1932) at 8–10 mm TL. LARVAL ECOLOGY

Larvae are herbivorous, primarily consuming algae. Like many species, the rasping mouthparts are used to scrape algae from plant stems and submerged debris. Feeding occurs for at least 1 year. Most larvae overwinter and metamorphose the following summer, although some may spend 2 winters prior to metamorphosis (Hedeen, 1971). During the winter, growth essentially ceases, and mortality from freezing may be high ­unless ponds and lakes are deep enough to prevent them from freezing solid. Larvae are found in well-­oxygenated ­waters, and Garnier (1883) observed them in fast-­flowing riffles, where they ­were extremely wary. ­Under laboratory conditions, embryos ­will not develop at temperatures >30.4°C (Moore, 1952). No development occurs at 7.8°C, and gill circulation fails to develop at 12.9°C. Moore (1952) suggested that the range of temperature tolerance was 14–30°C, and that L. septentrionalis did not possess any specific cold weather adaptations regarding larval development. In nature, however, most ­waters where this species deposits its eggs remain 80%) for slow-­moving prey, but low (3 yrs. In Québec, longevity also reached 4 yrs based on skeletochronology (Sagor et al., 1998), and some females can reach 5 yrs (Bastien and Leclair, 1992). However, in Wood Frogs from the Appalachian Mountains, maturity normally occurs 3 yrs ­after metamorphosis with a few maturing at 2 and 4 yrs (Berven, 1982a). Most mountain-­dwelling females (62%) matured at 4 yrs. Latitude also affects the timing of sexual maturity. In Québec, southern populations of L. sylvaticus matured ­later and ­were larger than southern conspecifics from low elevations studied by Berven (1982a), but they matured ­earlier and ­were smaller than Berven’s (1982a) southern frogs from higher elevations (Bastien and Leclair, 1992; Sagor et al., 1998).

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The results of several studies (Newman and Squire, 2001; Stevens et al., 2006a) suggest that Wood Frogs are not as philopatric as some authors (e.g., Berven and Grudzien, 1990) indicate. Still, Vasconcelos and Calhoun (2004) found that 98% of males and 88% of females returned to their pond of original capture. Most populations at nearby ponds have very similar allele frequencies, suggesting significant amounts of gene flow among closely spaced breeding ponds (Newman and Squire, 2001; Squire and Newman, 2002). In an example of breeding site flexibility, the majority of the population tends to move to adjoining ponds rather than breed in remnant pools when beaver ponds are altered by dam destruction (Petranka et al., 2004). When beavers altered the drainage patterns of formerly isolated ponds, allowing fish to invade, Wood Frog use declined in relation to the extent of fish invasion (more fish, less breeding), ­until they ­stopped breeding altogether and shifted to alternate ponds. If fish ­were pre­sent, they bred only in very shallow extensions of the pond, away from fish. When beaver activity was curtailed and the ponds restored to a fish-­free presence, Wood Frogs again used the site for breeding (Petranka et al., 2004). Overall, the general population remained stable despite the vari­ous habitat shifts over a 10 yr period. Site shifting also happens when excessive siltation occurs at formerly favorable breeding sites. If fish are pre­sent and fishless ponds are available, Wood Frogs ­will go to the fishless ponds, even if they formerly bred in the pond now containing fish (Hopey and Petranka, 1994), especially if the ponds ­were 15°C. Males are usually in close proximity to one another when calling, suggesting that pairing occurs haphazardly as frogs approach one another; reproductive males likely attempt amplexus with any frog moving in their proximity.

AQUATIC AND TERRESTRIAL ECOLOGY

Adult frogs prob­ably do not venture far from ­water, but Hale et al. (1977) noted that individuals ­were observed in “unlikely” canyons during the summer monsoon season. Some individuals seem to prefer to remain at a single location. Adult males have been reported to move up to 1,885 m, and adult females to 651 m (in Rorabaugh and Hale, 2005). Movements occur from the end of the dry season in June through the rainy season in August. Juveniles

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BREEDING SITES

Breeding occurs in permanent shallow pools within intermittent streambeds in the narrow rocky canyons inhabited by this species. Rorabaugh and Humphrey (2002) noted that the best breeding sites are plunge pools that have low mean flows (10 months, however. Summer transformation would allow recent metamorphs time to grow. In addition, it might allow juveniles to disperse along stream beds during brief periods of rain during the summer monsoons. Most recent metamorphs are 35–40 mm SUL, but froglets as small as 21 mm SUL have been reported (Rorabaugh and Hale, 2005). DIET

Lithobates tarahumarae is not a selective predator and ­will feed on just about any animal it can catch. Prey include invertebrates (beetles, sphinx moths, ­water bugs, scorpions, mantids, grasshoppers, wasps, spiders, caddisflies, katydids, moths, centipedes), fish (Gila ditaenia), juvenile mud turtles (Kinosternon sonoriense), and even snakes (Tantilla atriceps) (Zweifel, 1955; in Rorabaugh and Hale, 2005). Zweifel (1955) suggested that the prey ­were indicative of nocturnal feeding habits. PREDATION AND DEFENSE

This species dives into the ­water upon the approach of a potential predator and may utter a startle “squawk” (Campbell, 1934). They hide in leaf litter, ­under rocks, or in woody debris. According to Rorabaugh and Hale (2005), the skin secretions of L. tarahumarae are noxious to taste and may cause minor skin irritation. Specific predators have not been recorded, but it seems likely that a variety of mammals (Bassariscus astutus), birds, and snakes (including the Garter Snake Thamnophis cyrtopsis) prey upon juveniles and adults. Larvae may be eaten by aquatic invertebrates (Belostoma, Lethocerus), mud turtles, and garter snakes (in Rorabaugh and Hale, 2005).

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macrocirra; the cestode Ophiotaenia magna; the nematodes Falcaustra inglisi, F. lowei, Foleyellides striatus, Oswaldocruzia pipiens, Physaloptera sp., Rhabdias ranae, Subulascaris falcaustriformis; and unidentified acanthocephalans (Bursey and Goldberg, 2001). Hale and Jarchow (in Rorabaugh and Hale, 2005) also reported unspecified nematodes and cestodes in this species. Unidentified mites have been reported on this species (Lemos Espinal et al., 2013). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Adult Lithobates tarahumarae. Photo: Carl Gerhardt

Metals. Elevated levels of cadmium and arsenic ­were found in ­water where populations of L. tarahumarae dis­appeared in southern Arizona. Hale et al. (1995) speculated that acid-­soluble zinc leached from canyon walls, in conjunction with accumulations of insoluble cadmium in sediments, combined to have detrimental effects upon this species. This hypothesis has yet to be empirically examined.

POPULATION BIOLOGY

Virtually nothing is known about the population biology of this species. Campbell (1934) recorded it as “of rather common occurrence” at the permanent waterholes within the region. An estimate of the population size at its peak in the Santa Rita Mountains was 1,020 frogs in 1976, but this fell to 625 by 1977. By 1982, only 1–3 frogs ­were observed, with the last frog being sighted in 1983 (Hale et al., 1995). In unpublished data, Hale and Mays (in Rorabaugh and Hale, 2005) reported that maturity is reached at 2 yrs following metamorphosis, when males are ca. 64 mm SUL and females are 67 mm SUL. Longevity may extend to 6 yrs post-­metamorphosis. COMMUNITY ECOLOGY

Lithobates tarahumarae shares its habitat with Chiricahua Leopard Frogs (L. chiricahuensis) and Lowland Leopard Frogs (L. yavapaiensis). The nature of ­these species interactions is unknown. American Bullfrogs have colonized some streams within the range of L. tarahumarae, and ­these must be eradicated prior to the reestablishment of Tarahumara Frog populations DISEASES, PARASITES, AND MALFORMATIONS

Amphibian chytrid fungus (Batrachochytrium dendrobatidis, Bd) has been found in L. tarahumarae populations throughout the species’ range and is likely to have played a major role in the disappearance of this frog in Arizona and parts of México (Hale et al., 2005). Some infected populations have not declined, however. Antimicrobial peptides are pre­sent in the skin of this species and may provide some protection against the fungal pathogen (Rollins-­Smith et al., 2002). Tarahumara Frogs are parasitized by the trematodes Glypthelmins quieta, Haematoloechus breviplexus, Langeronia

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Habitat of Lithobates tarahumarae. Santa Rita Mountains, Arizona. Photo: James Rorabaugh

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Lithobates virgatipes 635

STATUS AND CONSERVATION

This species was extirpated by 1983 from southern Arizona due to undocumented ­causes, and its precarious status in the United States was recognized as early as 1974, when dead and ­dying frogs ­were discovered in the Atascosa-­Pajarito Mountains (Bury et al., 1980; Hale et al., 2005). Populations in the Santa Rita Mountains began declining in 1977. Many populations in México also are declining or have been extirpated (Hale et al., 2005), but the species is not protected ­there. Rorabaugh et al. (2020) summarize the history of population declines in the United States and México, as well as provide detailed accounts of re­introduction efforts. In order to have larvae and frogs to repatriate into formerly occupied habitat, 4 adults and 30 small tadpoles ­were imported into the US in 1999 from Sonora to begin studies on captive breeding, but ­these individuals died shortly thereafter. A partial egg mass was imported in 2000, tadpoles from which ­were released at several Arizona sites (Rorabaugh and Humphrey, 2002; Rorabaugh et al., 2020). An experimental population ­later was established in 2002 at the Kofa National Wildlife Refuge in western Arizona. This population continues to exist as of 2019 (Rorabaugh et al., 2020). Tarahumara Frogs (adults, juveniles, and larvae) ­were repatriated into Big Casa Blanca Canyon in Santa Cruz County beginning in 2004. Frogs to be repatriated ­were treated with itraconazole to counteract the effects of amphibian chytrid. Egg masses, large tadpoles, juveniles, and adults ­were observed subsequently, indicating reproduction and

Lithobates virgatipes (Cope, 1891) Carpenter Frog ETYMOLOGY

virgatipes: from the Latin words virgatus, meaning ‘striped,’ and pes, meaning ‘foot.’ The name refers to the striped markings on the rear feet. NOMENCLATURE

Conant and Collins (1998): Rana virgatipes Dubois (2006): Lithobates (Aquarana) virgatipes Fouquette and Dubois (2014): Rana (Lithobates) virgatipes Synonyms: None IDENTIFICATION

Adults. Adults are olive, greenish, tan, or brown with blackish, mottled spots dorsally. ­There are 2 yellowish,

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survival. However, numbers w ­ ere reduced by 2007 ­after post-­fire flooding and extensive sedimentation of the breeding pools, as well as continued die-­offs from Bd. Additional frogs ­were released from 2012 to 2015, and frogs have persisted in this and an adjacent drainage through 2019 (Rorabaugh et al., 2020). Another experimental population was established in 2014 at Sycamore Canyon in the Atascosa Mountains ­after eradication of American Bullfrogs, and small numbers continue to persist as of 2019 (Rorabaugh et al., 2020). The repatriated population is protected by Arizona state law. Suggested ­causes of declines have included habitat loss, the influence of toxic chemicals resulting from copper smelters, acid rain, competition, extreme weather, and disease, especially Bd. This pathogen may be the primary cause of decline, especially since some of the symptoms attributed to metal poisoning (dry skin, lack of righting response, partial paralysis) are also symptomatic of the final stages of chytridiomycosis. However, some populations in México have been infected for at least 17 yrs without declining (Hale et al., 2005), and Hale et al. (1995) noted that no abnormalities ­were pre­sent in histopathological examinations carried out by Elliott Jacobson at the University of Florida. Cold and pollution may increase the severity of the disease or at least make frogs more susceptible to it. The interaction of sublethal effects on the virulence of amphibian chytrid is not well understood. Much data on the life history and ecol­ogy of this species can only be found in unpublished reports. Rorabaugh and Hale (2005) have summarized some of this information.

reddish-­brown, or light brown dorsolateral lines ­running parallel along the dorsum; ­these lines begin ­behind the eye. A second series of lines runs from the edge of the upper jaw, below the tympanum, to the groin; the lower lines may be more distinctive and broader than the upper lines and contain light-­colored tubercles. Both sets of lines tend to fade out posteriorly. The tympanum and iris are bronze. Bellies are whitish with dark spots, although some individuals have a dull yellowish coloration on the lower surface of the head and forefeet. Blackish-­brown spots or blotches are found along the lateral sides of the body; ­these are more prominent than the dorsal markings. The rear of the femur has alternating light and dark bars, with the blackish-­brown variegated markings of the undersides extending onto the belly. The longest toes of the rear feet extend well beyond the webbing, unlike that of small L. grylio, which are similar in appearance to Carpenter Frogs and with which they may be confused. Considerable variation exists among frogs with regard to coloration and extent of dorsal and ventral blotching, and ­there are no differences between the sexes in

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color patterns. Fowler (1905) illustrates several examples of ventral pattern variation. Males have a swollen thumb, vocal sacs, and a tympanum larger than that of females. Males are slightly smaller than females. In New Jersey, adult males ranged from 39 to 62 mm SUL (Given, 1988a). Wright and Wright (1949) gave a range of 41–63 mm SUL for males and 41–66 mm SUL for females. In ­Virginia, males averaged 53 mm SUL (range 45–63 mm), and females averaged 53 mm SUL (range 42–66 mm) (Georgel, 2001). The largest reported male is 68 mm SUL, and the largest female 73 mm SUL (Standaert, 1967). Larvae. The dorsal larval coloration is grayish or brown with black spots. ­There is a distinctive stripe or row of dots formed along the lateral line pores on the dorsal tail fin. The tail musculature is not bicolored, a tail musculature stripe is pre­sent, and ­there are light spots surrounded by dark pigment near the edge of the tail fins. The intestine usually is not vis­i­ble, but it may be indistinct. Lower jaws are relatively narrow. The maximum larval size is 100 mm TL ( Jensen et al., 2008). The tadpole was described by Altig (1970). Eggs. The eggs of L. virgatipes are black above and creamy white to light tan below; they have only 1 envelope surrounding the vitellus. The envelope is a mean of 5.4 mm in dia­meter (range 3.8–6.9 mm), and the vitellus is a mean of 1.6 mm (range 1.4–1.8 mm) in dia­meter (Wright, 1932; Livezey and Wright, 1947). Eggs are deposited in firm, globular clumps and hatch in about 3–7 days. DISTRIBUTION

Carpenter Frogs occur in scattered populations from the southern half of New Jersey to northern Florida (Osceola National Forest). They are found solely on the Atlantic Coastal Plain, reaching as far inland as the Fall Line in South Carolina and adjacent North Carolina. Impor­tant distributional references include: range-­wide (Reed, 1957), Delaware (Conant, 1940; White, 1988; White and White, 2007), Florida (Stevenson, 1969; Krysko et al., 2019), Georgia (Wright, 1932; Neill, 1952; Williamson and Moulis, 1994; Jensen et al., 2008), Mary­land (Conant, 1947; Grogan, 1974; Harris, 1975; Given, 1999a; White and White, 2007; Cunningham and Nazdrowicz, 2018), Mid-­Atlantic (Beane et al., 2010), New Jersey (McCormick, 1970; Schwartz and Golden, 2002), North Carolina (Brimley, 1944; Meyers and Pike, 2006; Dorcas et al., 2007; Gaul and Mitchell, 2007), South Carolina (Neill, 1952), and ­Virginia (Funderburg et al., 1974b; Grogan, 1974; Tobey, 1985; Mitchell and Reay, 1999). FOSSIL REC­O RD

­There are no known fossils of this species.

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Distribution of Lithobates virgatipes SYSTEMATICS AND GEOGRAPHIC VARIATION

The Carpenter Frog is a member of the Lithobates catesbeia­ nus species group (Aquarana) of North American (Novirana) ranid frogs (Hillis and Wilcox, 2005). The species appears to be the basal member of the Aquarana clade and to have evolved from a monophyletic Aquarana ancestor on the Atlantic Coastal Plain. All other members of the Aquarana clade diverged from the ancestral L. catesbeianus group during the late Miocene to early Pliocene (Austin et al., 2003a). Changes in sea level during the Tertiary appear to have played a major role in the evolution of the Aquarana clade and likely contributed to the species’ current distribution. In this regard, a previous suggestion that the Catesbeianus species group is paraphyletic (Pytel, 1986) is not supported. Laboratory hybridization between ♂ L. virgatipes and ♀ L. pipiens does not proceed beyond the gastrula stage (Moore, 1949a). ADULT HABITAT

The Carpenter Frog is a species of highly acidic (pH 4.0–5.0) wetlands on the Atlantic Coastal Plain, although Gosner and Black (1957a) noted that low pH was not required by this species for embryonic development. It is found in a variety of aquatic habitats, including larger rivers, lakes, blackwater creek swamps, small, isolated wetlands, vernal pools, cranberry bogs, beaver ponds, seasonally flooded “Delmarva Bays,” Carolina Bays, pocosins (shrub bogs), and even man-­made impoundments, borrow pits, drainage ditches,

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and canals, as long as ­there is an abundance of floating or thick, emergent vegetation such as lily pads and green arrow arum (Peltandra virginica). ­These frogs are particularly associated with mats of sphagnum and are often called “sphagnum frogs” (Fowler, 1905). They avoid bogs in active cranberry farms, but inhabit bogs, ditches, and reservoirs in abandoned farms (Wen, 2015). In areas of prescribed fire, they only occur in wetlands burned >8 yrs previously, and then only if the habitat has been mulched (Klaus and Noss, 2016). In Mary­land, occupancy is associated with intermediate hydroperiods, acidic wetlands (pH 4.0–5.0), and a surrounding forest cover 50–500 m away from the wetland (Otto et al., 2007b). The amount of forest cover within 50 m of the high-­water mark is particularly impor­tant. In the New Jersey Pinelands, Carpenter Frogs are found in areas of elevated acidity, conductivity (ca. 50–80 μS cm-1), and nitrate-­nitrite levels, where they prefer unaltered habitats where the American Bullfrog is absent (Bunnell and Zampella, 1999; Zampella and Bunnell, 2000). In New Jersey, L. virgatipes is a species of the Pine Barrens, where the dominant pine is pitch pine (Pinus rigida) (McCormick, 1970). They are frequently associated with Atlantic white cedar (Chamaecyparis thyoides). Dominant trees associated with Atlantic white cedar include bald cypress (Taxodium distichum), tupelo gum (Nyssa aquatica), and red maple (Acer rubrum). In the South Carolina Coastal Plain, the species may be found in isolated, small wetlands interspersed within longleaf pine (Pinus palustris) forests located on sandhill ridges (Russell et al., 2002a). In all habitats, soils are organic, and standing ­water usually occurs year-­round except during serious droughts. Carpenter Frogs may inhabit rather small wetlands. For example, Russell et al. (2002a) recorded 307 dif­fer­ent Carpenter Frogs over a 2 yr period in 5 small, isolated wetlands in South Carolina; even a 0.38 ha pond contained 30 L. virgatipes. AQUATIC AND TERRESTRIAL ECOLOGY

Carpenter Frogs sit along the shorelines of slow-­moving streams and ponds, resting on sphagnum mats partially submerged in the ­water. ­There may be some differences in habitat use between the sexes, with males occupying the centers of wetlands and females more likely to be found nearer the shorelines (Standaert, 1967). Males, however, ­will call from shallow ­water near the shoreline, moving to open ­water following breeding. Activity occurs throughout the warm season into the autumn. In ­Virginia, for example, Carpenter Frogs may be found from March to November (Georgel, 2001). Carpenter Frogs can tolerate high temperatures, with the mean CTmax of adults from 35.8 to 38°C, depending upon

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acclimation temperature (Holzman and Mc­Manus, 1973). Overwintering occurs in aquatic habitats. Environmentally based mortality can occur both in summer and winter. For example, Standaert (1967) reported finding dead frogs in shallow pools in spring that prob­ably died during the winter due to anaerobic conditions. He further noted frogs that may have died during high summer ­water temperatures, which also produced anaerobic conditions. Males seemed to be particularly vulnerable to anaerobic conditions during what he termed the “summer torpor.” Movement patterns of this species are not well understood. Standaert (1967) anecdotally mentioned that small frogs dispersed from a large pond to a nearby smaller pond in fall, remained ­there during the following spring, then returned to the larger core pond as smaller adjacent ponds dried during the summer. Thus, ­there may be routine periodic movements among adjacent ponds, depending on hydroperiod or other ­factors. Carpenter Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities ( Jaeger and Hailman, 1971, 1973). Given (1988a) confirmed night feeding. They likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Calling begins in the spring and may extend throughout the summer. In ­Virginia, for example, calls can be heard from March to September (Georgel, 2001). The advertisement call of this species is a series (1–10) of loud “ca-­thunk ca-­thunks” that sound like carpenters hammering nails (dominant frequency of 460–720 Hz; lesser defined peak at 1,500–2,500 Hz; 110 dB) (Given, 1987, 1990). Call intensity increases with male body mass. The calling male may be difficult to locate, despite the loud, distinctive call. Most males call ­every night throughout their breeding season, and Given (1988b) recorded a median residency of 40.5 nights (range 8–94). Calling normally occurs from dusk to dawn throughout the main portion of the breeding season, with peak levels at 23:00–01:00 hrs in New Jersey (Given, 1987) and 21:00 hrs in ­Virginia (Georgel, 2001). In August and September in ­Virginia, calling becomes more diurnal and tapers off at night. Not all males call with equal intensity. First year males tend to put less effort into calling than larger, older males. In Given’s (1988a) study, 9 frogs called with low intensity, 26 with average intensity, and 7 with high intensity. Males call from shallow-­water habitats located 0.1–0.5 m from the shoreline while sitting among emergent vegetation. Vegetation must extend at least 1.7 m from the ­water’s edge, but ­water depth per se is not impor­tant to territorial establishment. Males call in a high posture, with their heads up at a slight ­angle and their lungs inflated. They tend to call nightly

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638  RANIDAE

from the same locations, and they ­will defend their calling stations in aggressive encounters with other males (Given, 1988b). Such territorial defense lasts about 2 weeks ­until a male has completed his breeding season. Males establish from 0 to 4 territories (median 1.5) and spend about 84% of their time in such territories (Given, 1988b). Territories are ca. 1 m2 in area, and the mean distance a male moves from 1 territory to establish another is 22.5 m (range 4–200 m). Males frequently return to a previously established territory. When males detect another frog (­whether male or female), they switch from the advertisement call to a single-­note aggressive call (dominant frequency of 565–720 Hz; secondary frequencies of 1,140–1,380 Hz; 17–32 ms call duration). Intruding males respond by uttering a single-­note aggressive call. The aggressive call is higher in primary frequency than the advertisement call, but the secondary frequency of the advertisement call is higher than the secondary frequency of the aggressive call (Given, 1999b). In par­tic­u­lar, males have a higher rate of aggressive calls in response to frequencies of 1,500 Hz than to other frequencies. Essentially what the male does is to modify the secondary frequency of his advertisement call ­toward that of the aggressive call, while maintaining aspects of the advertisement call that are impor­tant to female choice (Given, 1999b). If an intruder is a male, ­there may be a series of aggressive calls between them, followed by a brief wrestling match that residents usually win. Males have a release call that is uttered by 1 male ­toward another if amplexus is attempted. Females also have a low-­intensity, single-­note call during courtship that has higher dominant and secondary dominant frequencies than the male aggressive call (dominant frequency of 720–780 Hz; secondary frequencies of 1,480–1,520 Hz; 16–23 ms call duration) (Given, 1993). If the intruder is a female, the male ­will amplex her, and she ­will begin searching for an oviposition site. Females initiate courtship by moving ­toward a calling male and uttering an encounter call. Males respond with a single or multinote aggressive call and move ­toward her, followed by amplexus. The multinote call is only given in response to a combination of the female’s call and vibrations in the ­water. The mixture of sound and vibrations thus helps the male to home in on the position of the female. In playback experiments, females do not respond to variation in call frequencies or repetition rates, but they may use differences in the harmonic structure to differentiate between large and small males (Given, 1993). Lithobates virgatipes thus has an unusual call repertoire in which the same aggressive call is used ­toward both sexes. The aggressive response of the territorial male is consistent, and ­whether fighting or mating ensues depends on the response of the frog that approaches him. Male mating success is correlated with size (greater for larger frogs) and the number of nights a male calls. Losers of

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aggressive bouts and smaller males may take up positions as satellite males near a large resident. ­These satellites remain with a resident from 1 to 7 days. They may utter a single-­ note call si­mul­ta­neously with the resident, but they assume a more crouched posture (Given, 1988b). BREEDING SITES

The species breeds in ponds that have intermediate to long hydroperiods (Eason and Fauth, 2001; Otto et al., 2007b). Most studies suggest optimal breeding sites are permanent wetlands with a long hydroperiod (Given 1988b; Zampella and Bunnell, 2000; Georgel, 2001), as the larvae may overwinter prior to metamorphosis in some populations. Otto et al. (2007b), however, reported Carpenter Frogs in seasonal wetlands. Some breeding ponds can be quite small (i.e., < 1 ha) (Russell et al., 2002a). REPRODUCTION

Calling occurs from spring throughout the summer, at least in the northern portions of the range. Specific times include mid-­March to early August (North Carolina: Gaul and Mitchell, 2007), late April–­early August (New Jersey: Fowler, 1905; Conant, 1947; Given, 1987, 1988a, 1990), and a peak from April to August, with calls heard from March to September (­Virginia: Georgel, 2001). Wright (1932) heard calls from June to August in the Okefenokee Swamp, but he acknowledged that the breeding season prob­ably began at least in March. I have heard them chorusing in north Florida on 20 April, and ­there are specimens in the Florida Museum of Natu­ral History that ­were collected from March to December. ­After the first several months of the breeding season, calling becomes much more sporadic. It is not clear ­whether breeding takes place throughout the calling period, or ­whether calling takes place over a more extended time than ­actual oviposition. Eggs masses are oviposited near the surface in shallow ­water (0.05–1.0 m in depth), attached to vegetation or debris (Wright, 1932; Given, 1990). The sometimes elongate globular mass is ca. 6.5 cm × 6.5 cm to 5 cm × 10 cm and often has a bluish tinge (Livezey and Wright, 1947). ­There appear to be few ­actual counts of the number of eggs per mass, with most authorities repeating Wright and Wright’s (1949) estimate of 200–600 eggs per mass. Wright (1932) gave counts of 349 and 474 from 2 masses. LARVAL ECOLOGY

­Little information is available on the larval ecol­ogy of L. virgatipes. In New Jersey, tadpoles first appear in late May within the shore zones of ponds. Density then increases steadily through peak metamorphosis in September, when they can reach nearly 120 per ha (Standaert, 1967). Most tadpoles metamorphose in the autumn (September to October), despite

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Lithobates virgatipes 639

against the bacteria Escherichia coli and Staphylococcus aureus (Conlon et al., 2005b). POPULATION BIOLOGY

Tadpole of Lithobates virgatipes. Photo: John Bunnell

suggestions in the lit­er­a­ture (Wright, 1932; Wright and Wright, 1949) that all tadpoles overwinter and metamorphose the following spring or summer. It is likely, however, that some tadpoles do overwinter, especially ­those resulting from egg deposition late in the season. Wright and Wright (1949) reported that young frogs ­were 23–31 mm SUL following metamorphosis, but Standaert (1967) noted that the mean metamorphic size in New Jersey was 32 mm SUL (range 28–36 mm). ­These postmetamorphs disperse rapidly with autumn rains and quickly enter overwintering sites. DIET

Feeding occurs throughout the breeding season, although it appears to be depressed during peak breeding, as calling males concentrate on attracting mates and defending territories. Feeding increases rapidly following the breeding season. In ­Virginia, insects constituted 85% of the items consumed and 94% of the prey volume (Georgel, 2001). Prey included beetles, collembolans, flies, ants, dragonflies, leafhoppers, true bugs, homopterans, a larvae L. virgatipes, and a larval Lesser Siren (Siren intermedia) (Georgel, 2001). Males tended to consume more beetles of the species Donacia rufescens than females, perhaps consistent with differences in habitat use between the sexes. This beetle frequents lily pads and emergent vegetation in open ­water habitats. Females also ate prey of a wider variety of taxa (35 vs. 26) than males. A small amount of vegetation may be ingested with the invertebrate prey. Prey volume is correlated with the frog’s head width, meaning that large frogs eat more and larger prey than small frogs.

Tadpoles that metamorphose in summer may reach the minimum size of sexual maturity prior to their first overwintering period and breed the following spring. For ­those that overwinter ­because of late-­season hatching (August) and metamorphose in spring (April–­May), sexual maturity is not reached ­until mid-­to late summer of the first warm season activity period (i.e., about 10–12 months ­after egg deposition). The size at maturity for males is 40–45 mm SUL in ­Virginia (Georgel, 2001) and 42–44 mm SUL in New Jersey (Standaert, 1967). Standaert (1967) noted, however, that ­there was considerable variation in size at sexual maturity. For example, 95% of males ­were mature by 48 mm SUL, whereas 5% ­were mature by 40 mm SUL. He considered frogs 38–47 mm SUL as juveniles. Populations may experience considerable turnover from 1 year to the next. For example, only 3 of the 26 males followed by Given (1988a) ­were seen the year ­after first marking. Growth rates may be extremely rapid for newly metamorphosed frogs, with rates of 7 mm/month recorded (Standaert, 1967). They usually enter their first overwintering period at 34–36 mm SUL, but a few frogs may reach 49 mm SUL by October of their first year. ­After emergence, growth rates continue at ca. 5 mm/month, allowing sexual maturity by mid-­June. Male growth rates are lower than ­those of females, and males reach a smaller asymptotic size than females. Longevity exceeds 3 yrs. Calling is energetically expensive for males throughout the extended breeding period. As the season progresses, dry body mass and ­percent lipid both decrease, although both show a sharp increase in August ­after the end of the breeding season, presumably as males begin concentrated feeding (Given,

PREDATION AND DEFENSE

­ hese frogs are wary and cryptic as they sit among sphagT num, and quickly sink out of site at the approach of a potential predator. They ­will swim short distances and seek shelter in submerged vegetation. Predators include ­water snakes (Nerodia sipedon) and Black Racers (Coluber constrictor) (Wright, 1932; Jensen et al., 2008), and undoubtedly many types of birds and mammals. Larvae are likely attacked by a variety of predaceous insects and vertebrates, such as birds, turtles, and snakes. The skin contains antimicrobial peptides that may aid in defense

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Adult Lithobates virgatipes. Photo: David Dennis

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640  RANIDAE

1988a). For example, Given (1988a) mentions a male with a growth rate of 0.13 g/day prior to the breeding season, 0.03 g/day during the breeding season, and 0.10 g/day ­after the breeding season. Smaller males (i.e., first year adults) have a lower calling effort than larger males and tend to increase their body mass more than large males. This indicates a trade-­off between reproductive effort (calling) and growth among the smaller males. The sex ratio at breeding ponds is male biased. ­After hibernation, Standaert (1967) found that ­there ­were as many as 2.5 males per female in April, but by June the ratio had dropped to 1.65:1.00. During the latter part of the activity period, the sex ratio became female biased. In addition, sex ratios changed by position in the pond and by time of day. ­These results suggest differences in be­hav­ior, habitat choice, and activity between males and females that bias determinations of the sex ratio. COMMUNITY ECOLOGY

Many authors have noted the association of Carpenter Frogs with acidic wetlands. As noted by Gosner and Black (1957a), normal embryonic and larval development can take place at higher pHs than usually encountered in the boggy or swampy areas frequented by Carpenter Frogs, although a low pH is lethal to many other anuran eggs and larvae. It may be that by occupying acidic habitats, Carpenter Frogs are able to reduce interspecific competition with other anurans, particularly American Bullfrogs and Pickerel Frogs. Other ranids (L. clam­ itans, L. sphenocephalus), however, are somewhat acid tolerant and are often found in the same habitats as L. virgatipes, whereas the hylids Acris crepitans and Dryophytes versicolor are not (Zampella and Bunnell, 2000). Thus, the life stage (larval, adult, or both) at which competition might occur among species, and ­whether and how acidity might reduce competition, is not well understood. When calling, L. virgatipes and L. clamitans form mixed-­ species aggregations along a shoreline. ­There is a slight degree of habitat partitioning when calling, with L. clami­ tans calling from vegetation at the shoreline and L. virgatipes calling from locations 0.1–0.5 m into the wetland (Given, 1990). Male L. clamitans are capable of displacing male L. virgatipes from calling sites. When choruses are mixed, L. virgatipes tend to associate with conspecifics rather than the larger and heavier L. clamitans; L. clamitans shows no such preference. DISEASES, PARASITES, AND MALFORMATIONS

A single individual tested for amphibian chytrid fungus at the Savannah River Site in South Carolina was negative (Daszak et al., 2005). Amphibian chytrid fungus was not found on this species in Florida, based on museum specimens

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Habitat of Lithobates virgatipes. New Jersey Pinelands. Photo: John Bunnell

(Karwacki et al., 2021). Perkinsea and Ranavirus ­were found on specimens from the state in the 1970s, but their prevalence was rare. Ranavirus has also been reported from this species in North Carolina (Lentz et al., 2021). SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. The lethal and critical pH levels for this species are 3.5 and 3.6–3.8, respectively (Gosner and Black, 1957a). Bunnell and Zampella (1999) found Carpenter Frogs in wetlands with a pH of 3.8–4.1 in the Pine Barrens of New Jersey. Carpenter Frogs can tolerate ­water to a pH of 9.2 with no ill effects. STATUS AND CONSERVATION

This species occurs over a discontinuous range on the Atlantic Coastal Plain. It may be locally abundant, although its secretive nature means it could be easily overlooked. Some populations undoubtedly have been lost due to habitat destruction or alteration. For example, Brady (1927) recorded this species as widespread in ­Virginia’s Dismal Swamp, but subsequent attempts to locate it ­there ­after years of intensive forest cutting have proven unsuccessful (Pague and Mitchell, 1987). However, Russell et al. (2002b) found no effects on resident L. virgatipes inhabiting isolated wetlands (to 1.06 ha) up to 1.5 yrs following clearcutting in South Carolina. The wetlands ­were left intact, and wetland vegetation was undisturbed by forestry treatments, thus allowing this highly aquatic species to persist. In New Jersey, Weir et al. (2009) reported a slight positive trend in occupancy in wetlands monitored from 2001 to 2007. In Mary­land, Given (1999a) found the species at only 8 of 22 historically reported populations in that state (a total of 9

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Lithobates yavapaiensis 641

of 34 sites surveyed). Otto et al. (2007b) ­later recorded them at 18 of 40 study sites on the western and southern portions of the Delmarva Peninsula of Mary­land. In ­Virginia, ­there are only 6 populations known from the state; the largest population is at Ft. A.P. Hill. This species is considered Endangered in Mary­land.

Every­thing that is known concerning the life history of L. virgatipes is based on studies in New Jersey and ­Virginia. Nothing is known about its habits, breeding phenology, activity patterns, or reproductive biology in the more southern parts of its range. Life history studies in the South are long overdue.

Lithobates yavapaiensis (Platz and Frost, 1984) Lowland Leopard Frog

some individuals; this species lacks the light halo seen in some other Southwestern leopard frogs. Spots are absent on the head in front of the eyes. An incomplete light stripe is pre­sent on the upper lip but does not extend in front of the eyes. The groin region contains yellow pigment that may extend onto the venter and the underside of the legs. ­There is a dark reticulation pattern on the posterior portion of the thighs, and the dorsal portions of the thighs are barred. Venters are cream colored, and chins are light. Rear toes are fully webbed. Males have prominent external vocal sacs and enlarged thumb pads. This species is longer limbed than the similar L. onca. Males are smaller than females. In the type series, males averaged 54.7 mm SUL (range 50–57 mm), and females averaged 63.5 mm SUL (range 59–72 mm). Other morphological mea­sure­ments are provided by Platz and Frost (1984). Degenhardt et al. (1996) give the size range as 46–72 mm SUL for males and 53–87 mm SUL for females, whereas Goldberg (2019h) rec­ords males at 43–68 mm SUL (mean 56.7 mm) and females at 53–89 mm SUL (mean 69.7 mm) in his museum-­based sample. Larvae. Larval L. yavapaiensis have a dusky yellow-­olive dorsum. The sides of the tadpoles are paler than the dorsum and have numerous obscure dark spots. The venter is white with a yellow cast to the throat. The tail muscle is olive gray with scattered dark gray spots, and tail fins are shallow and yellowish white, with discrete gray spots. The iris is bronze. The larvae reach a TL of ca. 77 mm, with the tail nearly twice the body length. A detailed description of larvae and mouthparts is in Scott and Jennings (1985). Eggs. The vitellus is surrounded by 3 envelopes. Storer (1925) provided the only mea­sure­ments of the eggs and noted they had already begun development: vitellus 1.78– 2.11 mm (mean 1.97 mm), inner capsule 2–2.5 mm (mean 2.25 mm), ­middle capsule 2.22–2.72 (mean 2.35 mm), and outer capsule 4.23–4.78 (mean 4.48 mm). Eggs are deposited in an irregular jelly mass about 75 mm in dia­meter.

ETYMOLOGY

yavapaiensis: named for Yavapai County, Arizona, where the holotype and paratype specimens ­were collected. The county was named for the Yavapai Nation. NOMENCLATURE

Stebbins (2003): Rana yavapaiensis Dubois (2006): Lithobates (Lithobates) yavapaiensis Fouquette and Dubois (2014): Rana (Lithobates) yavapaiensis Synonyms: None Leopard frogs have long been recognized for their phenotypic variation. Dif­fer­ent phenotypes have been considered to reflect polytypic variation of a wide-­ranging species (L. pipiens), to be the result of clinal variation, or to be members of a wide-­ranging multispecies complex (the Leopard Frog complex) (Ruibal, 1957; Moore, 1975; Hillis, 1988). Some assessments (e.g., Moore, 1944) mixed individuals from a variety of locations, resulting in considerable taxonomic confusion. Much of the scientific lit­er­a­ture uses the name Rana pipiens for frogs now recognized as Litho­ bates yavapaiensis, or refers to them as the “lowland type” or form (e.g., Frost and Platz, 1983). Readers should verify locations when using the older lit­er­a­ture. IDENTIFICATION

Adults. Lithobates yavapaiensis is a smooth-­skinned, light brown, gray-­brown, or olive leopard frog with prominent, lighter-­colored dorsolateral folds. ­These folds are poorly defined and are interrupted posteriorly and deflected medially. The typical pattern is discontinuous, with the posterior section inset medially. The inset has a series of dots with minimal or no similarity to the raised granular tissue of the primary fold (see Fig. 1(5) in Pauly et al., 2020a). Dark brown spots are pre­sent, but they may be pale brown in

Dodd_Canada_int_5pgs_B4.indd 641

DISTRIBUTION

Lithobates yavapaiensis historically was found from the Bill Williams River drainage of eastern Mohave County,

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642  RANIDAE

Despite similarities, ge­ne­tic analy­sis has confirmed that ­these leopard frogs are the invasive L. berlandieri. Morphological characters used to distinguish ­these species overlap in the frogs now found in California, which complicates field identification. Lithobates berlandieri in this region show extensive variation in the condition of the inset of the dorsolateral folds, a key character, more so than they do in their native range (Pauly et al., 2020a). FOSSIL REC­O RD

No fossils are known. Holman (2003) noted Miocene (Hemphillian) fossils from Navajo County in northern Arizona that belonged to the L. pipiens complex. The relationship between ­these fossils and L. yavapaiensis is unknown. SYSTEMATICS AND GEOGRAPHIC VARIATION Distribution of Lithobates yavapaiensis. Dark gray indicates extant populations; light gray indicates extirpated populations.

Arizona ( Jones, 1981), southeast in an arc to Hidalgo, Catron, and Grant counties in southwestern New Mexico. Additional populations occur in Santa Cruz and Cochise counties, Arizona, and in Surprise Canyon in the western part of the ­Grand Canyon. In México, the species occurs in Sonora, but ­little is known about its status ­there. It once was widespread in riparian habitats in southern Arizona (Gila and Salt drainages) and possibly along the lower Colorado River into México, but ­these populations have been extirpated (e.g., in the Santa Catalina Mountains; Lazaroff et al., 2006). Likewise, ­there have been no reports of Lowland Leopard Frogs in California (San Felipe–­ Carrizo drainages eastward to the Colorado River in Imperial, Riverside, and San Bernardino counties) since 1965, and the species is likely extirpated in the state. ­There are no reports from New Mexico since 1985. Grismer (2002) stated that Clarkson and DeVos (1986) observed 2 L. yavapaiensis in the border region with Baja California in 1981, but the latter authors reported seeing frogs of the “leopard frog complex.” ­These could have been L. berland­ ieri. Impor­tant distributional references include: Jones (1981), Platz (1988b), Jennings and Hayes (1994a, 1994b), Degenhardt et al. (1996), Gelczis and Drost (2004), Brennan and Holycross (2006), Oláh-­Hemmings et al. (2009), Bezy and Cole (2014), Murphy (2019), Pauly et al. (2020a), and Holycross et al. (2021). Although extirpated since 1965 in California, leopard frogs resembling L. yavapaiensis have been observed in habitats formerly occupied by that species in the state.

Dodd_Canada_int_5pgs_B4.indd 642

Lithobates yavapaiensis is a member of the Novirana clade of North American ranid frogs. It is an associate of the mostly lowland and tropical leopard frog group (or Scurrili­ rana) (Hillis and Wilcox, 2005). Its closest relatives in the United States include L. berlandieri, L. blairi, L. onca, and L. sphenocephalus. Microsatellite markers ­were characterized by Savage and Jaeger (2009). Oláh-­Hemmings et al. (2009) affirmed the close, but distinct, relationship between L. onca and L. yavapaiensis and noted that ­there are 2 distinct evolutionary lineages within the species. One group includes the populations in Arizona and northern México, whereas the other group is found only in Surprise Canyon in the western ­Grand Canyon. Lithobates onca and L. yavapaiensis diverged during the early Pleistocene. Lithobates yavapaien­ sis and L. onca also have closely related but distinct antimicrobial skin peptides (Conlon et al., 2010). Variation in leopard frog phenotypes from the American Southwest has been recognized for some time. The species can be separated from other leopard frogs by a combination of morphological, biochemical, auditory, and ge­ne­tic characteristics (Platz and Platz, 1973; Platz and Mecham, 1979; Frost and Platz, 1983). Frost and Bagnara (1976) presented a ­table comparing vari­ous phenotypes among leopard frog populations. ­Under laboratory conditions, crosses between ♀ L. yavapa­ iensis and ♂♂ of other members of the Leopard Frog complex (L. chiricahuensis, L. magnaocularis, L. pipiens from Vermont) can produce relatively high rates of hybrid offspring (Platz and Frost, 1984). ­These hybrids, however, do not produce offspring when backcrossed with parental species. Crosses between L. forreri and L. yavapaiensis ­were unsuccessful, as ­were crosses between ♂ L. yavapaiensis and ♀ L. pipiens from Arizona.

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Lithobates yavapaiensis 643

ADULT HABITAT

This species occurs in small, slow-­flowing streams and rivers, springs, and rock pools in Sonoran Desert Scrub and pinyon-­ juniper communities. Rocky rivers, small, intermittent creeks, open marshlands surrounding streams, and even the grassy marshes along large rivers such as the lower Colorado ­were inhabited. Jennings and Hayes (1994a) noted that man-­made stock ponds, ditches, and canals ­were occupied in the Imperial Valley of California. Preferred microhabitats include small spring runs where they enter large streams, which provide potential refugia ­under streamside debris piles. Deeper plunge pools with tree root masses and vegetation that provide protection against desiccation and predators also are favored, as are areas with moderate stream gradients that minimize the effects of flashflooding (Wallace et al., 2010). In southeastern Arizona, Wallace et al. (2010) found them in pools averaging 1.4 m in depth and 94.4 m3 per pool. This species is reported from habitats dominated by bulrushes, cattails, and riparian grasses near or ­under a partial cottonwood-­willow canopy ( Jennings and Hayes, 1994a). In a survey of intermittent streams inhabited by this species in the Sonoran Desert, Zylstra et al. (2015) found them in pool complexes (1–17 pools per complex) that ranged from 10.1 to 32.9 km2 in area at elevations of 849–1,313 m. ­Water availability averaged 63% but varied considerably among seasons. In New Mexico, the species occurs to an elevation of about 1,700 m. Not all potential habitats are occupied. For example, Wallace et al. (2010) found them at 15 of 33 sites surveyed in southeastern Arizona. Jennings and Hayes (1994a) provided historical photo­ graphs of former habitat along the lower Colorado River near ­Needles and at Harper’s Well Wash (San Felipe Creek) in Imperial County.

24°C) per note. Brennan and Holycross (2006) characterize it as a “low, grunting noise that resembles the sound of a fin­ger stroking an inflated balloon.” The first note is shorter than the following 6–15 notes, and the notes last 3–8 sec depending upon the number of notes and temperature. Pauses between notes and the duration of the notes tend to decrease as the call sequence progresses. Pulse rates average 12/sec (11 pulses in the first note to 3–4 in the last of the series) at a dominant frequency of 1.8 kHz (Frost and Platz, 1983). This produces a somewhat high-­pitched call that sounds something like an insect. Frost and Platz (1983) provided a sonogram comparing this species’ advertisement call with ­those of other Southwest leopard frogs. BREEDING SITES

Breeding takes place in shallow ­water in pools, tinajas (bedrock pools with ­little or no soil deposition along the bank), and springs, especially along intermittent streams. Optimal sites have vegetated banks with some canopy cover. REPRODUCTION

Breeding in this species is tied to predictable stream flows when eggs and larvae are not likely to be washed away by floods or adversely affected by drought. This species breeds ­earlier than the other Southwest ranids. In the California desert, breeding occurred from January to March, when ­water temperatures ­were 10–18°C (Ruibal, 1962). In Arizona, the breeding period is from late January to late May (Collins and Lewis, 1979; Frost and Platz, 1983; Degenhardt et al., 1996; Sartorius and Rosen, 2000). In addition, this species is known to deposit eggs in September and October (Platz and Platz, 1973; Collins and Lewis,

AQUATIC AND TERRESTRIAL ECOLOGY

­ here is ­little information available on habitat use. IndividuT als likely stay within close proximity to aquatic habitats. Daily or seasonal movements are likely localized and take place along watercourses, much as in L. onca, but longer distances may be traversed between pool complexes during favorable wet weather. Seim and Sredl (in Sredl, 2005a) found that juveniles ­were associated with small pools and marshy areas, whereas adults ­were found more often in larger pools. During drought conditions, the frogs retreat into mammal burrows, rock cracks, and fissures, and deep within dry mud cracks (in Sredl, 2005a). CALLING ACTIVITY AND MATE SE­L ECTION

The advertisement call of L. yavapaiensis is a series of short chuckles (or a trill) with a low number of pulses (12 at

Dodd_Canada_int_5pgs_B4.indd 643

Eggs of Lithobates yavapaiensis. Photo: Cecil Schwalbe

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644  RANIDAE

1979; Frost and Platz, 1983; Sartorius and Rosen, 2000). Goldberg (2019h) reported the smallest mature male at 43 mm SUL, and the smallest mature female at 53 mm SUL. He further described oogenesis and spermiogenesis and suggested that L. yavapaiensis has some degree of spawning activity in all 4 seasons of the year. Eggs are deposited in spheroidal masses in shallow ­water. The masses are attached to vegetation, bedrock, or gravel up to 1 m below the ­water’s surface. Most masses, however, are at or just below the surface. Late spring egg masses may be only half the size of ­those oviposited ­earlier in the spring. Eggs take 15–18 days to hatch in 14°C ­water, and most egg masses ­will produce larvae (Sartorius and Rosen, 2000). Eggs exposed to air experience the most mortality. Normal development for most leopard frog populations occurs at ­water temperatures >8°C. However, the minimum temperature for egg development was 11°C in the California desert (Ruibal, 1962). The maximum temperature that L. yavapaiensis eggs can survive is 28–29°C (Ruibal, 1962). LARVAL ECOLOGY

­ here is very ­little information available on larval ecol­ogy. T Larvae have been found throughout the year at some localities. Some larvae overwinter, perhaps from the autumnal breeding period. Sredl (2005a) gives the larval period as 3–4 months or as long as 9 months. Newly transformed froglets are 25–29 mm SUL (Platz, 1988b). In experimental ­trials, larval L. yavapaiensis are able to detect the presence of the nonnative sunfish Lepomis cyanellus and respond by swimming faster when chemical cues from this predator are pre­sent than they do in control treatments

(Shah et al., 2010). Larvae also reduce their levels of activity, and indeed develop dif­fer­ent body shapes, including deeper tail fins, larger tail muscle heights, and smaller tail muscle areas (Sosa et al., 2009). Still, ­these authors could not demonstrate a survival advantage to this phenotypic plasticity. Although tadpole be­hav­ior and morphology ­were altered in ­these mesocosm treatments, it is unknown ­whether similar be­hav­ior occurs in the wild or ­whether the altered be­hav­ior affects the survivorship of wild tadpoles. In any case, the tadpole reaction to the nonindigenous predator suggests that they are capable of rapid learning. DIET

Larvae are herbivorous. ­There is no published information on the diet of postmetamorphs of this species. Like other leopard frogs, they prob­ably feed on a variety of invertebrates and even small vertebrates in relation to their availability. PREDATION AND DEFENSE

Lithobates yavapaiensis seeks refuge among roots and streamside debris along small creeks. Jennings and Hayes (1994a) provided a photo­graph of a California L. yavapaien­ sis with its rear limbs held into the body and the forelimbs brought up over and shielding the eyes. The frog is crouched against the substrate. Perhaps such an immobile posture did not draw attention to the frog or made it more difficult for a small predator to attack. Tadpoles may be able to detect chemical cues emanating from predators and alter their be­hav­ior accordingly. ­There is ­little information on predation other than Jones’ (1990) and Jones and Hensley’s (2020) reports of larvae and adults being consumed by the Garter Snake Thamnophis cyrtopsis. Thamnophis eques also eats this species ( Jones et al., 2020). Ravens (Corvus corax) and Turkey Vultures (Cathartes aura) also take larvae and metamorphs (Brattstrom, 2019). POPULATION BIOLOGY

Tadpole of Lithobates yavapaiensis. Photo: Cecil Schwalbe

Dodd_Canada_int_5pgs_B4.indd 644

Populations of adults breeding in selected pools may be small. For example, Sartorius and Rosen (2000) counted only 3–20 frogs per transect in surveys conducted from February to September at a population in the Santa Catalina Mountains. Survivorship was lowest during the autumn and winter months, and highest in spring and summer. Wallace et al. (2010) reported up to 104 frogs per survey (mean 15.4) in 15 canyons in southeastern Arizona. Frogs live at least 3 yrs post-­metamorphosis (Sredl, 2005a). As expected for a ranid frog living in the desert, stream dynamics play a critical role in this species’ population dynamics. Based on 16 yrs of survey data involving 33 pool

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Lithobates yavapaiensis 645

are not likely to be recolonized if distances between sites are large (2–8 km). DISEASES, PARASITES, AND MALFORMATIONS

Adult Lithobates yavapaiensis. Photo: Dennis Suhre

complexes in 4 canyons, Zylstra et al. (2015) found that monthly adult survival rates ranged from 72 to 99% in summer and 59–94% in winter. Annual adult survival was estimated at 37%. Survival and recruitment rates increased with the extent of available surface ­water. Recruitment occurred primarily in winter and ranged from 1.9 to 3.8 individuals per season per pool. Frog abundance varied across years and seasons, but no discernible population trends ­were apparent. In a further analy­sis of 6 stream reaches in 2 catchment basins, Zylstra et al. (2019b) found low annual survivorship ranging from 7 to 14% that varied with temperature and surface ­water availability. When ­water was at 100% of maximum pool depth, monthly survival was 88%; it was 81% at 50% ­water depth, and was only 36% at 20% maximum ­water depth. Severe drought contributed to population extirpation at 1 of the catchments studied. Localized extinctions are more common during drought years, as expected, and colonization rates increase when larval and dispersal periods occur during wetter than normal years (Zylstra et al., 2019a). Frogs can only persist at sites where ­water occurs consistently and where sediment loads from upstream regions are low. The location of pools is patchy and, in conjunction with climate variation, the overall population structure is one of a metapopulation in constant states of change. In Zylstra et al.’s study, 52% of the pool complexes initially surveyed (1996) ­were occupied by L. yavapaiensis, but this dropped to 0% in their northern population complex by 2015, and 38–61% in their southern pool complex. Drought, combined with a lack of surface ­water, inhibited per­sis­tence and recolonization of habitats. In the absence of drought, populations are unlikely to be extirpated. Following extreme droughts, however, habitats

Dodd_Canada_int_5pgs_B4.indd 645

Lithobates yavapaiensis possesses antimicrobial skin peptides that may aid in defense against Escherichia coli and Staphy­ lococcus aureus (Conlon et al., 2009). The virulent pathogen Batrachochytrium dendrobatidis (Bd) has been found in fairly high prevalence (10 of 13 populations examined) in this species in Arizona (Bradley et al., 2002; Schlaepfer et al., 2007). Fully 43% of the Lowland Leopard Frogs examined ­were positive for Bd. Schlaepfer et al. (2007) suggested that the pathogen becomes virulent during the winter in conjunction with other ­factors, and that thermal springs may offer refuge from the pathogen. Forrest and Schlaepfer (2011) found an inverse relationship between ­water temperature and chytrid presence, with chytrid detection dropping dramatically at ­water temps of >30°C. Indeed, the prevalence of the pathogen varies spatially and temporally. Over a 5 yr period covering 12 localities, Savage et al. (2011) found that 2 populations never exhibited Bd, 10 populations exhibited Bd in winter, and 5 populations had winter mortality. No mortality was observed in summer, and Bd prevalence was low (1.6%) in infected populations. In winter, infection intensities decreased from 2008 to 2010, although mortality and infection prevalence did not change throughout the winter among the 10 localities. Savage et al. (2011) also noted higher mortality rates at non-­thermal springs in winter as opposed to thermal springs, further implicating higher temperature as having a positive influence on survivorship. Large frogs had higher mortality rates and higher infection intensities than survivors. Lowland Leopard Frogs are parasitized by the trematodes Cephalogonimus brevicirrus, Glypthelmins quieta, Haema­ toloechus complexus, and Megalodiscus temperatus. They also are parasitized by the nematodes Falcaustra catesbei­ ana, Physaloptera sp., and Rhabdias ranae (Goldberg et al., 1998a). SUSCEPTIBILITY TO POTENTIAL STRESSORS

NaCl. Lowland Leopard Frogs ­were found historically in brackish ­water along desert streams (Ruibal, 1959). Ruibal (1959) recorded salinities of 1.75–9.0 ppt at locations with frogs, with reproduction occurring at salinities of 2–2.75 ppt. This population no longer exists. STATUS AND CONSERVATION

At one time, this frog was quite abundant in southern Arizona. For example, King (1932) noted they ­were common

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646  RANIDAE

Habitat of Lithobates yavapaiensis. Photo: Dennis Suhre

along the stream in Sabino Canyon in 1930. Populations in the Santa Catalina Mountains, in riparian habitats of southern Arizona (Gila and Salt river drainages), and along the lower Colorado River have been extirpated (Vitt and Ohmart, 1978; Lazaroff et al., 2006), as have populations in the Imperial Valley of California ( Jennings and Hayes, 1994a, 1994b). Lowland Leopard Frogs ­were not found in previously occupied habitats in California during surveys from 1983 to 1987 (Clarkson and Rorabaugh, 1989). Jennings and Hayes (1994a) summarized the historical distribution and decline of this species in that state. The last verified collection of Lowland Leopard Frogs in California was from near Calexico in 1965. Lithobates yavapaiensis

Dodd_Canada_int_5pgs_B4.indd 646

appears to have dis­appeared from New Mexico by 1985 (Degenhardt et al., 1996). Clarkson and Rorabaugh (1989) reported that populations in upland areas of Arizona ­were still largely intact as late as 1987, and Sredl (1998) noted fairly high occupancy rates at historical localities, with many new populations discovered. Still, Griffis-­Kyle et al. (2018) considered this species one of the most vulnerable amphibians in the Desert Southwest ­because of its dependence on springs and stream habitats that are threatened throughout the region. The likely ­causes of decline include a variety of ­factors, especially habitat alteration and outright destruction as their springs, riparian environments, and rivers ­were modified and ­water reallocated for ­human use. Altered hydrology coupled with increasing drought due to climate change are the biggest threats to this species. In order to conserve this species, riparian areas must be managed to allow per­sis­tence and connectivity among populations (Wallace et al., 2010; Zylstra et al., 2015, 2019a, b). Vegetation changes and the introduction of L. catesbeia­ nus into many isolated spring habitats likely hastened population declines. Competition also may have occurred between this species and the introduced Rio Grande Leopard Frog (L. berlandieri), which has become widespread in southern Arizona (Clarkson and Rorabaugh, 1989). Bury and Lukenbach (1976) noted that some leopard frogs in the lower California deserts did not appear to be native, and L. berlandieri is now known to be widespread in former L. yavapaiensis habitat ­there. Ranid declines in Arizona, including L. yavapaiensis, are correlated with the presence of introduced crayfish and Bd (Lazaroff et al., 2006; Witte et al., 2008). Declines are thus the product of multiple ­causes acting in­de­pen­dently or cumulatively on a population-­by-­population basis. Evoked vocal responses have been used to improve detection during surveys for this species. By using evoked calls in initially ­silent populations, detectability improved by 71%. However, responses to evoked calls ­were not entirely species specific, as L. yavapaiensis males also responded to male L. chiricahuensis calls about 45% of the time (Schlaepfer et al. 2021). Much information on the ecol­ogy and life history of L. yavapaiensis is contained in unpublished reports. Sredl (2005a) summarizes some of this information. The species is protected by state law in Arizona and is considered Endan­ gered in New Mexico.

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Rana aurora 647

Rana aurora Baird and Girard, 1852 Northern Red-­legged Frog Grenouille à pattes rouges du Nord ETYMOLOGY

aurora: Latin for ‘red,’ referring to the frog’s coloration.

ranids. They are deposited in a loose and viscid jelly mass 15–25 cm in dia­meter, with each mass containing 500–1,300 eggs. The bluish-­tinged mass is attached to vegetation at the surface of the ­water. ­There are 3 indistinct envelopes surrounding the egg. The outer envelope is 10–14 mm in dia­meter; the ­middle envelope is 6.25–7.9 mm (mean 6.8 mm) in dia­meter; and the inner envelope is 4–6.7 mm (mean 5.7 mm) in dia­meter. The vitellus is 2.3–3.6 mm in dia­meter. Eggs ­were described by Livezey and Wright (1947).

NOMENCLATURE

Dubois (2006): Rana (Amerana) aurora Fouquette and Dubois (2014): Rana (Rana) aurora Synonyms: Rana agilis aurora, Rana temporaria aurora IDENTIFICATION

Adults. This is a large (males to 68 mm SUL, females to 101 mm SUL; Storm, 1960; Jennings and Hayes, 1994b), rather smooth-­skinned frog, although some individuals have a granular skin texture. The dorsum is brown, reddish brown, or greenish gray and peppered with small black spots with a few large spots in between. A prominent dorsolateral fold consisting of granular glands is pre­sent, and ­there usually is a light lip-­line from the eye to the shoulder, bordered above by a dark mask. The venter is heavi­ly infused with gray. The groin has prominent, irregular, black and gray mottling. The lighter mottling may range from white to red to green to yellow. The venter is yellow with a superimposed tinting of red. The undersides of the arms, legs, and abdomen are red; this red coloration is not apparent in juveniles. Legs are long. Male Rana aurora usually lack a vocal sac (Hayes and Krempels, 1986). Dunlap (1955) and Dumas (1966) provide extensive information on morphological comparisons with other Pacific Northwest ranids. Females are larger than males. Adult females ­were 72–93 mm SUL (mean 84 mm) and males ­were 49–65 mm SUL (mean 59 mm) in an Oregon population (Storm, 1960). In British Columbia, males ­were 45–60 mm SUL and females ­were 62–80 mm SUL (Licht, 1974). Larvae. The tadpole is tan to dark brown with numerous, scattered clumps of golden flecks. The venter is off-­white or brassy and iridescent in coloration, and ­there are 2 or 3 transverse bands of clumped, brassy pigment cells. The dorsal and ventral tail fins are the same depth as the tail musculature, and they may be marked with minute dark spots and have a golden tone. Small tadpoles (10°C (Licht, 1969c). Movement to breeding sites begins soon ­after emergence. Movements occur at night and are facilitated by rain and cloud-­cover. ­After breeding, the adults disperse into thick, moist vegetation (Twedt, 1993). Like California Red-­legged Frogs, Northern Red-­legged Frogs are sometimes found at significant distances from ­water. Dumas (1966), for example, recorded them up to 914 m from the nearest ­water, whereas Nussbaum et al. (1983) noted terrestrial movements 200– 300 m from ­water. In a mark-­recapture study, Hayes et al. (2001) caught both males and females from 1.1 to 2.4 km (straight-­line distance) between captures. ­These results suggest that R. aurora can make terrestrial movements of considerable distances ­either between breeding sites or from breeding sites to terrestrial refugia and foraging habitat. CALLING ACTIVITY AND MATE SE­L ECTION

Calling normally occurs from midwinter to early spring, although advertisement calls have been heard in September (Leonard et al., 1997b). Males arrive at breeding sites prior to females, and are active diurnally, sitting around the margins of ponds and on stream banks in full sunlight. ­After about a week, vocalization begins as temperatures increase to >6°C consistently over several days; once initiated, calling can occur at temperatures as low as 4°C (Storm, 1960; Calef, 1973b; Brown, 1975b). Males usually call nocturnally while completely submerged in ­water usually >60 cm in depth (range 16 cm–1.8 m; Licht, 1969c). Occasional calls are made during the day, especially in the early morning. Frogs call from the bottom of the pond or river while concealed in vegetation; they ­will surface for air occasionally. Male R. aurora position themselves a few meters apart from other males and call from locations more than 1 m from the pond or stream shoreline. They apparently do not defend territories, and overt aggression does not occur. Calling frequency increases in the vicinity of another male, and indeed any movement near a calling male ­causes him to call continuously for up to 30 sec. Movements by calling males back and forth between vari­ous locations within a wetland or pond may occur during the breeding season. Most such movements are for short distances, although Calef (1973b) recorded a few frogs moving >300 m. The advertisement call of R. aurora is low in volume and has poor carry­ing capacity in air (normally 15 m away, although calls given underwater can be detected 10 m distant using underwater hydrophones. Licht (1969c) recorded listening to a “resounding” chorus of R. aurora using hydrophones, a chorus that could not be detected out of ­water! The call consists of 2–5 notes (usually 3–4), with each note consisting of 5–6 pulses. The dominant frequency ranges between 450 and 1,300 Hz, with some notes extending to 7.5 kHz. Calls last about 1 sec depending on the number of notes. Storm (1960) and Licht (1969c) described the call as a repetitive series of “uh uh uH UH,” and Licht (1969c) included a sonogram. Amplexus is axillary (Storm, 1960). A male may attempt to amplex a female even prior to beginning his vocalizations. When a female is receptive, amplexus leads to egg deposition. Males are tenacious. If unreceptive, the female cannot dislodge a per­sis­tent male simply by uttering a release call with its associated abdominal vibrations. Instead, she must roll to her side while extending her back legs. She remains in this position with all legs stiff and extended ­until the male releases her (Licht, 1969b). Even so, a male may remain clasped to a posturing female for as long as 15 min. Kicking ­toward the male or assuming an arched body posture is not as effective in discouraging an ardent male. The Northern Red-­legged Frog also has a release call that is directed ­toward conspecific males should amplexus be attempted. Many males may try to amplexus a female si­mul­ta­neously. Amplexus has been recorded with other ranids, Ambystoma gracile, and even an apple (Storm, 1960; Nussbaum et al., 1983). Males make 2 dif­fer­ent calls during amplexus that are distinct from the advertisement call. The first is directed ­toward receptive females and is uttered about once per second more or less continuously. This call is a single unpulsed note of 750–1,500 Hz that is directed into the female’s ear. A second amplexus call, more like the male’s release call, appears directed ­toward unreceptive females and is uttered in a series of 1–8 notes that are structurally dif­fer­ent from the amplexus call directed ­toward receptive females. When a female attempts to displace a male, he increases the speed, intensity, and repetition rate of this amplexus call. Males make this call 2 or 3 times a minute when clasped to an inactive female, but if the female attempts to displace him the repetition rate increases to 10–13 times per minute. Licht (1969b, 1969c) provided sonograms of ­these clasping calls and photo­graphs of a female’s be­hav­ior while discouraging a suitor. BREEDING SITES

Breeding sites are varied, from temporary to permanent wetlands to slow-­moving (25 cm–5 m in depth and at least a meter (to 4.5 m) from the shoreline of the pond. In British Columbia, Licht (1971) found that egg masses ­were attached to submerged vegetation in at least 46 cm of ­water. Normally, the egg mass is completely submerged, although portions of egg mass might break ­free of the vegetation and float to the surface. In streams, egg masses are deposited in deep ­water (60 cm–1.2 m) in the deepest part of the channel, usually on the downstream side of the current. Egg masses may take on a greenish tint due to algal growth as development proceeds. Algae produce oxygen, which is beneficial to the developing embryos. Egg masses are spaced several meters apart in the immediate vicinity of the calling location of the successful male. Thus, ­there is an evenness of spacing, although Licht (1969c) recorded 10 masses in a 1.8 m2 area. Egg masses are positioned so that they ­will be exposed to direct sunlight in ­water with virtually no flow. However, some egg masses ­will be lost as ponds desiccate or due to the sudden onset of freezing temperatures. Eggs also experience increased mortality in habitats containing large quantities of silt, perhaps in conjunction with the physico-­chemical properties of the soils involved (Platin and Richter, 1995). Females likely breed ­every year and stay at a breeding pond only ­until they oviposit. Males stay throughout the breeding period. LARVAL ECOLOGY

Egg mass of Rana aurora. Photo: Brome McCreary

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The period of embryonic development lasts about 35 days in ­water that ranges from 4.5 to 7.8°C (Brown, 1975b); Storm (1960) recorded embryonic development over a period of 35–50 days ­under field conditions. In contrast, Rombough and Trunk (2017) recorded development (deposition to hatching) in only 17–33 days. Eggs deposited early in the season when ­waters are cold take longer to develop than eggs deposited late in the season, accounting for ­these discrepancies. The jet-­black larvae hatch at 11–12 mm TL (Storm, 1960; Brown, 1975b), and the larval period is 110 days in ­water ranging from 7.3 to 23.5°C

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density, although tadpoles reaching a large size have a decreased predation rate and a greater chance of survival than small tadpoles. Food does not appear to be a limiting ­factor, as tadpole survivorship is in­de­pen­dent of density, even at densities of 75 larvae per square meter. Calef (1973a) estimated that high tadpole mortality occurred in the first 4 weeks ­after hatching, with higher survivorship through time as the tadpoles grew. Only 5% of the tadpoles that hatched ­were estimated to reach metamorphosis. Licht (1974) obtained a similar estimate. DIET Tadpole of Rana aurora. Photo: William Leonard

(Brown, 1975b). Calef (1973a) estimated the larval period in British Columbia as 11–14 weeks, with tadpole developmental rates directly correlated with the number of degree-­days since hatching. Larvae are significant grazers on algae and have an impor­tant role in structuring the periphyton community (Dickman, 1968). Transformation occurs at about 29 mm TL, with recently completely metamorphosed juveniles ranging from 20 to 29 mm TL (Storm, 1960; Calef, 1973a; Nussbaum et al., 1983). Brown (1975b) reported that larvae reach a maximum size of 34 mm TL at Gosner stage 39, but Storm (1960) found larvae nearly 60 mm TL. ­Because egg masses contain many eggs, and ­there may be many masses deposited in a pond, tadpole densities immediately ­after hatching can reach 500 per m2 (Calef, 1973a). In laboratory tests, tadpoles associated with siblings during early development (Blaustein and O’Ha­ra, 1986), but they soon dispersed rather rapidly and evenly across a pond’s substrate. Subsequently, no kin-­based associations ­were observed. Tadpoles are capable of selecting certain substrate features based on early experience, and they can retain their preferences throughout development. They also can develop a substrate preference at any stage of development, so ­there is no early learning or imprinting. Wiens (1970) suggested that a ready capacity to learn suitable substrate patterns would be advantageous to tadpoles, especially ­those that hatched initially onto unsuitable substrates. However, tadpoles reared in featureless environments did not show a substrate preference. Predation can have a significant effect on larval survivorship. In experimental mesocosms in natu­ral lakes, larvae ­were twice as likely to survive when salamanders (Taricha granulosa) ­were absent as when salamanders ­were pre­sent (Calef, 1973a). Tadpoles are eaten in proportion to their

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Northern Red-­legged Frogs are mostly insectivorous, with larger frogs eating larger prey than smaller or recently metamorphosed frogs (Licht, 1986b). The diet consists largely of spiders, beetles, aphids, flower flies, leafhoppers, crane flies, spittlebugs, caterpillars, isopods, and other small invertebrates (Fitch, 1936; Licht, 1986b). PREDATION AND DEFENSE

The most impor­tant predators on postmetamorphic R. au­ rora are garter snakes (Thamnophis sp.). Adult R. pretiosa feed upon small R. aurora (Licht, 1986b), and juveniles and adults are preyed upon by American Bullfrogs (Lithobates catesbeianus) (Twedt, 1993; Kiesecker and Blaustein, 1997a; O’Loughlin et al., 2016). Birds (herons) and mammals (raccoon, mink) also likely take postmetamorphs, and predation by Northern Harriers has been reported on adults (Todoroff, 2021). Eggs (Licht, 1969a) and tadpoles are highly palatable. Larvae are eaten by Common Garter Snakes (Thamnophis sirtalis), larval and paedogenic adult salamanders (Taricha granulosa, Ambystoma gracile), fish (Salmo gairdneri), and predaceous aquatic insects (­giant diving bug Lethocerus americanus; dragonfly naiads) (Storm, 1960; Calef, 1973a). Eggs and embryos are eaten by leeches (Batrachobdella picta) and amphipods (Crangonyx spp.) (Hudgens and Harbert, 2019). Like many anurans, larval Northern Red-­legged Frogs appear to be able to detect predator chemicals in their environment and alter their be­hav­ior accordingly. For example, small larvae reduce their levels of activity in the presence of chemicals emanating from newts (Taricha granulosa) fed on a diet of tadpoles, but not from newts fed on insects. They also are able to detect chemicals from injured or disturbed conspecifics (Wilson and Lefcort, 1993; Kiesecker et al., 1999). Indeed, disturbed tadpoles show an increased amount of ammonium secretion, and larvae exposed to increased levels of ammonium respond with antipredator be­hav­ior similar to that of disturbed tadpoles (Kiesecker et al., 1999). In contrast, Adams and Claeson (1998) could not demonstrate trap avoidance by large

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R. aurora tadpoles in response to damage release chemicals. Response to ­these chemicals may be related to size (large tadpoles are not as prone to predation as small tadpoles), or perhaps the chemosensory response does not occur ­under field conditions. Changing be­hav­ior in the presence of an unseen threat would likely enable larvae to decrease their risk of predation, so it seems plausible that a lack of response might be size related. Adult Northern Red-­legged Frogs are wary, especially during the day. At night, however, males may be approached easily. They remain stationary on land ­until a certain approach distance has been breached, then quickly jump into the ­water. This approach distance is farther for large predators (ca. 1 m) than it is for small predators (80 sites in the debris-­ avalanche zone, with breeding activity at 40 locations (Crisafulli et al., 2005). As time passes and ponds become suitable, R. aurora should reoccupy virtually all its former territory in the region. POPULATION BIOLOGY

Many Northern Red-­legged Frogs may deposit eggs in a single pond. For example, Calef (1973a, 1973b) counted 618 and 620 egg masses over a 2 yr period at a breeding pond in British Columbia and estimated the male breeding popula-

Adult Rana aurora. Photo: David Dennis

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tion as 1,770 one year and 3,600 the next. He estimated that ­there ­were initially 300,000 tadpoles pre­sent, a number that quickly fell to 75,000 ­because of predation. Only about 15,000 tadpoles (5%) metamorphosed. In another British Columbia study, Licht (1974) found that only 52% of metamorphosing juveniles survived the first year. Once reaching adulthood, survivorship can be high (69%) from one year to the next. Many authors have speculated that the size at and timing of metamorphosis influence postmetamorphic fitness. Chelgren et al. (2006) tested this hypothesis with R. aurora and found that large metamorphs had higher rates of survival than small metamorphs, and that survival was inversely proportional to the number of days metamorphosis was delayed ­after the first metamorphs emigrated from the breeding pond. Thus, ecological ­factors within the pond that delay growth (e.g., food resources, density, competitors) have impor­tant subsequent effects on metamorph survival and fitness. Juveniles are capable of rapid growth (0.09–0.18 mm/ day). Sexual maturity is reached by males at 45–50 mm SUL the first year following metamorphosis; females attain sexual maturity at 60 mm SUL during the second year following metamorphosis (Licht, 1974; Hayes and Hayes, 2003). Several authors have estimated longer times to maturity, such that individuals do not reach maturity ­until their third or fourth year following metamorphosis (Nussbaum et al., 1983; Leonard et al., 1993; Jennings and Hayes, 1994b); ­these may be overestimates, although more data are needed from multiple sites. Longevity is >10 yrs based on rec­ords in captivity (Cowan, 1941). DISEASES, PARASITES, AND MALFORMATIONS

Eggs that do not develop or are placed in very cold ­water are colonized quickly by fungus. Larvae are infected by the parasitic yeast Candida humicola (Lefcort and Blaustein, 1995). Tadpoles infected by this yeast have higher thermal preferences (“behavioral fever”) than normal tadpoles, and they exhibit an inability to discriminate chemosensory cues from potential predators. In experimental ­trials, they suffered increased levels of predation when compared with noninfected tadpoles (Lefcort and Blaustein, 1995). ­These results suggested that the pathogen had sublethal effects that decreased the ability of the larvae to survive to metamorphosis, regardless of the direct effects of infection. Iridovirus has been reported from larval R. aurora in northern California (Mao et al., 1999). Batrachochytrium dendrobatidis (Bd) has been found in larvae from Redwood National Park in northern California at a rate of 6.4% of ­those examined, especially ­those with deformed oral disks (Nieto et al., 2007). Additional reports of suspected or confirmed chytridiomycosis are in Green et al. (2002),

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Adams et al. (2010), and Ecoclub Amphibian Group et al. (2016). Unlike tadpoles infected by parasitic yeast, Northern Red-­legged Frog larvae experimentally infected with Bd show no evidence of behavioral fever or altered thermoregulation (Han et al., 2008). Parasites of larvae include protozoans (Apiosoma sp., Epistylis sp., an unidentified trichodinid, Ichthyobodo sp.), the trematodes Brachycoelium lynchi, Gyrodactylus aurorae, and Ribeiroia sp., and unidentified leeches in the oral cavity (Ingles, 1936; Mizelle et al., 1969; Nieto et al., 2007). Adults are parasitized by leeches (Batrachobdella picta) (Licht, 1974). The trematode Ribeiroia ondatrae has been found in free-­living R. aurora. This trematode ­causes malformations, and 10 dif­fer­ent malformations ­were observed in 7.9% (9 of 114) of the frogs examined from throughout the Pacific Northwest ( Johnson et al., 2002). ­These included missing eyes, multiple limbs and other limb deformities, and cutaneous fusions of the hind limb. From 1 to 78 metacercariae ­were found per larva or recent metamorph. Other trematodes infecting R. aurora include Gorgoderina multilobata, Margeana (Glypthelmins) californiensis, and Megalodiscus microphagas (Zamparo and Brooks, 2005). Rana aurora and R. draytonii also have closely related but distinct antimicrobial skin peptides (Conlon et al., 2006). ­These peptides aid in the frog’s defense against bacterial pathogens. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Nitrates and sulfates. Larval survivorship is decreased at nitrate concentrations of 5 and 20 mg/L in the presence of the ­water mold Saprolegnia, but the nitrate itself did not significantly affect survivorship (Romansic et al., 2006). At higher concentrations, embryos of R. aurora suffer 100% mortality at >105 mg/L NH4-­N (ammonium nitrate) and 918 mg/L NO3-­N (sodium nitrate) (Schuytema and Nebeker, 1999a). Growth of R. aurora tadpoles is inhibited at concentrations >134 mg/L NH4-­N (Nebeker and Schuytema, 2000), and decreases in length or mass of R. aurora embryos are evident at >13.2 mg/L NH4-­N and 50 m from ­water, and Cook et al. (2012) observed 60 juvenile R. boylii in a residential area from 2008 to 2011 where they ­were found 16–331 m from a nearby stream. The largest single observation was of 40 juveniles on 1 night in October. In addition, numbers of juveniles ­were observed on nearby roadways, with numerous frogs killed by vehicles. The overwintering be­hav­ior is unknown, but overwintering may occur ­under rocks in the heavi­ly shaded and cool tributaries to the main stream channels or perhaps even ­under rocks in nearby forests within a few meters of a stream (Storer, 1925; Zweifel, 1955). Nussbaum et al. (1983) reported unearthing small R. boylii at a rock outcrop 50 m from a river in April. According to Kupferberg (1996), adults are seen in small tributaries prior to the mating season in spring. Storer (1925) suggested that this species does not enter dormancy ­because of the relatively mild temperatures throughout much of its range. In many of the small creeks, winter rainfall ­causes considerable scouring flow across rocky substrates, making dormancy within stream channels risky. Like most frogs, Foothill Yellow-­legged Frogs are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities ( Jaeger and Hailman, 1973). Rana boylii likely has true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males have a guttural, grating advertisement call that has 1 pitch or a slightly rising inflection. Rombough and Hayes (2005) described it as “a series (5–7 notes) of distinct, rubbery clucks.” According to Stebbins (1951), the frog sometimes utters groups of 4–5 croaks lasting ca. 0.5 sec each with ca. 0.5 sec intervals between croaks. This series may be followed by “a prolonged rattling sound lasting 2.5 seconds.” In addition, R. boylii produces several dif­fer­ent calls while submerged, suggesting an extensive vocabulary that is capable of conveying information to other Foothill Yellow-­legged Frogs. MacTague and Northen (1993) reported 4 dif­fer­ent calls produced underwater: a short, unpulsed call, pulsed calls of 2 intermediate lengths, and a long call where groups of pulses form notes. ­These calls could be used in a territorial context (advertisement and defense) as well as to attract mates. Rana boylii also makes a faint short, pulsed call when the frogs are handled. Calling is associated with ­water temperatures >10°C and ­water depths of >10 cm (peak ca. 20 cm); the onset of calling occurs ­later at deeper ­water depths and cooler temperatures, and varies annually and geo­graph­i­cally (Wheeler et al., 2018). Males arrive at the breeding sites and remain in the

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same location throughout the breeding season. Females arrive asynchronously throughout the breeding season and do not remain ­after oviposition has been completed. Wheeler and Welsh (2008) suggested that males defended optimum calling sites that may not be directly associated with an oviposition site; they observed male–­male aggression that indicated territorial defense. Not surprisingly, the daily operational sex ratios at breeding sites are male biased, although the seasonal operational sex ratio actually may be female biased (Wheeler and Welsh, 2008). It appears large males have an advantage when mating in comparison to small males, although strict size-­assortative mating does not occur. In Wheeler and Welsh’s (2008) study, for example, amplexing males ­were larger than males that never mated within a breeding season. Thus, mating is likely not random, but it is not known ­whether females choose mates or how amplexus is initiated. Occasionally, R. boylii ­will attempt to mate with other species, such as males amplexing female L. catesbeianus (Lind et al., 2003). BREEDING SITES

Breeding occurs only in relatively slow-­moving streams on a rock or boulder substrate, which allows for the development of sediment bars and backwater habitats. Preferred sites include river bars offering protection from swift currents and in proximity to the confluence of tributaries where rivers are wide and shallow. Oviposition also occurs in off-­channel scour pools or troughs. Frogs may deposit eggs in narrow or deep channels, but eggs oviposited in ­these areas have poor survival in both wet (from scouring effects) and dry (prone to desiccation) years. Optimal sites may be used repeatedly from one year to the next (Lind et al., 2016). REPRODUCTION

Emergence occurs several weeks prior to the initiation of breeding, and breeding is closely linked with stream hydrology. Reproduction occurs ­after the high-­flow stream discharge which results from winter rainfall and high-­ elevation snowmelt; it is correlated with warming air and ­water temperatures and decreases in stream flow. Reproduction is thus timed to reduce the threat of eggs being washed away via stream scouring. Based on histology, Goldberg (2019d) noted that spermiogenesis occurs in the spring and that females are capable of breeding from spring through fall in California. The smallest mature male was 33 mm SUL, whereas the smallest mature female was 44 mm SUL in his museum-­based sample. The breeding season of R. boylii lasts from 3 to 7 weeks, from late March to early June, depending upon elevation, location, and environmental conditions (Storer, 1925; Fitch, 1936; Wright and Wright, 1949; Zweifel, 1955; Jennings and

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Eggs of Rana boylii. Photo: Koen Breedveld

Hayes, 1994b; Jones et al., 2005; Wheeler et al., 2018). For example, the breeding season lasted 19–52 days (mean 49.5) in Del Norte County, California, over a 6 yr period (Wheeler and Welsh, 2008), and Kupferberg (in Wheeler and Welsh, 2008) found the breeding season lasted 18–63 days (mean 34) in Mendocino County, California. Plasticity in the timing of reproduction allows this species to take advantage of optimum stream conditions in an environment subject to stochastic changes in rainfall and river flow patterns. In contrast, Rombough and Hayes (2007) suggested that R. boylii was an explosive breeder in Oregon, and that the breeding season only lasted about a week. ­These differences may be related to differences in the availability of habitat and to physical differences in streams between locations. Breeding occurs over an extended period in spring, but the precise dates of reproduction depend on ­water flow and temperature. In Oregon, for example, breeding occurs from April to June (Nussbaum et al., 1983). In California, breeding begins from early April to early May, depending on river flows, with river flows of 0.10–0.6 m/s producing the greatest amount of breeding activity. River flow levels usually are optimal >30 days following the initiation of the breeding season, although reproduction may be halted or delayed by high stream flows that might dislodge eggs or wash them away. The initiation of breeding occurs ­earlier in low-­flow years than in high-­flow years and, as such, the amount and timing of precipitation may have a direct impact on the duration of the breeding season and an indirect impact on reproductive output. This is ­because the number of egg masses produced at a location may be correlated with maximum annual river flow during the breeding season. Oviposition also occurs ­later in cooler and deeper ­water.

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­ ater temperatures should be >9.5–12°C (Zweifel, 1955; W Kupferberg, 1996; Wheeler et al., 2018). Eggs are attached to substrates (cobbles and boulders, rarely on bedrock or vegetation) within the stream in shallow ­water (4–43 cm) of low velocity. Wide, shallow streams are preferred, with egg mass abundance positively associated with distance (ca. 0.5–4 m) to the shoreline (Lind et al., 2016). Attachment sites are on the downstream sides of cobbles and boulders, where stream velocity is slower than in the main channel. The location of egg deposition depends on ­water level. In high-­flow years, eggs may be deposited along the shoreline; in low-­flow years, eggs may be placed as much as 1.25 m from shore (Kupferberg, 1996). The location of the egg mass ensures adequate oxygenation as cool ­water passes around it. Kupferberg (1996) reported a mean of 19 clutches per site at 15 study sites in northern California, whereas Lind et al. (2016) counted 568–731 masses in 15.6 km of the South Fork Trinity River; at Hurdygurdy Creek, they counted 84–140 masses over 4.7 km. The greatest density of egg masses was reported by Bourque and Bettaso (2011) as 323.5 masses/km along the Mad River in Humboldt County, California. Egg masses are frequently covered in sediment ­after a few days, at which time they match the brown alga Nostoc that is pre­sent in the same habitats. The resemblance may help conceal the eggs. The male amplexes the female and then rides along with her as she selects an oviposition site. Females prepare oviposition sites by scraping a rock surface with their hind or front limbs to remove algae and debris, thus making the rock less slippery. This pro­cess normally lasts 50%; ­these effects result in an increase in the extent of Cladophora turfs (Kupferberg,

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1997c). By feeding on periphyton, R. boylii larvae actually increase area-­specific primary productivity by 10%, although biomass-­specific productivity decreases. As a result, the abundance of invertebrate consumers and their predators declines in areas grazed by Foothill Yellow-­legged Frog larvae. Rana boylii can experience resource competition from the introduced American Bullfrog (L. catesbeianus). In field experiments, bullfrog larvae selectively graze on the most nutritious algae (Cladophora with diatoms) and by ­doing so reduce survivorship and mass at metamorphosis of ­those sympatric R. boylii larvae that do survive (Kupferberg, 1997a). According to Kupferberg (1997b:155), “a small impact of bullfrogs on algal quality, thus resulted in a large impact on Rana boylii ­because ­there is such a strong correlation between algal quality and the biomass of R. boylii metamorphosing from the enclosures.” Thus, negative effects on native ranids can result by the way bullfrog larvae selectively graze on food resources. Large overwintering bullfrog larvae can also get a head start on newly hatched R. boylii larvae by outcompeting them physically for access to high quality food. DISEASES, PARASITES, AND MALFORMATIONS

Rana boylii is parasitized by the introduced Asian copepod Lernaea cyprinacea, outbreaks of which are correlated with unusually warm summers (Kupferberg et al., 2009). In California, outbreaks of this parasite ­were associated with increased levels of malformations (to 26.5% vs. 1.1% of uninfected larvae), particularly at infestation sites around the hind limbs and cloaca. Malformations included missing hands, missing limbs, fused digits, abnormal limb development, and extra limbs. Recent metamorphs ­were generally smaller if infested with copepods as larvae compared with uninfected froglets, and infestation rates ­were higher in downstream sites compared with upstream sites. The amphibian chytrid fungus Batrachochytrium dendro­ batidis has been found in R. boylii from California (Green et al., 2002; Ecoclub Amphibian Group et al., 2016; Adams et al., 2017a, b) and from natu­ral populations in the Pacific Northwest (Adams et al., 2010). Indeed, this fungus was found in museum specimens collected from California as early as 1966 (Padgett-­Flohr and Hopkins, 2009). It does not kill postmetamorphic Foothill Yellow-­legged Frogs outright, but it does substantially retard growth by as much as 50% (Davidson et al., 2007). Rana boylii has peptides in the skin that inhibit the growth of amphibian chytrid and likely prevent substantial mortality in adults. However, the effectiveness of the peptides is reduced when frogs are exposed to certain contaminants such as the pesticide carbaryl.

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Foothill Yellow-­legged Frogs are parasitized by the trematodes Clinostomum sp., Deropegus [Halipegus] aspina, Glypthelmins quieta, Gorgoderina multilobata, Haematoloe­ chus sp., H. kernensis, H. variplexus, Megalodiscus microph­ agus, and M. temperatus; Distoichometra bufonis and unidentified dilepinid cestodes; the nematodes Cosmocer­ coides variabilis, Falcaustra pretiosa, F. ranae, Hedruris sp., and Rhabdias ranae; and acanthocephalans (centrorhychid and oligocanthorhychid cystacanths) (Ingles and Langston, 1933; Ingles, 1936; Walton, 1941; Lehmann, 1960; Goldberg and Bursey, 2002a; Bursey et al., 2010). SUSCEPTIBILITY TO POTENTIAL STRESSORS

Metals. Mercury has been found in R. boylii from California at concentrations of 0.030–0.65 μg/g wet weight (Hothem et al., 2010). Values >0.3 μg/g wet weight exceed EPA standards for mercury concentrations in fish. Chemicals. Death occurs in about 18 hrs at 30 mg/L of carbaryl, a broad-­spectrum insecticide, and it significantly reduces larval activity levels at 2.5 mg/L (Bridges and Semlitsch, 2000). At low concentrations (0.48 mg/L), it has no effect on survival. The estimated minimal lethal concentration is 4.8 mg/L (Davidson et al., 2007). However, carbaryl, even at low doses, reduces the effectiveness of skin peptides in repelling amphibian chytrid fungus in postmetamorphs and thus has sublethal effects that might be overlooked in dosage experiments alone. The agricultural pesticides chlorpyrifos, diazinon, malathion, and their oxon derivatives are toxic to larval R. boylii at concentrations found in the environment. LC50 (1–4 day) values are 3.0 mg/L for chlorpyrifos, 2.14 mg/L for malathion, and 7.49 mg/L for diazinon. Oxon derivatives are 10–100 times more toxic than their parental forms (Sparling and Fellers, 2007). ­These pesticides depress normal cholinesterase activity. STATUS AND CONSERVATION

This species has dis­appeared from much of its historic range ( Jennings, 1988a; Jennings and Hayes, 1994a; Bury, 2008), and this decline occurred in 50) are large and distinct, and the dorsal skin ranges from smooth to rather rough in texture. Dorsolateral folds are prominent, and the jaw has a distinct light stripe. Venters are yellowish or buff colored, and melanophores are absent from the medial abdominal area. A diffuse, dark reticulation pattern is evident in the groin, which is yellow green. Most of the ventral yellow coloration is located posteriorly and on the lower limbs. Jennings and Hayes (1994b) noted that the iris is brown with gold iridophores. Males have prominent nuptial pads. Dunlap (1955) provided extensive information on the color pattern and morphology of this species. Females are slightly larger than males. In Oregon, males reach a maximum SUL of 57 mm whereas females reach a maximum SUL of 66 mm (Briggs and Storm, 1970). In California, males averaged 43.9 mm SUL (range 28–53 mm) and females 56.4 mm SUL (range 45–70 mm) based on museum specimens (Goldberg, 2020i); the smallest mature female was 48 mm SUL, whereas the smallest mature male was 40 mm SUL. Jennings and Hayes (1994b) give a maximum size of 75 mm SUL. Larvae. The tadpole is brown or olive brown dorsally and on the tail musculature. The belly is pale and has a golden iridescence, and the tail often has small blotches. Larvae reach a mean maximum size of 53–66 mm TL. Sype (1975) provides a description of the eggs and larvae from fertilization through metamorphosis. Albino tadpoles ­were reported by Altig and Brodie (1968) from Oregon, and Brome McCreary observed albino tadpoles and a recent metamorph (photos in this volume). Eggs. Eggs are black dorsally and cream colored ventrally. ­There are 3 distinct envelopes surrounding the vitellus. The outer envelope is ca. 11.3 mm; the ­middle envelope is

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ca. 5.8 mm in dia­meter; the inner envelope is ca. 4.9 mm in dia­meter; and the vitellus is ca. 2.3 mm (Livezey and Wright, 1947). Eggs are deposited in a solid jelly mass. DISTRIBUTION

The Cascades Frog is found in 3 disjunct regions: the Olympic Mountains of Washington, the Cascades Mountains of Washington and Oregon, and the Klamath-­Siskiyou Mountains in northern California. The species occurs at elevations of 230–2,740 m. Impor­tant distributional references include: Slater (1955), Metter (1960), Bury (1973), Hayes and Cliff (1982), Nussbaum et al. (1983), Leonard et al. (1993), Jennings and Hayes (1994b), McAllister (1995), Corkran and Thoms (2020), and Flaxington (2021). FOSSIL REC­O RD

No fossils are known. SYSTEMATICS AND GEOGRAPHIC VARIATION

­ here have been a number of studies that suggested someT times conflicting phylogenies among Pacific Northwest ranids (Zweifel, 1955; Dumas, 1966; Case, 1978; Farris et al., 1979, 1982; Green, 1986a, 1986b; Macey et al., 2001; Hillis and Wilcox, 2005). This species is a member of the R. muscosa complex (sometimes termed the Rana boylii species group; Macey et al., 2001) of the Amerana clade of North American ranid frogs. It is most closely related to R. aurora, and somewhat more distantly to R. muscosa and R. sierrae. Green

Distribution of Rana cascadae

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(1986a) described the chromosomes and hypothesized that R. cascadae was related more to R. aurora–­R. draytonii and R. pretiosa–­R. luteiventris than to R. boylii and R. muscosa. The diploid chromosome number is 26 (Haertel et al., 1974). Populations of R. cascadae show a high degree of ge­ne­tic differentiation at a local scale, and ­there is a discordance between the results of molecular analyses depending upon ­whether mtDNA or nuclear DNA is studied. mtDNA results suggest a 2–3 my separation between the Olympic and Cascades populations, but this is not supported by microsatellite data, which indicate a more recent separation. However, microsatellite results indicate a break between the Oregon and Washington Cascades populations at the Columbia River. Monsen and Blouin (2003) suggested that ­there ­were 2 distinct population segments within R. cascadae, Washington/ Oregon and California, but that the 3 disjunct populations should be managed for conservation separately. Laboratory hybridization experiments suggest ge­ne­tic incompatibility with R. draytonii (Zweifel, 1955). Hybridization occurs with R. pretiosa ­under laboratory conditions, with larvae completing metamorphosis (Haertel and Storm, 1970) or not (Dumas, 1966). Hybridization was more successful when R. cascadae was the female parent. Green (1985c) documented hybridization with R. pretiosa in nature, but the hybrids ­were infertile ­because of a lack of chromosome pairing during meiosis. ADULT HABITAT

The Cascades Frog is found in boreal habitats (especially associated with firs [Abies] and arborvitae [Thuja]) in the high Cascade and Olympic Mountains of the Pacific Northwest. At one time, however, they ­were also known from low elevation sites on the Olympic Peninsula (Leonard et al., 1993). Habitats include ponds, lakes, slow-­moving steams through wet meadows, meadow wetlands, and riparian areas along fast-­moving, steep mountain streams. Both permanent and ephemeral habitats may be occupied, but ephemeral habitats must have saturated areas for the species to survive. This species prefers old-­growth forest habitats but can be found in less abundance in mature second-­growth and young forest stands (Aubry and Hall, 1991). Bosakowski (1999) found most Cascades Frogs in open meadows surrounded by conifer saplings, and Cole and North (2014) found positive associations at high elevations with many (>30) pools, a lack of trout, wet meadows, and distances 5 km (maximum 5.75 km) over substantial (to 770 m) gains in elevation. Some frogs even managed to climb 36° slopes for 2 km. Such dispersal distances suggest substantial amounts of gene flow within mountain basins. In contrast, only 4% of marked adults moved >200 m, although 4 frogs moved 2 km. In another example, from 1 to 32% of marked juveniles moved between breeding ponds and summer habitats in Idaho (Pilliod et al., 2002). When juveniles stop dispersing, they tend to remain in the vicinity of the newly ­adopted ponds rather than return to their natal pond. ­There is a ­great deal of variation in landscape ecol­ogy from one season to the next and among individual frogs. Some frogs remain within the same areas virtually all their lives and repeatedly use the same breeding and overwintering sites. ­Others remain in an area for 1–2 yrs before moving to a dif­fer­ent area, and still other frogs roam at considerable distances across the landscape. Turner (1960b) recaptured frogs as far as 610 m from the point of original capture. Indeed, the activity range of R. luteiventris in Yellowstone varied between 230 and 3,312 m2 and was not related to the sex or age of the frog. Breeding ponds do tend to be isolated in some areas, with the extent of ge­ne­tic isolation a function of distance between breeding ponds, regardless of ­whether distance is mea­sured linearly or along drainages (Goldberg and Waits, 2010). Goldberg and Waits (2010) suggested that urban and rural developed lands ­were most responsible for ge­ne­tic isolation, and that agricultural lands and grasslands provided the least re­sis­tance to isolation in northern Idaho. CALLING ACTIVITY AND MATE SE­L ECTION

Males arrive at breeding sites 1–36 days (median 6.5 days; Davis and Verrell, 2005) prior to females, and they establish the oviposition site. Calling occurs by day or at night depending upon population. The call of this species sounds like repeatedly clicking one’s tongue against the roof of the mouth (Morris and Tanner, 1969). ­There are from 4 to 50 notes per call, with the rate of note production directly correlated with temperature; the warmer the temperature, the more notes are produced. Males can produce 300–480 notes per minute, and calls last from 4 to 10 sec. Calls are produced

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both in air as the frog sits in the shallow ­water along a pond’s edge or while the male is submerged. Calls are not very loud (in contrast to descriptions of deep resonant calls by Svihla, 1935), and a ­human observer can only hear a call 14°C along the Wasatch Front in Utah (Morris and Tanner, 1969). Tadpoles metamorphose prior to midsummer, when ­water heats up to lethal temperatures. As with most frogs, breeding does not occur at ­every potential breeding site. For example, breeding occurs in

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14–26% of potential breeding sites examined in the Greater Yellowstone Ecosystem (Patla, 1997; Corn et al., 2005; Corn, 2007; Gould et al., 2019). In northern Idaho, Lucid et al. (2020) found frogs at 204 of 433 (47%) potential sites. The reasons for this are complex, involving the interrelationship of hydroperiod, ­water depth, vegetation, evapotranspiration, and current environmental conditions (i.e., drought versus wet years) (Gould et al., 2019). For example, conditional breeding probabilities of R. luteiventris are associated positively with runoff and maximum ­water depth, as is breeding per­sis­tence. However, deep wetlands had lower probabilities of breeding per­sis­tence than shallow wetlands with low runoff. With high runoff, breeding per­sis­tence was higher in deeper wetlands. As such, ­whether a site ­will be used for breeding in any par­tic­u­lar year is the result of a myriad of ­factors. REPRODUCTION

Male Columbia Spotted Frogs have sperm in their testis in June and July, and females have mature oocytes from July to September (Goldberg, 2019g). As such, spermiogenesis occurs in spring and early summer, and females likely carry mature ova through the winter, allowing spawning on emergence in spring. Goldberg (2019g) found the smallest mature male to be 42 mm SUL, whereas the smallest mature female in spawning condition was 55 mm SUL. Additional information on spermiogenesis and oogenesis is found in this paper. Breeding occurs over a wide range of dates, depending upon location, temperature, and elevation. For example, breeding occurs from mid-­March to early April in Utah (Hovingh, 1993b; Cuellar, 1994) and the Palouse Region of Idaho and Washington (Davis and Verrell, 2005), between late March and late May in northeastern Oregon and eastern Washington (Svihla, 1935; Metter, 1960; Bull and

Egg mass of Rana luteiventris. Photo: Brome McCreary

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Hayes, 2001), and in May in Alaska (Ream et al., 2019). Breeding normally occurs in mid-­to late May and June in Yellowstone National Park (although adults have been observed in April: Yeager, 1926 [as “northern frogs”]; Turner, 1955, 1958), and into early July in interior British Columbia (Logier, 1932). Breeding dates also change with elevation. For example, breeding occurs from late March to May at low elevations in Montana, but in June at higher elevations (Black, 1970; Werner et al., 2004). Similar elevation-­based trends are reported in Alberta, with breeding occurring from early May to June (Salt, 1979). Even within a localized area, frogs ­will breed ­earlier in warm-­water spring habitats than in ponds fed by cold-­ water springs. Breeding also can be accelerated by warm weather or delayed or interrupted by cold weather. During cold weather, an amplexed pair ­will move to deep ­water ­until temperatures rise again. Males enter breeding ponds prior to females, and females are ready to ovulate as soon as they enter the breeding ponds. Eggs are deposited ­either day or night in a jelly mass 10– 20 cm below the ­water’s surface attached to floating or emergent vegetation. Masses mea­sure 75–200 mm in dia­meter. Within a short time, the attachments give way and the eggs tend to flatten out and float at the surface. Some of the eggs are attached to one another by a small gelatinous cord, and a single egg may be connected to 5 other eggs through the cord or outer membranes. This species has communal oviposition sites, and the egg masses tend to be in close proximity to one another. For example, Morris and Tanner (1969) recorded 50 egg masses within an area 75 cm in dia­meter, and Bull and Marx (2002) noted up to 135 egg masses per oviposition site. Koch and Peterson (1995) observed communal sites with 34 and 45 egg masses, whereas Hossack et al. (2013c) counted from 1 to 57 egg masses per site at 26 of 60 wetlands sampled. Only a few clutches may be oviposited away from the larger mass of eggs, but ­there may be multiple communal oviposition sites per lake. Each mass normally contains approximately 325–950 eggs and mea­sures ca. 20 cm × 15 cm. Specific counts include a mean clutch size of 605 eggs (range 430–725) one year in Utah and 746 (range 147–1,160) the next (Morris and Tanner, 1969); 325–710 (mean 444) eggs per clutch in western Utah (Cuellar, 1994); a mean of 539 eggs per clutch (range 206–802) in Wyoming (Turner, 1958b); a range of 150–2,000 in Colorado, with most masses containing 500–600 eggs (Livo, 1998); 2 masses of 1,000 and 1,500 eggs in eastern Washington (Svihla, 1935); and a maximum clutch size of 2,400 (Livezey and Wright, 1947). Algae may enter the egg capsules, giving them a greenish appearance as development progresses. Egg mortality results chiefly from freezing and desiccation, although embryos within the jelly

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mass may survive for some time out of ­water (Hossack, 2006). Presumably, such embryos would have the chance to survive short-­term desiccation ­until reinundated. Hatching occurs in 12–21 days, with metamorphosis following in about 60 days of further aquatic development (Turner, 1958b), although Black (1970) stated hatching occurs in only 4 days. In interior British Columbia, Logier (1932) suggested that larvae overwinter and do not metamorphose ­until the following early summer. Hatching and development rates are directly tied to ­water temperature, with ­later hatching and slower growth at low temperatures. At hatching, larvae are 7–10 mm TL. Columbia Spotted Frog females may not produce a clutch of eggs ­every year, and at most they produce only 1 egg mass per female. In Yellowstone National Park, for example, successful egg deposition takes place only once ­every 2–3 yrs, although males breed ­every year (Turner, 1958b). The number of egg masses at a location also varies annually. For example, Hovingh (1993b) reported from 2 to 33 clutches at a site in Tule Valley, Utah, although at other sites as many as 462 masses ­were counted at a single location in a single year. In northeastern Oregon, Bull and Hayes (2001) counted from 3 to 39 egg masses at 6 breeding ponds. In British Columbia, ­there ­were only a handful to nearly 90 egg masses per site at 24–25 ponds over a 2 yr period; the largest number was at a single spring-­fed pond (Swan et al., 2015). ­Whether ­these values reflect differences in resource availability, environmental conditions, or fluctuations in breeding female population size is unknown. Swan et al. (2015) noted that the number of egg masses was positively correlated with pond elevation, mean temperature, mean pH, and the presence of fish in a pond in the drawdown zone of the Kinbasket Reservoir in British Columbia. LARVAL ECOLOGY

Larvae feed on decomposing plant material and green algae. They also may be able to derive nutrition from bacteria in the ­water (Burke, 1933), and they ­will feed on carrion such as dead conspecifics during the ­later stages of development. Tadpoles form aggregations that function to stir up food from the substrate or to conserve heat and thus speed up development (Carpenter, 1953b). ­These aggregations may produce audible clicking sounds as the tadpoles smack their lips and rise to the surface to gulp air. Larvae can be quite abundant in optimal sites. Salt (1979) reported densities of 150–225 larvae/ m2 at sites in Alberta. At another site, he noted from 10 to 60 larvae in a single square meter. Carpenter (1953b) recorded from 1,000 to 1,500 tadpoles within a 30 cm × 30 cm area. The normal lethal temperature limit of larval development is 28°C (Licht, 1971), although Brues (1932) reported larvae of this species “swimming about in ­water ranging from 39.2 to 41.6°C” in Yellowstone National Park. The CTmax for

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larvae is 37.9°C (Katzenberger et al., 2018). Thermal analyses suggest that microclimate temperatures ­will be more impor­tant than mean air temperatures in this species’ response to climate change. ­There is considerable variation in the maximum size of tadpoles. Morris and Tanner (1969) reported that larvae normally reach 50–55 mm TL just prior to metamorphosis, but that some tadpoles reached 70 mm TL. Turner (1958b) recorded the maximum size as 60 mm TL. Recent metamorphs, however, are 16–25 mm TL (Turner, 1958b; Livo, 1998; Russell and Bauer, 2000). Likewise, the larval period is quite variable, ranging from 70 to 85 days in Wyoming (Turner, 1958b) and from 122 to 209 days in Utah (Morris and Tanner, 1969). Most authorities state that tadpoles do not overwinter. Still, large tadpoles have been found in January in Montana (Black, 1970). DIET

This species consumes a variety of invertebrates, particularly snails, Gammarus shrimp (or scuds), sow bugs, spiders, beetles (both aquatic and terrestrial), moths, ­water striders, ants, lepidopteran larvae, and many types of flies and fly larvae (Ruthven and Gaige, 1915; Tanner, 1931; Moore and Strickland, 1955; Turner, 1959b; Miller, 1978). The type of prey may change seasonally, depending on availability. Plant debris and inert detritus may be consumed as prey are captured and forced into the mouth. No diet specialization is evident, and frogs likely consume what­ever small invertebrates come within feeding range. Feeding occurs mostly during the day and at dusk. Although small frogs naturally consume small prey, large frogs eat a wide variety of different-­sized prey. Debris may be ingested in conjunction with feeding, and Turner (1960b) recorded an instance of a large stone becoming lodged in the intestine of a R. luteiventris, which undoubtedly would have caused its death. PREDATION AND DEFENSE

When disturbed in ­water, Columbia Spotted Frogs dive into the substrate and bury themselves in mud. If on land, Black (1970) reported they might attempt escape by jumping down mouse burrows. Likely predators of postmetamorphic Columbia Spotted Frogs include mammals (Coyotes, Otters, Mink), birds (Ravens, Northern Harriers, Herons, Egrets, Sandhill Cranes, ­Great Gray Owls, Pygmy Owls, Ring-­billed and California Gulls), garter snakes (Thamnophis sp., T. elegans), and trout (Salt, 1979; Koch and Peterson, 1995; Drost, 2020). Eggs masses are attacked by predaceous diving beetles, crane flies, and leeches (Haemopis marmorata) (Hovingh, 1993b). Larvae are prob­ably consumed by predaceous invertebrates (dytiscid beetles), a variety of birds including Gray Jays, and

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garter snakes (Turner, 1960b; Black, 1970). Hatchlings may be eaten by conspecifics (Rombough, 2016). POPULATION BIOLOGY

Age at sexual maturity is variable depending upon environmental conditions and elevation. For example, sexual maturity is delayed in this species in Yellowstone National Park, with males breeding first at about at 4 yrs (at >45 mm SUL) and females at 5–6 yrs (at >60 mm SUL). In coastal populations, however, maturity of both sexes is reached at 2 yrs. Longevity is approximately 10 yrs for males and 12–13 yrs for females in natu­ral populations at Yellowstone (Turner, 1960b), but only 3–4 yrs in coastal populations and 5 yrs for males and 7 yrs for females in the Nevada ­Great Basin (Reaser, 2000). The rigors of Yellowstone and other interior regions thus have major consequences on the demography and longevity of Columbia Spotted Frogs. Young Columbia Spotted Frogs in Yellowstone grow rapidly ­after metamorphosis and are generally 25–30 mm SUL when they emerge from overwintering the following year. If newly metamorphosed frogs are ca.16 mm SUL, then this indicates an initial growth spurt of 9–14 mm (Turner, 1960b). By the end of the first year, juveniles ­will have grown 19% of their total body size, and they ­will have grown 52% by the end of the second full year ­after transformation (Turner, 1960c). In Yellowstone, they do not reach 50 mm SUL ­until their fourth or fifth full season ­after metamorphosis. Turner (1960b) provided growth curves for his Yellowstone population and speculated that differences in male and female growth rates first become apparent in their fourth year. Some populations of R. luteiventris grow much faster than ­these rates, perhaps reflecting differences in growing season, resource availability, and environmental conditions among populations. Although a variety of aquatic sites are used by this species for reproduction, McCaffrey et al. (2014) found that survivorship from eggs to metamorphosis was highest in semiperma­ ere oviposited in nent ponds despite the fact that most eggs w permanent ponds. Larval recruitment into the population as metamorphs varied annually from one pond type (ephemeral, semipermanent, permanent) to another. In warmer years, permanent ponds produced the most metamorphs, but in cooler years, the only successful metamorphs came from semipermanent and ephemeral ponds. Overall net recruitment and stochastic population growth rates ­were highest in semipermanent ponds with homogenous hydroperiods, but lowest when pond hydroperiods ­were heterogenous, regardless of pond type (McCaffrey et al., 2014). Postmetamorphic survivorship can be relatively high. Funk et al. (2005b) estimated a juvenile survival rate of 33%, indicating that ­those juveniles that survived dispersal had a relatively good chance of establishing themselves at the

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Adult Rana luteiventris. Photo: Dirk Stevenson

ponds to which they immigrated. It is thus not surprising that populations of Columbia Spotted Frogs historically could be substantial. In the early 1950s, for example, Turner (1960b) estimated ­there ­were 1,200–1,850 frogs occupying his 28 ha Yellowstone study site. In western Utah, Cuellar (1994) estimated that ­there ­were 100 frogs/ha. However, breeding populations are often small. The sex ratio of frogs in a population can vary depending on age-­class (Reaser, 2000). Since males remain at breeding sites longer than females and call from communal oviposition sites, sex ratios at breeding ponds might be expected to be male biased (Davis and Verrell, 2005). For example, Morris and Tanner (1969) reported 5 males per ­every female at their Utah breeding sites. In the Palouse Region, males sometimes exceeded females by 11 to 1 at small breeding ponds (Davis and Verrell, 2005). In contrast, Reaser (2000) reported female-­biased sex ratios at most Nevada ­Great Basin sites she studied, and found that that sex ratios varied annually among sites. Likewise in Turner’s (1960b) study, the overall sex ratio was female biased (1:2). Sex ratios ­were 1:1 ­until the fifth year, when highly skewed female-­biased sex ratios ­were observed. ­These observations, coupled with Reaser’s (2000) analy­sis of lines of arrested growth, suggest that females live longer than males, perhaps ­because of differential survivorship. In general, however, overall annual survivorship is high for adults. In the mountainous West, landscape features play impor­ tant roles in the ge­ne­tic structuring of populations of Columbia Spotted Frogs. In an extensive analy­sis of 6 microsatellite loci, Funk et al. (2005a) demonstrated that mountain ridges and elevation act as ge­ne­tic barriers and thus increase ge­ne­tic differentiation among sites. Most populations consist of multiple ponds, except in the case of isolated wetlands, and ge­ne­tic relatedness is more evident within

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mountain basins than between them. Gene flow also occurs more extensively among low-­elevation populations than among ­those at high elevations; ge­ne­tic variation among populations is strongly negatively correlated with elevation. Low-­elevation populations are impor­tant sources of immigration and ge­ne­tic variation. If ­these populations are eliminated, their loss could affect the per­sis­tence of high-­elevation populations, even without direct habitat threats. In the mountainous West, connectivity among populations is an impor­tant ­factor as to occupancy (Murphy et al., 2010a; Gould et al., 2012). Columbia Spotted Frogs do not occupy all available habitats within a region. For example, Goldberg and Waits (2009) recorded R. luteiventris in 31% of 105 wetlands surveyed on private lands in Idaho, and predictive models suggested that breeding should occur in only 8–15% of wetlands in their survey area. In the Salmon River Mountains of Idaho, connectivity among nutrient-­ poor high mountain lakes is negatively correlated with distance between sites, the presence of predatory fish, and the topographic complexity between sites; site productivity (based on a heat load index) and growing season (in terms of frost-­free periods between sites) ­were positively correlated with gene flow (Murphy et al., 2010a). Together, ­these results suggest that Columbia Spotted Frog populations function in a dynamic source-­sink population structure in this region. The abundance of this species throughout its range is not uniform among populations representing the 3 main evolutionary lineages. In analyses of long-­term data, Hossack et al. (2013d) showed ­there was significant variation in growth that was associated with landscape characteristics, drought frequency, and vari­ous management protocols, particularly the construction of breeding wetlands. Egg mass abundance declined by 3% annually from 1991 to 2011 across the range. Despite this, egg mass abundance was estimated to have increased by 32% in the ­Great Basin clade and 47% in the Utah clade. In the Northern clade, egg mass abundance declined by 12%. Population growth estimates (based on egg mass counts) ­were negative for 7 of 19 sites within the Utah clade, 5 of 8 sites in the ­Great Basin clade, and 40 of 65 sites in the Northern clade. Small, temporary breeding sites ­were most susceptible to drought, whereas even a few large wetlands (natu­ral or restored) buffered the effects of drought throughout a region (Hossack et al., 2013d). Locally, ­there was a strong negative impact of ­human disturbance on growth (egg mass counts) within populations. Columbia Spotted Frogs are easily observed when pre­sent during monitoring programs. During surveys of potential habitat, for example, detection probabilities ­were generally high in Glacier and Yellowstone National Parks (75–95%), and adjusted occupancy rates ranged from 17 to 26% of habitats examined over a 2 yr period (Corn et al, 2005).

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DISEASES, PARASITES, AND MALFORMATIONS

Peptides in the skin may help in defense against bacterial and Bd infections (Goraya et al., 2000; Loudon et al., 2020). The amphibian fungal pathogen Batrachochytrium dendrobatidis (Bd) has been detected in this species in the Greater Yellowstone Ecosystem (Corn, 2007), Oregon (Adams et al., 2010), and elsewhere in British Columbia (Brunet et al., 2020), Idaho, Montana, Utah, Washington, and Wyoming (Muths et al., 2008; Russell et al., 2010; Gaulke et al., 2011; Araos et al., 2017; Erdmann et al., 2018); ­these latter authors found the pathogen in 35–40% of the individuals sampled. In Oregon, 12 of 14 sites had positive indications of Bd, and 71 of 198 (35.9%) frogs tested ­were positive (Adams et al., 2010); 1 larva also tested positive. Ranavirus has been detected in the Greater Yellowstone Ecosystem (Patla et al., 2016) and in northern Idaho (Russell et al., 2011). Rhizobacter and Chryseobacterium are the main bacteria found on this species (Loudon et al., 2020). Rhizobacter relative abundance is negatively correlated with the ability of antimicrobial peptides to inhibit Bd, but is not associated with Bd status. ­There is no relationship between metabolites and Bd. Bacterial communities and Bd differ by location on the frog, which suggested to Loudon et al. (2020) a strong environmental influence. Rana luteiventris are inhabited by consistent core bacterial communities, but frogs also ­house transient bacteria that are site specific. Columbia Spotted Frogs are parasitized by the nematodes Aplectana gigantica, Gyrinicola batrachiensis, and Spi­ ronoura pretiosa and by the trematodes Glypthelmins sp., Gorgoderina tanneri, G. translucida, Haematoloechus parviplexus, H. similiplexus, Halipegus sp., Haplometrana intestinalis, and H. utahensis (Olsen, 1937, 1938; Turner, 1958a; Waitz, 1961; Hossack et al., 2013c). Males and females are subject to equal parasitic loads. Hossack et al. (2013c) found 66% of the 205 larvae they examined ­were parasitized by G. batrachiensis, with from 1 to 107 worms per larva. Leeches may attach themselves to the gular region of tadpoles (Carpenter, 1953b). Malformations of R. luteiventris have been reported at frequencies of 21–31% at certain sites in the Pacific Northwest, with from 82 to 144 Ribeiroia metacercaria per frog ( Johnson et al., 2002; Roberts and Dickinson, 2012). Malformations included cutaneous skin fusions (extra webbing), missing digits (ectrodactyly), polydactyly, ectromelia (missing limb), partial limbs (hemimelia), femoral projections, limb rotation, and other miscellaneous deformities. SUSCEPTIBILITY TO POTENTIAL STRESSORS

Chemicals. The herbicide Roundup® is toxic to larvae, with an LC50 (24 hrs) of 1.65 mg/L (King and Wagner, 2010). The LC50 decreases at 15 days to 0.98 mg/L.

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Metals. The presence of heavy metals, ­either singly or in combination, may alter the ability of Columbia Spotted Frog larvae to respond to predators. When taken in combination, sublethal effects of metals are more evident at lower dosages than they are singly. For example, combinations of cadmium and zinc are much more toxic to larvae than ­either metal is by itself. Medium levels of zinc and medium to high levels of lead decrease the fright response of tadpoles, and exposure to soils containing metals may delay metamorphosis and decrease larval fright responses (Lefcort et al., 1998). The LC50 (4 day) of specific metals is: zinc (28.4 ppm), cadmium (15.8 ppm), and zinc + cadmium (4.44–4.52 ppm). ­These results suggest that sites contaminated by a variety of metals could be both toxic and have sublethal effects on tadpole survival. In a further experiment, Lefcort et al. (1999) showed that in the presence of lead-­, zinc-­, and cadmium-­contaminated soil, tadpoles metamorphosed at an older age than non-­metal-­exposed animals. In the absence of heavy metals, tadpoles reduced snail (a potential competitor) recruitment. ­Because heavy metals disproportionately harmed tadpoles, the negative effects of tadpoles on snails ­were ameliorated in polluted environments. The presence of snails did not alter tadpole development but did increase metal loads in tadpoles. UV light. Columbia Spotted Frogs have high levels of photolyase, an enzyme impor­tant in DNA repair from UVB radiation. Spotted frogs reared in ambient UVB and in situations where UVB is shielded have similar levels of hatching success and embryo development (Blaustein et al., 1999). STATUS AND CONSERVATION

Columbia Spotted Frogs are susceptible to habitat loss, as are all frogs. Populations have dis­appeared from many historic locations, such as along the Wasatch Mountain front in the Bonneville Basin of Utah, the northern ­Great Basin, and in southwestern Alberta (Weller and Green, 1997; Corn, 2000; Wente et al., 2005). Declines are evident even in some protected areas, such as the Greater Yellowstone Ecosystem, but not in ­others, for example Glacier National Park (Hossack et al., 2015). Wente et al. (2005) estimated that R. luteiventris occupied only 53% of its historically documented locations in northern Nevada and eastern Oregon. Suggested ­causes include the outright destruction of habitats, habitat fragmentation, ­water modification proj­ects (channelization, diversion, irrigation, floodplain alteration), and the introduction of nonnative species (crayfish, fish, American Bullfrogs, raccoons) (Hovingh, 1993b). Other human-­related mortality occurs from roadkill, use as bait, malicious killing, and trampling by human-­introduced livestock (Turner, 1960b; Ross et al., 1999). ­There is ­little doubt that the introduction of nonnative salmonid fishes into high-­elevation lakes and streams has

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Habitat of Rana luteiventris. Utah. Photo: Breck Bartholomew

adversely affected Columbia Spotted Frog populations, even in remote wilderness (Pilliod and Peterson, 2000, 2001). Fish come to occupy nearly all frog habitats (­those >1 ha and >4 m deep), and many fish-­uninhabited sites are too small, shallow (800 m from its release pond, although most frogs dispersed over much shorter distances (50–510 m, mean 80 m). Whereas most interpond movement likely occurs along watercourses, ­these frogs are capable of making straight-­line movements over inhospitable terrain of several hundred meters or more. Seasonal migrations occur between summer breeding and feeding sites and sites where frogs spend the winter. At high

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690  RANIDAE

elevations, Mountain Yellow-­legged Frogs tended to remain within a lake or adjacent stream, with mean movements of 77 m over the course of August (Matthews and Pope, 1999). In September, movements increased, with mean straight-­line movements of 145 m and circuitous movements of 315– 466 m. ­These autumnal movements decreased substantially (to a mean of 43 m) by October, as frogs took up winter residences. Accordingly, home range estimates ­were 385 m2 in August, 5,336 m2 in September, and 53 m2 in October (Matthews and Pope, 1999). Temperatures during autumn movements can be quite cool (5.5–12.5°C). Movements generally follow watercourses, but frogs occasionally move overland as much as 66 m between wetland sites (Matthews and Pope, 1999). Overwintering by larvae and postmetamorphs occurs in permanent lakes and streams in the high mountains, where activity may occur only during a brief 3–5 month summer season. ­These sites may be several hundred meters from the summer feeding areas and are usually deep and of large surface area. Matthews and Pope (1999) tracked frogs to nearshore habitats ­under ledges and in deep ­water crevices in fractured bedrock. The crevices ­were 0.2–1.2 m below the ­water surface in Matthews and Pope’s (1999) study. Some of the high-­elevation lakes and streams may be covered by ice 6–9 months of the year, and if winters are severe with oxygen depletion, nearly all Mountain Yellow-­legged Frogs ­will die.

Eggs of Rana muscosa. Note how silt attaches to the egg membranes. Photo: Adam Backlin

The calls are comprised of a series of raspy scraping sounds, often ending in a loud, accentuated note (Lang et al., 2009). Rana muscosa also makes a call with a stuttered note. Calls appear to require a ­great deal of effort. The calls of R. mus­ cosa and R. sierrae have dif­fer­ent properties and are easily distinguished using oscillogram and spectrogram analyses (Vredenburg et al., 2007). Rana muscosa, for example, does not have transitions between pulsed and noted sounds whereas R. sierrae does.

likely extends from early April to mid-­May. For example, Storer (1925) reported finding a pair in amplexus in early April; Zweifel (1955) found evidence of breeding from late April to early May; and Stebbins (1951) reported egg deposition in early to mid-­May. In the Sierra Nevada, it seems likely that reproduction occurs much ­later, from mid-­May to early July, depending upon elevation. At high elevations, the frogs must wait ­until ice and snow melts from the breeding sites. Santana et al. (2015) studied the effect of dormancy on captive R. muscosa. They found that dormancy resulted in positive effects on reproductive be­hav­ior, particularly vocal advertisements, female receptivity, amplexus, and oviposition. Results confirmed that a period of dormancy is necessary for successful reproduction, since only previously dormant females produced eggs and only previously dormant males successfully fertilized eggs. Body conditions ­were slightly less in previously dormant frogs, but no mortality resulted.

BREEDING SITES

LARVAL ECOLOGY

Eggs are deposited along streams and in higher lakes, tarns, and meadow ponds. In streams, eggs may be attached to rocks on the stream bottom, whereas in lakes they may not be attached and are allowed to float ­free. Eggs also may be oviposited in lentic pools in close proximity to lakes, streams, or meadow wetlands. The species tends to avoid the smallest streams.

Larval Mountain Yellow-­legged Frogs prefer relatively high ­water temperatures at breeding sites in the high mountains and actively seek warm (to 27°C) microhabitats in shallow ­water during the day. At night, they ­will move to deeper ­water as the shallows become cool. Larvae may form aggregations similar to ­those of R. cascadae and appear to follow much the same thermal pattern of diel habitat use as that species ( Jennings and Hayes, 1994b). Larval R. muscosa overwinter in streams and lakes at high-­elevation sites for 1 or 2 winters prior to metamorphosis. During severe winters, nearly all larvae die due to oxygen depletion when ­waters

CALLING ACTIVITY AND MATE SE­L ECTION

REPRODUCTION

­ ittle information is available on reproduction in this L species. In the southern part of its range, the breeding season

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Rana muscosa 691

POPULATION BIOLOGY

Nothing is known concerning the population biology of this species in the Transverse Ranges. At high elevations, it likely has similar population and demographic characteristics as other high-­elevation ranids such as R. cascadae and R. sierrae. COMMUNITY ECOLOGY

Tadpole of Rana muscosa. Photo: Chris Brown

freeze. However, some larvae can survive nearly anoxic conditions. Larvae have a lower critical oxygen tension than postmetamorphs, allowing them a greater tolerance of low PO2. In addition, they are able to reduce energy and oxygen consumption at low PO2 (Bradford, 1983). Despite their diurnal preference for higher ­water temperatures, larvae thrive in cold ­water, and Brattstrom (1963) recorded them in ­water at temperatures of 4–9°C. In the Transverse Ranges, metamorphosis usually occurs in the same season as oviposition, although Zweifel (1955) reported 1 observation that suggested at least some tadpoles overwinter. Overwintering usually would not be advantageous in the mountain streams ­because of the potential for scour flooding during the winter wet season. Recently transformed individuals are 20–30 mm SUL. DIET

Adults are known to consume large quantities of anuran tadpoles, particularly Anaxyrus canorus and Hyliola regilla (Pope, 1999; Pope and Matthews, 2002). Other prey includes spiders, ticks, harvestmen, beetles, flies, ants, bees, wasps, ­water striders, and aquatic Hemiptera (true bugs) (Long, 1970). Lemm (2006) stated that cannibalism had been reported, but this observation was likely in reference to R. sierrae. Males and females have similar diets, and based on gut analyses, some feeding occurs year-­round. Prey size is proportional to frog size, and prey items are selected opportunistically.

­ ecause of the short growing season, adult R. muscosa must B feed rapidly in order to grow and reproduce during the 3–5 month activity season. Pope and Matthews (2002) noted a correlation between the distribution of larval Hyliola regilla and Anaxyrus canorus in Kings Canyon National Park and suggested that an abundance of anuran larvae was essential to the growth and good body condition of R. muscosa. Adult R. muscosa may seek out ­water bodies with anuran larvae in order to take advantage of this nutritious food source in an other­wise food-­limited environment. DISEASES, PARASITES, AND MALFORMATIONS

The virulent pathogen Batrachochytrium dendrobatidis (Bd) was reported from R. mucosa tadpoles, metamorphs, and postmetamorphs in the Sierra Nevada as early as 1998 (Fellers et al., 2001a; Rachowicz et al., 2006; Woodhams et al., 2007; Smith et al., 2017). Backlin et al. (2013) noted its presence at all 9 known populations in southern California. Green et al. (2002) also reported this pathogen in R. muscosa but without locality data, although likely referring to the Fellers et al. (2001a) study. Russell et al. (2019) found that only 9% of R. muscosa they examined had Bd, with a survival rate of ca. 70–75%; males had lower survivorship than females. Russell et al. (2019) showed that the presence of the pathogen, even without obvious mortality, leads to reduced survivorship in wild populations. As noted below, Bd was likely responsible for the mass mortality observed at Kings Canyon National Park instead of

PREDATION AND DEFENSE

When disturbed, R. muscosa dive into the w ­ ater and submerge their bodies in the muddy substrate. As the frog buries itself, silt is stirred up, making it difficult to locate the well-­camouflaged frog. Potential predators include mammals (black bears, coyotes, ­humans), birds (Brewer’s Blackbirds, Ravens, Eared Grebes, American Dippers, American Kestrels), snakes (Thamnophis elegans), and lizards (Elgaria coerulea) on adults, subadults, and larvae (Fellers et al., 2007a). In experiments, trout rapidly consume both newly hatched and larger tadpoles (Vredenburg, 2004).

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Adult Rana muscosa. Photo: Chris Brown

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692  RANIDAE

the bacterial pathogen Aeromonas hydrophila, although this species plus the bacteria Enterobacter aerogenes and E. agglomerans ­were isolated from dead individuals (Bradford, 1991). Ranavirus also has been reported from Mountain Yellow-­legged Frogs, but Smith et al. (2017) suggested that it has not contributed to population declines in the Sierras. Endoparasites include the trematode Langeronia brenesi (Goodman, 1989). SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. Tadpoles do not occur in lakes with a pH ≤5.2, although adults inhabited lakes with a pH as low as 5.0 (Bradford et al., 1998). Embryos are much more sensitive to low pH than adults. Chemicals. The pesticides DDE, γ-­chlordane, trans-­ nonachlor, chlorpyrifos, and diazinon have been found in frog tissues from the high Sierras in areas that could have received contaminants borne on the wind from the San Joaquin Valley (Fellers et al., 2004, 2007a). Healthy frog populations in a wind-­protected site did not have high concentrations of pesticides, although ­these frogs may have been the closely related R. sierrae. It is uncertain ­whether organophosphate pesticides, which do not ­favor bioaccumulation, have played a role in frog declines in the high Sierras. UVB radiation. UVB radiation does not affect hatching success or developmental rates of embryos (Vredenburg et al., 2010a). STATUS AND CONSERVATION

The Mountain Yellow-­legged Frog has dis­appeared from most of its historic distribution in the Transverse Ranges and disjunct populations in southern California (Backlin et al., 2013). In addition, the species has declined in the southern Sierra Nevada, including habitat-­protected sites in Kings Canyon and Sequoia National Parks, where more than half of the historic populations have dis­appeared (Bradford, 1991; Bradford et al., 1994b; Jennings and Hayes, 1994a, 1994b; Vredenburg et al., 2005; Brown et al., 2014a; Rothstein et al., 2020). Even when pre­sent, Brown et al. (2014a) found few individuals in watersheds. Remaining populations are characterized by low ge­ne­tic diversity, evidence of ge­ne­tic bottlenecks, and a high degree of historical isolation with ­little gene flow (Schoville et al., 2011). The presence of nonnative predaceous fish in alpine lakes is correlated with an absence of R. muscosa tadpoles (Bradford, 1989; Bradford et al., 1998) and has fragmented remaining populations (Bradford et al., 1993). When fish are removed, frog populations recover rapidly (Vredenburg, 2004). Schoville et al. (2011) noted that the high degree of isolation ­will require individual management protocols.

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Habitat of Rana muscosa. Photo: Chris Brown

Batrachochytrium dendrobatidis (Bd) has decimated amphibians throughout the Western states, including R. muscosa, and may be the primary cause of population declines in this species (Bradford et al., 2011). The first documentation of its effects on R. muscosa was by Bradford (1991), who noted the extinction of this species at a site in Kings Canyon National Park. He observed that tadpoles and recent metamorphs had symptoms akin to the bacterial pathogen Aeromonas hydrophila, but the mortality event was more likely due to Bd. By 2005, Mountain Yellow-­ legged Frogs in Kings Canyon National Park ­were nearly all infected (96.7%) with Bd. Rachowicz et al. (2006) ­later found that 19% of 144 R. muscosa populations in the southern Sierra Nevada ­were infected during both years of their survey (2003–2004), with another 16% of the populations uninfected the first year but infected the second year. Following an outbreak, population size could be reduced as much as 88%. Despite the lethality of the fungus in labora-

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Rana pretiosa 693

tory and field settings, however, it is clear that at least some frogs survived and reproduced to allow the per­sis­tence of infected frog populations over a considerable period (Briggs et al., 2005). Infected frogs that survive appear to have low fungal loads, survive between years, and may even lose and regain the infection (Briggs et al., 2010). Survivorship within infected frog populations may result from density-­dependent host-­pathogen dynamics. This is especially true with regard to the pathogen’s life history characteristics, which allow for optimal encysting and the rapid production of zoospores at temperatures of 17–25ºC within a host’s cell wall. As temperatures drop to 7–10ºC, greater numbers of zoospores are produced which remain infectious for a long time (Woodhams et al., 2008). In effect, the pathogen responds to decreasing temperatures with increased fecundity as maturation rate slows, and infectivity increases as growth decreases (Woodhams et al., 2008). ­These results help explain why Bd is successful at colder temperatures. The fungus also is likely aided in its per­sis­tence within frog populations by its infection of the long-­lived tadpole and by nonhost fungal reservoirs (Briggs et al., 2005). Secretions from R. muscosa possess peptide mixtures which inhibit the growth of the pathogen (Woodhams et al., 2007). The peptides are produced by a variety of symbiotic bacteria and may be impor­tant in assisting disease re­sis­tance and defense in at least a few frogs. Still, Bd is likely the proximate cause of Mountain Yellow-­legged Frog disappearance throughout much of its former range. What­ever its origin, the pathogen seems to have affected frogs initially from 1979 to 1983. Bradford’s (1991) population was extinct by 1989. ­Because of population losses, Mountain Yellow-­legged Frogs have been repatriated into sites where they occurred formerly, in the hopes of reestablishing the species. In Sequoia National Park, for example, Fellers et al. (2007a) repatriated eggs, tadpoles, subadults, and adults into 4

previously occupied sites. Although survivorship was good the first year, the repatriation effort soon failed. Since fish ­were not pre­sent at any of the sites, Fellers et al. (2007a) attributed the failure to the effects of Bd or possibly the continuing effects of contaminants. In 2010, an experimental reestablishment effort began at Indian Creek in Hall Canyon in Ventura County. Tadpoles collected during a salvage operation at Dark Canyon ­were raised at the San Diego Zoo and bred in captivity. Progeny of ­these animals, including 300 eggs and 36 tadpoles, ­were released at this location in 2010; however, ­these individuals ­were not detected during surveys in 2011. In 2011, 313 tadpoles and 270 eggs ­were placed at the Indian Creek location; tadpoles ­were detected during subsequent monitoring in 2011. Additional re­introduction efforts have centered on the Angeles National Forest. In 2018–2019, more than 1,000 Los Angeles Zoo–­bred Mountain Yellow-­legged Frogs and tadpoles ­were released into a tributary to Cooper Canyon south of Palmdale. Since 2006, more than 3,800 captive-­bred Mountain Yellow-­legged Frog tadpoles and subadult males have been released into their historical habitat in the San Gabriel, San Bernardino, and San Jacinto mountains in Southern California. In addition, 50 tadpoles and 15 frogs from ­Little Rock Creek ­were relocated to the Los Angeles Zoo as the 52,730 ha Bobcat Wildfire threated to destroy R. muscosa habitat in October 2020. Hammond et al. (2020) have shown that using PIT tags with a long-­range PIT tag reader significantly increases detection probability of translocated individuals. The IUCN lists this species as Endangered, and the species is listed as Endangered ­under the US Endangered Species Act of 1973. The state of California also protects this species as Endangered. A conservation assessment designed to guide research and conservation activities on southern California National Forests has been prepared (Brown et al., 2014b).

Rana pretiosa Baird and Girard, 1853 Oregon Spotted Frog Grenouille maculée d’Orégon

NOMENCLATURE

ETYMOLOGY

pretiosa: Latin for ‘to be prized’ or ‘worth the effort.’ The name may refer to the reward of catching such an attractive frog.

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Dubois (2006): Rana (Amerana) pretiosa Fouquette and Dubois (2014): Rana (Rana) pretiosa Synonyms: Rana temporaria pretiosa Much of what has been written on Rana pretiosa includes information on the Columbia Spotted Frog, R. luteiventris (e.g., Nussbaum et al., 1983). The latter species was separated from R. pretiosa based on molecular data, and its distinctiveness has been subsequently confirmed (Green et al., 1997; Funk et al., 2008). Readers should consult the

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694  RANIDAE

location of past studies to verify ­whether the information contained refers to R. pretiosa or R. luteiventris. IDENTIFICATION

Adults. Adults are brown or reddish brown and become redder with age. Very old adults can approach a brick-­red coloration covering the dorsum. Juveniles, however, are brown or even olive green. The head and dorsum have black spots with light centers; ­these spots become darker and larger with age. When viewed from above, the eyes point upward rather than laterally, and they tend to focus dorsally as one might expect from a frog that spends much time in the ­water. The frog has a tan to orange dorsolateral fold that is lighter than the rest of the dorsum. The fold begins ­behind the eyes and tends to break up as it extends posteriorly. Juvenile venters are white or cream with red pigments ­under the legs and abdomen. Pigments become brighter with age. Adults have a bright orange-­red pigmentation ventrally that extends from the posterior of the frog ­toward its chest but not up the sides of the frog. Throats have a tan, brown, or gray mottled appearance on an other­wise white background. Mottling is absent on the groin or comprised of a black or gray coloration on a light background. This mottling pattern does not contain lines and spots of black, green, yellow, or red as it does in R. aurora. Webbing is complete on the hind feet. Dunlap (1955) provided extensive information on the color pattern and morphology of this species. Males are smaller than females. In the Cascades of Washington, males averaged 57 mm SUL (range 46–66 mm) and females 75 mm SUL (range 59–89 mm) (McAllister and Leonard, 1997); in Thurston County, Washington, males averaged 56 mm SUL (range 46–65 mm), whereas females averaged 66 mm SUL (range 51–76 mm). In Oregon, males averaged 54.6 mm SUL (range 50–64 mm) and females 68.8 mm SUL (range 55–78 mm) (Pearl et al., 2018). In British Columbia, males reach 64 mm SUL and females 80 mm SUL (Licht, 1974, 1975). The maximum recorded size is 107.5 mm SUL for a female in Washington at a locality where many females exceed 100 mm SUL (Rombaugh et al., 2006). Larvae. The tadpole is tan to dark brown to greenish, with a tail that may have small flecks or blotches. Oregon Spotted Frog larvae are similar to larvae of R. aurora but have a lighter (white-­or aluminum-­colored) belly. They also may have a pale gold coloration around the margin of the belly. Older tadpoles usually have metallic flecking on the head, body, and anterior tail musculature which is not accompanied by the deep gold or brassy pigments observed in R. aurora. The tail fin and tail musculature are of equal height. This species’ larvae may be indistinguishable from ­those of R. cascadae. The tadpole may reach 70 mm TL.

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Eggs. Eggs are black dorsally and white ventrally. The outer envelope is large, mea­sur­ing 10–15 mm in dia­meter. The vitellus averages 2.3 mm in dia­meter. Eggs have 3 gelatinous envelopes (Rombough and Bowerman, 2021). Previous reports of 1 or 2 jelly envelopes are in error and not infrequently based on confusing R. pretiosa with R. luteiven­ tris (e.g., Livezey and Wright, 1947; Morris and Tanner, 1969; see Rombough and Bowerman, 2021, for details). Rombough and Bowerman (2021) have noted that egg, membrane (vitelline/fertilization), and envelope dimensions change with stage of development and age, and that ­there is regional and individual variation in egg dimensions. Eggs are oviposited in rounded or globular jelly masses, but they are not as firm as in other species. The eggs also appear rather far apart from one another. DISTRIBUTION

Rana pretiosa historically occurred from northeastern California through southwestern British Columbia. Many populations have dis­appeared, however, and ­until recently ­there ­were only 3 known extant populations in British Columbia (lower Fraser River Valley), 6 in Washington, and 24 in Oregon (Pearl et al., 2009b). All populations in northeastern California (Pit River drainage, 5 historic localities) and the Willamette Valley in Oregon (Bury, 2008), and many populations from a large area of western Washington (Olympic Peninsula, Mount Rainier), have been extir-

Distribution of Rana pretiosa. Dark gray indicates extant populations; light gray indicates extirpated populations.

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Rana pretiosa 695

pated. However, Bohannon et al. (2016) found 13 previously unknown breeding sites in the Northern Puget Sound Basin of Washington, including in 3 new river drainages (Samish, Nooksack, and Sumas). Impor­tant distributional references include: range-­wide (Nussbaum et al., 1983; Green et al., 1997), British Columbia (Cowan, 1936; Weller and Green, 1997; Matsuda et al., 2006; Ovaska and Govindarajulu, 2010), California ( Jennings and Hayes, 1994b), and Washington (Meek and Elliot, 1899; Slater, 1955; Leonard et al., 1993; McAllister et al., 1993; McAllister, 1995; McAllister and Leonard, 1997; Bohannon et al., 2016). FOSSIL REC­O RD

No fossils are recognized.

28–87 m) in the Puget Trough (Bohannon et al., 2016). The species requires lakes, ponds, and slow-­moving streams with abundant shallow-­water emergent vegetation for concealment. Wetlands with 50–75% open ­water are preferred. The species selects wetlands with herbaceous and shrub macrohabitats that form continuous floating mats or mats interspersed with ­water (Popescu et al., 2013). In terms of microhabitat, tall and less dense vegetation and thick, submerged vegetation is preferred regardless of the floristic composition. Frog presence is associated with 3 m) lakes and ponds in the high mountains of the Sierra Nevada (Knapp et al., 2003). A ­great depth allows them to avoid being completely frozen in ice in winter or stranded in drying ponds in summer, as frequently occurs in shallow ­water bodies. Deep, cold ­water holds oxygen longer and offers the potential to avoid anoxic conditions that occur during the very long winters at high elevations. Larvae prefer cold ­water, and they are not found in ­water >27°C (Mullally and Cunningham, 1956b). Still, they may congregate in the warmer reaches of the pond during the day. At lower elevations, transformation occurs during the same season as oviposition, and larvae do not overwinter; at intermediate elevations, it appears that some frogs may overwinter whereas ­others do not (Zweifel, 1955). DIET

Egg masses of Rana sierrae. Photo: Roland Knapp

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Very ­little information is available. Larvae are benthic herbivores. Adults have been reported to feed on the larvae of Anaxyrus canorus, unspecified small invertebrates, and dragonfly nymphs (Mullally, 1953). They prob­ably eat any invertebrate they can put into their mouth.

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Rana sierrae 705

PREDATION AND DEFENSE

This species is extremely wary on land and readily jumps to ­water at the approach of an observer. They swim to the substrate and bury themselves in mud and vegetative debris or hide ­under rocks. Mullally and Cunningham (1956b) noted 1 individual that took refuge in a rodent burrow. In the ­water, frogs are much less wary, and they easily blend in with vegetation and the substrate. When attacked, this species may scream or puff up its body to distract or thwart potential predators (Camp, 1917). ­There is very ­little information on predators, but they likely include a number of species of birds (crows) and mammals. Coyotes have been reported to feed on tadpoles (Moore, 1929), and garter snakes (Thamnophis couchii, T. elegans) eat postmetamorphs (Mullally and Cunningham, 1956b; Brown et al., 2020). POPULATION BIOLOGY

Males reach sexual maturity at ca. 44 mm (4.3–4.5 mm) and females at 45 mm (44–47 mm). Growth rates are similar between the sexes, but vary considerably among individuals (Fellers et al., 2013). Asymptotic sizes (a mean of 65 mm [64–67 mm] for males and 76 mm [75–77 mm] for females) are reached at 3–4 yrs ­after sexual maturity, therefore at ca. 10 yrs of age (3 yrs as a tadpole and 6 yrs post-­ metamorphosis). Longevity is 14–16 yrs. Stream populations tend to be small (< 20 individuals), with annual survival rates of 0.55–0.89 (Brown et al., 2019, 2020). However, C. Brown et al. (2019, 2020) recorded stream populations of 227 and 547 individuals, and a beaver pond meadow held an estimated 36 males and 33 females. C. Brown et al. (2019) noted sex ratios of 0–0.52 males, but some streams had only females, perhaps suggesting differences in habitat occupancy during the survey periods. In

other populations, sex ratios ­were essentially 1:1 (Brown et al., 2020). Intermittent creeks actually may hold larger populations than small, permanent streams, perhaps ­because of the lack of predators (Brown et al., 2020). In small streams, recruitment is low, relatively small numbers of tadpoles are observed in any 1 year, and adults tend to live long, up to 13 yrs (Brown et al., 2020). In a stream-­meadow complex, Fellers et al. (2013) reported a population size of 45–115 adults (mean of 86, with 45 males and 41 females) over a 9 yr period based on a capture-­mark-­recapture survey protocol. During this period, they marked 348 males, 316 females, and 93 juveniles, and sex ratios ­were close to 1:1. Rana sierrae at the low-­elevation (2,200 m) site had high overwintering survival rates (mean 70%, range 44.6–95%) and grew at a fast rate (K = 0.73– 0.78). Annual survivorship was correlated with precipitation from the previous year. Frogs tended to be sedentary, with a minimum convex polygon home range estimate of 139 m2 (range 1.5–5,380 m2), but they had low site fidelity from one year to the next. Median distance movements between years ­were 84 m (range 1–1,365 m) and did not vary by sex. Although Bd was pre­sent and had been in the population for at least 13 years, the population showed no evidence of decline. Alpine populations tend to be larger than subalpine populations. For example, Joseph and Knapp (2018) estimated the population size of a reintroduced population of R. sierrae at their alpine site as 768 adults over the course of a 12 yr study, with only 172 adults at their subalpine site. COMMUNITY ECOLOGY

The introduction of nonnative trout species throughout the western states has had devastating effects on a number of amphibian species, including the Sierra Nevada Yellow-­ legged Frog. For example, the distribution of R. sierrae was strongly negatively correlated with the presence of trout in Yosemite National Park, based on a survey of 2,655 ponds and marshes; the frog was only found in 282 wetlands (Knapp, 2005). Although adult trout do not eat adult frogs, the trout consume a ­great number of insects and insect larvae that would other­wise be available to frogs. Trout also consume R. sierrae larvae. Thus, trout adversely affect frogs through trophic interactions as well as predation; in essence, they disrupt natu­ral food webs (Finlay and Vredenburg, 2007). DISEASES, PARASITES, AND MALFORMATIONS

Adult Rana sierrae. Photo: Dana Drake

Dodd_Canada_int_5pgs_B4.indd 705

The virulent pathogen Batrachochytrium dendrobatidis (Bd) was reported from both tadpoles and recent metamorphs based on collections from 1998 to 2000, 2001, and 2005 throughout the Sierra Nevada (Fellers et al., 2001, 2011;

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706  RANIDAE

Smith et al., 2017), and it prob­ably acts on R. sierrae populations much as it does on R. muscosa populations (see R. muscosa species account). Bradford et al. (2011) suggested that Bd was likely the leading cause of population declines in this species. Larvae typically have abnormalities of the oral disk, including missing keratinized mouthparts, depigmented jaw sheaths, and swollen labial papillae. Tadpoles with abnormalities ­were found at 70% of the sites and from 4.2 to 100% of the tadpoles examined per site. Sierra Nevada Yellow-­legged Frogs in Yosemite National Park ­were infected at a high percentage (67.5%) with the pathogen during surveys in 2005. Russell et al. (2019) found that ca. 30% of adult R. sierrae they examined had Bd, with a survival rate of ca. 50–60%; males had lower survivorship than females. Although mass die-­offs are sometimes associated with the pathogen, Russell et al. (2019) showed that the presence of the pathogen even without obvious mortality leads to reduced survivorship in wild populations. Using data from a 12 yr capture-­mark-­recapture study, Joseph and Knapp (2018) showed that high Bd infection intensities ­were associated with an increase in detectability and a reduction in survivorship in post-­reintroduction frog populations. When average infection intensities ­were high across a population, adults ­were more likely to gain infections and less likely to lose them. Survival also was intensity-­ dependent, and not surprisingly, the most heavi­ly infected frogs experienced the greatest mortality. Secretions from R. sierrae possess peptide mixtures that inhibit the growth of Bd (Woodhams et al., 2007). The peptides are produced by a variety of symbiotic bacteria and may be impor­tant in assisting in disease re­sis­tance and individual survival. Ranavirus also has been reported from Mountain Yellow-­legged Frogs, but Smith et al. (2017) suggested that it has not contributed to population declines in the Sierras. SUSCEPTIBILITY TO POTENTIAL STRESSORS

pH. Increased levels of acidity have been mentioned as a pos­si­ble cause of the decline of this species throughout much of its range. The LC50 for embryos is a pH of 4.37 and for hatchlings 7,000 surveys over a 20 yr period, the abundance of R. sierrae increased 7 fold at a rate of 11% per yr. The increases occurred in hundreds of populations throughout Yosemite. Laboratory ­trials suggested that recovery resulted from reduced susceptibility to Bd, as well as the removal of introduced nonnative fish. The results of Knapp et al. (2016) and Brown et al. (2014a) appear quite contradictory, as the latter found ranid frogs at only 47% of watersheds that had locality data from 1990 to 2001 and 2% of sites with locality data prior to 1990. ­These differences may reflect differences in sampling, that is, in lakes versus watershed streams where frogs ­were less abundant or con­spic­u­ous. Rana sierrae has been reintroduced into a number of streams in the northern part of its range to augment existing populations. Frogs ­were released in 2017 (22 individuals) and 2018 (60 individuals) in the Plumas National Forest; as of late 2018, 52 frogs ­were known to be still alive. Over-­ winter survival was lower (29%) for captive-­released frogs than for wild frogs (56%). Brown et al. (2020) suggested that augmentation programs should continue by releasing adults rather than eggs or tadpoles, inasmuch as ­these latter life stages have extremely poor survivorship in the streams they studied. This program is ongoing.

Dodd_Canada_int_5pgs_B4.indd 707

In a comparison of wild and captive-­raised and released frog movement patterns, Keung et al. (2021) found that wild frogs moved ­little (median of 50 m during the year following release. Although released frogs occasionally moved >100 m (128 m over 22 days, 346 m over 53 days), captive-­raised frogs mostly settled at a release site. All frogs stayed within 1 m of ­water during movements. As the season progressed ( June–­October), movement tendencies decreased. ­These results suggest that re­introduction programs need to account for ­water availability throughout the year at a release site, especially during times when streams dry or become intermittent (Keung et al., 2021). It seems clear that the presence of pathogens and the effects of climate drive post-­reintroduction population dynamics in this species and affect the success of re­introduction programs ( Joseph and Knapp, 2018). This species is considered Threatened in California and a Sensitive Species in Nevada. It also is protected by the US Endangered Species Act of 1973 as Endangered.

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F ­ amily Scaphiopodidae

Scaphiopus couchii Baird, 1854 Couch’s Spadefoot

couchii: a patronym honoring Darius N. Couch (1822– 1897), a US Army officer who collected vast numbers of natu­ral history specimens in México at his own expense while on leave.

when viewed dorsally. Tails are slender and sometimes blotched or have a dark spotted pattern. Jaws are narrow to medium in width and never cuspate. Larvae reach a maximum size of 25 mm TL. Descriptions are in Wright (1929) and Altig (1970). Eggs. Eggs are black dorsally and white ventrally with a colorless jelly mass surrounding them. ­There is 1 thick, gelatinous capsule surrounding the vitellus mea­sur­ing 2.4– 3.5 mm in dia­meter (Ortenburger, 1924; Grubb, 1972). The vitellus is 1.4–1.6 mm in dia­meter (Livezey and Wright, 1947).

NOMENCLATURE

DISTRIBUTION

ETYMOLOGY

Synonyms: Scaphiopus couchii rectifrenis, Scaphiopus laticeps, Scaphiopus rectifrenis, Scaphiopus varius, Spea laticeps IDENTIFICATION

Adults. Coloration is highly variable from reticulated green and black markings to mottled shades of black, green, and yellow, or brown to nearly uniform green with black spots. ­There also may be sexual dichromatism in color pattern, with males a uniform light green and females a darker yellowish green. Hind limbs are short with rear feet having a black, keratinized spade for digging. Venters are cream to dirty white. Eyes are large with a characteristic vertical pupil, and ­there is no boss between the eyes. Males have a large vocal sac that when distended is nearly 3 times the size of the head. Additional descriptions are in Strecker (1908). Adults are not sexually size dimorphic. The mean SUL of adult males in Arizona was 69.2–74.4 mm SUL (­Sullivan and ­Sullivan, 1985); Jennings and Hayes (1994b) gave an adult size range of 45–82 mm SUL in California. Based on museum specimens, Goldberg (2018d) recorded males from 54 to 75 mm SUL (mean 60.7 mm) and females from 51 to 83 mm SUL (mean 61.7 mm). The maximum size is 90 mm SUL (Degenhardt et al., 1996). Larvae. Tadpoles are small and blackish brown, and are lighter ventrally and on the tail than on the body. A profusion of coppery-­bronze flecks may provide an iridescent sheen. Larvae have an overall appearance of being short and plump, with the widest part of the body ­toward the posterior

Dodd_Canada_int_5pgs_B4.indd 708

Couch’s Spadefoot occurs from western Baja California and the Mojave Desert of California eastward across southern Arizona, New Mexico (including northward along the Rio Grande Valley), and Texas to southwest Oklahoma. The range includes much of central Texas west of the Balcones Escarpment (Smith and Buechner, 1947) and extends south well into central México. Isolated populations occur in southeastern Colorado and east central Arizona (Petrified

Distribution of Scaphiopus couchii

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Scaphiopus couchii 709

Forest National Park). The species is absent from mountainous regions, such as the Mogollon Rim, southern Rockies in New Mexico, and Sierra Madre in México. Impor­tant distributional references include: Arizona (Brennan and Holycross, 2006; Murphy, 2019; Holycross et al., 2021), Baja California (Grismer, 2002), California (Tinkham, 1962; Vitt and Ohmart, 1978; Jennings and Hayes, 1994b; Flaxington, 2021), Coahuila (Lemos Espinal and Smith, 2007b), Colorado (Hammerson, 1999), New Mexico (Van Denburgh, 1924; Degenhardt et al., 1996), Oklahoma (Sievert and Sievert, 2006), and Texas (Dixon, 2000, 2013; Tipton et al., 2012; Davis and LaDuc, 2018). FOSSIL REC­O RD

Pleistocene fossils of Couch’s Spadefoot are known from Arizona, New Mexico, Sonora, and Texas (Holman, 2003). The species is readily separated from other fossil scaphiopodids by a variety of osteological characters (see Holman, 2003), and is closely related to the fossil spadefoot Scaphio­ pus alexanderi.

cacti. In Texas, it is associated with black calcareous soils in the east and with any friable, well-­drained, sandy soil ­toward the western part of the range; the species prefers mixed scrub vegetation (Wasserman, 1957; Dayton et al., 2004). In California and New Mexico, it is associated with dry washes and the edges of sand dunes, such as Algodones Dunes and White Sands, presumably where it can burrow down and be in contact with moist soil (Mayhew, 1965; Degenhardt et al., 1996). It is also associated with irrigated agricultural lands. Dayton and Fitzgerald (2006) developed a habitat suitability model with high predictability for Couch’s Spadefoot in Big Bend National Park, Texas. They found that this species was associated with clay loam soils with good water-­holding capacity, temporary pools, and alluvial floodplains. Most populations occurred along the Rio Grande River or in the northern regions of the park at low elevations (80%), when air temperatures are around 20.5ºC, and when soil temperatures are >15ºC. They are generally not active at air temperatures 29°C (Pearson, 1955; Johnson, 2003; Ryan et al., 2015). Gosner and Black (1955) reported a minimum daylong temperature of 9.6°C as necessary for emergence in New Jersey, although Ball (1936) recorded activity at 7.5–8.6ºC in Connecticut. ­These differences are slight, but they could reflect latitudinal variation in temperature tolerance and acclimation.

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718  Scaphiopodidae

Eastern Spadefoots disperse extensively into the uplands surrounding breeding sites, and immigration and emigration patterns suggest directed rather than random movements (Dodd and Anderson, 2018). Timm et al. (2014) found that adults moved a mean of 130 m (range 3–449 m) immediately postbreeding on Cape Cod. They have been found >800 m from breeding sites in Florida (Pearson, 1955), and I found them widely dispersed in the sandhill habitats of north central Florida 95–914 m (mean 539 m) from the nearest breeding site (Dodd, 1996). Adults disperse exclusively at night (Todd and Winne, 2006). However, dispersal occurs slowly, with individuals usually moving only short distances from one movement to the next. As such, it can take considerable time to traverse long distances between terrestrial and breeding sites. Nearly all of their time is spent in burrows about 7–30.5 cm in length and from 2 to 4 cm in dia­meter in Florida (Pearson, 1955), and 0.10–0.96 m in depth on Cape Cod (Timm et al., 2014). They ­will also occupy gopher tortoise (Gopherus polyphemus) burrows and stump holes in the South (Dziadzio and Smith, 2016; Murphy et al., 2021). As winter approaches, spadefoots dig deeper into the substrate. Most animals do not dig more than 18 cm in depth, at least in the South. The burrow itself does not consist of an open tube, but instead may be filled with loose sand and soil through which the spadefoot makes its way each time it enters and departs from the burrow. Pearson (1955) noted that the burrow is open when the spadefoot is away foraging (photo in Johnson, 2003). They may remain within a burrow for as many as 104 consecutive nights (mean 9.5 nights). Pearson (1955) observed that spadefoots ­were active on only 8% of the nights sampled, and thus would be foraging or other­wise away from the burrow on only 29 nights during an entire year in Florida. In contrast, Ryan et al. (2015) and Timm et al. (2014) recorded activity on 43 and 61% of the nights that spadefoots ­were monitored in Connecticut and Rhode Island, respectively. Eastern Spadefoots tend to remain in the same burrow, and only 1 frog is found per burrow. However, they can also shift burrows during an activity season, the mean distance usually being approximately 3.3 m (range 0.6–11.6 m in Florida; Pearson, 1955) and ≤50 m on Cape Cod (Timm et al., 2014, with 1 individual moving 304 m). Johnson (2003) recorded 1 spadefoot using 4 burrows within 2.0 m, and Timm et al. (2014) found that most tracked spadefoots used ≤3 burrows. They may remain in the new burrow or return to the first one. Pearson (1955) provides examples of how spadefoots used multiple burrows over a period of up to 10 months. Spadefoots also overwinter in burrows, according to Pike (1886), on south-­facing slopes, at least in the Northeast. In the North, spadefoots have been found to a

Dodd_Canada_int_5pgs_B4.indd 718

depth of several meters under­ground (Pike, 1886; Ball, 1936). Home ranges are used from one year to the next; that is, S. holbrookii is rather philopatric in its choice of terrestrial habitat (Pearson, 1957; Timm et al., 2014). Home ranges of Eastern Spadefoots in Florida ­were mea­sured from 0.65 to 82.1 m2 (mean 9.9 m2). Males had average home ranges slightly smaller (8.4 m2) than females (10.3 m2), but ­there was such variance as to preclude statistical significance (Pearson, 1955). Home ranges within a sex did not overlap, leading Pearson (1955) to suggest that individuals ­were territorial to members of their own sex. It appears that most time is spent within a par­tic­u­lar area within a season, although occasional movements well outside the core area are not uncommon (about 10% of movements are of this type). Home ranges ­were much larger on Cape Cod, 45–21,108 m2, with no difference in ­either Minimum Convex Polygon estimates or home range length between males and females (Timm et al., 2014). Movements also vary with habitat type. For example, Pearson (1955) found greater movements in ecotonal habitats than within core hammock (forested) habitats. Shorter-­ distance movements also occur where spadefoot densities are highest. Patterns of movement and burrow use within home ranges ­were illustrated by Pearson (1955, 1957) and Timm et al. (2014). ­After breeding, Eastern Spadefoots return to the terrestrial home range from which they departed. Population density appears to be rather high in areas near breeding sites. Pearson (1955) estimated from 493.4 to 976.0 spadefoots/ha on a single 73 m2 plot over a 1 yr period, depending on month, and from 237 to 511.2 spadefoots/ha on another 5,520 m2 plot over a 2 yr period. However, he was not confident that the assumptions of mark-­recapture studies ­were met, as the spadefoots ­were usually inactive, and the marked animals did not seem to mix randomly with the total population; he thus considered his estimates as “very inaccurate.” ­Later adjusted counts still gave estimates as high as 631 spadefoots/ha, with considerable monthly variation in estimate (Pearson, 1957). Density decreases during the course of a calendar year within an area, and survivorship is highest as the spadefoots overwinter. Eastern Spadefoots are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their daily activities ( Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Eastern Spadefoots likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Males call both diurnally (reviewed by Neill, 1957a) and nocturnally. Calling begins as soon as individuals arrive at

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Scaphiopus holbrookii 719

the breeding site and may extend over a period of several days depending on environmental conditions. For example, Church et al. (2002) recorded from 9 to 50 adults at a breeding pond in ­Virginia over a period of 8 days. The distinctive call is difficult to describe, but has been likened to the second syllable in a long, drawn-­out meow of a Siamese cat (“owww, owww”), an explosive grunt, languishing moans, a young crow, “waagh,” or “waaank” (Palis, 1994). Calling occurs as males float in the ­water. They arch their heads and bodies convexly backward, allowing the expansion of the vocal pouch. Calls are uttered at 4–6 sec intervals at a fundamental frequency of 170 cps and a trill rate of 177 (McAlister, 1959). Not all males call, as many seem to await the approach of anything that might be a female, which they readily clasp. McAlister (1959) described the vocal anatomy and method of call production. Amplexus is inguinal. Many if not most females are amplexed as they make their way ­toward the breeding site rather than at the site. Even as they move ­toward ephemeral wetlands, females may be clasped by more than 1 male, although the primary suitor ­will issue a warning call and vigorously kick rival males that attempt to amplex the pair (Ball, 1936). The female swims during amplexus and determines the oviposition site. Once the eggs are expelled and fertilized, amplexus ends and the adults separate. Interspecific amplexus has been observed between S. holbrookii and D. chrysoscelis (Palis, 2019). Eastern Spadefoot calls have been recorded with dominant frequencies of 1,300–1,550 cps at 18ºC for durations from 0.52 to 0.62 sec (Blair, 1958b). The calls of S. holbrookii and S. hurterii are essentially the same in call characteristics and sound. BREEDING SITES

Reproduction usually occurs in small, shallow (100 metamorphs) occurred on only 5 occasions, however. Greenberg and Tanner

Dodd_Canada_int_5pgs_B4.indd 723

(2005a) noted that 4 ponds acted as source populations, and then only during some years. Some of the source ponds acted as “sinks” in other years, that is, no recruitment occurred despite a breeding event; 3 ponds always acted as sinks. At a small north central Florida temporary pond, ­there was only a single breeding event in 5 yrs, but no tadpoles ­were observed (Dodd 1992), although a few metamorphs ­were subsequently captured on the inside of a pond-­encircling drift fence. Breeding events without any recruitment appear to be common in Eastern Spadefoot populations. Greenberg and Tanner (2005a) further reported that from 0 to 4,648 metamorphs ­were produced per breeding event, that apparent population trends reflected breeding effort, that recaptures ­were rare, and that capture rates fluctuated substantially from one year to the next. The total number of breeding adults captured was positively associated with juvenile recruitment when summed over a 2 week period following the initial breeding event, but not for comparisons of breeding adults and subsequent recruitment based solely on the number of adults captured during the initial event (Greenberg et al., 2017a, b). Over the 9 yr study, from 0 to 7,370 recruits ­were captured in any 1 yr, whereas 15,145 metamorphs ­were captured in total over the study period (Greenberg and Tanner, 2005a). Some interpond movement was recorded, mostly to nearby sites. One spadefoot traveled 416 m between breeding sites, however. Average air temperature does not influence juvenile recruitment. As is evident from the previous paragraphs, populations of this species vary widely in abundance and demographic characteristics among years and breeding ponds, making ­simple trend analyses difficult at the landscape level (Greenberg et al., 2018a). In an example of this, t­ hese authors found that recruitment was negatively correlated with the abundance of adult S. holbrookii 2 years following breeding events, but positively correlated 4 years ­later. Low statistical power hampered an ability to detect trends even over a 24 yr period. COMMUNITY ECOLOGY

In mixed larval groups, Scaphiopus holbrookii larvae have complex and sometimes subtle interactions and effects upon both other anuran larvae and larval predators. For example, S. holbrookii is a superior competitor to Cope’s Gray Treefrog (Dryophytes chrysoscelis) in laboratory ­trials, and they actually grow better in the presence of Anaxyrus americanus and Lithobates sphenocephalus larvae than in their absence. Alford (1989a) suggested that spadefoots ­were competitively dominant to ­these species. However, Anaxyrus and Lithobates larvae scrape periphyton off mesocosm walls and release nutrients that enhance the growth of phytoplankton, a better food source for spadefoot larvae than filamentous algae (Wilbur et al., 1983). This may allow them to

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724  Scaphiopodidae

grow faster without directly negatively impacting the other species. In another experiment, the sunfish Enneacanthus completely eliminated larval S. holbrookii (Kurzava and Morin, 1998). In mesocosms where hydroperiod is controlled (for 100 days, 50 days), S. holbrookii larvae generally survive well at both low and high densities, achieving as high as an 80% survivorship (Wilbur, 1987). However, adding newts (Notophthalmus viridescens) to experimental tanks nearly eliminates survivorship, as newts seem to prefer Scaphiopus larvae to Anaxyrus, Dryophytes, and Lithobates tadpoles in mixed-­species assemblages (Morin, 1983; Wilbur, 1987). When newts are added to a mesocosm featuring only Scaphiopus, the initial density of the tadpoles (low or high) affects the larval period and mass at metamorphosis. At high tadpole densities, the newts tend to become satiated so that survivors are actually larger than they would be at low densities, where food might be more abundant but tadpoles are easy targets for the predators. Likewise, the absence of newts, or their introduction early in development, increases mean larval mass, especially when tadpoles are initially in high densities (Alford, 1989a). Increasing the number of predators further tends to increase the mass of surviving larval spadefoots; although survivorship decreases dramatically, the larval period decreases but mean body mass and growth rates increase (Morin, 1983). DISEASES, PARASITES, AND MALFORMATIONS

The amphibian chytrid fungus (Batrachochytrium dendroba­ tidis) has been found on S. holbrookii at Cape Cod, Mas­sa­ chu­setts (Tupper et al., 2011, 2014). Ranavirus was found in S. holbrookii in Illinois (Kirschman et al., 2017) and North Carolina (Lentz et al., 2021). Larval Eastern Spadefoots are sensitive to ranavirus, more so than to FV3. However, FV3 does not appear to result in larval mortality (Hoverman et al., 2011). Parasites include protozoans (Nyctotherus cordiformis, Octomitus intestinalis, Opalina oblanceolata, O. obtrigonoi­ dea, O. carolinensis, O. triangulata, Trichomonas augusta), cestodes (Distoichometra bufonis, proteocephalid cysts), and nematodes (Agamonema, Cosmocercoides dukae, Oswaldo­ cruzia leidyi, O. pipiens, Physaloptera, Rhabdias, R. ranae) (Brandt, 1936). An Eastern Spadefoot with malformed fin­gers is shown in Gibson and Anthony (2019). SUSCEPTIBILITY TO POTENTIAL STRESSORS

No information is available. STATUS AND CONSERVATION

Determining the status of Eastern Spadefoots can be difficult, since individuals are generally rarely observed, despite

Dodd_Canada_int_5pgs_B4.indd 724

Breeding habitat of Scaphiopus holbrookii. Photo: John Bunnell

intensive sampling, especially in the northern portions of the species’ range. For example, the first rec­ords of Eastern Spadefoots from ­Great Smoky Mountains National Park ­were only made in the late 1990s, despite intensive observations within the park for more than 70 yrs (Irwin et al., 1999). C.C. Abbott (1884) noted that Eastern Spadefoots bred in a nearby sinkhole 10 yrs apart. Nichols (1852) reported breeding at a site only 4 times in 30 yrs. Indeed, the species may be relatively common, despite the rarity of sightings or museum specimens from specific localities (Gentry, 1955). Individual rec­ords may be the result of chance observations (Ball, 1936; Brodman, 2003; in Montanucci, 2006), whereas intensive sampling at appropriate times of the year may prove them to be more widely distributed than previous observations would suggest (e.g., Rice et al., 2001; Gibson and Anthony, 2019). In this regard, Devan-­ Song et al. (2021) successfully used eyeshine surveys to detect large numbers of terrestrial adults and postmetamorphs in areas where few sightings had been recorded previously, perhaps offering an alternative sampling method for a species somewhat mistakenly thought to be overly secretive. Eastern Spadefoots may have been able to follow sandy river terraces into mountainous regions or into other­wise unsuitable habitat, where they could dig into sandy soils (Netting and Wilson, 1940; Klemens, 1993; Montanucci, 2006: Raithel, 2019). ­These areas are often affected by river damming and other riparian modifications, as well as development along river terraces. Habitat specificity plus subsequent fragmentation may account for the scarce and disjunct rec­ords from uplands habitats and from certain portions of its range. Thus, peripheral populations may be particularly vulnerable to habitat modification or loss, such as on Long Island and in southern Connecticut. This is also

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Scaphiopus hurterii 725

true of their temporary shallow breeding sites, which are virtually unprotected throughout the species’ range. Some well-­documented populations, such as in Ball (1936), are now extirpated. Moran and Button (2011) developed a GIS model to identify Eastern Spadefoot habitat in eastern Connecticut that may aid in the identification of habitat in other regions, especially in the North along riparian habitats. Although S. holbrookii may be found in wetlands surrounded by pine plantations, their numbers may be small, and it is unclear the extent to which site preparation and planting affects population viability. For example, Russell et al. (2002b) found only 6 spadefoots over a 2 yr period at 5 wetlands embedded within a pine plantation in South Carolina. The low number could reflect habitat disturbance, or perhaps appropriate environmental stimuli such as heavy bouts of rainfall ­were absent. In Florida, Enge and Marion (1986) found only 26 Eastern Spadefoots in maximum or minimum clearcut and site prepared pine plantations, but found 4,696 individuals in an uncut forest wetland over the same 1 yr sampling period. Roller chopping and heavy-­duty site preparation would seem particularly detrimental to this fossorial species. It is clear that large-­scale development proj­ects can have very negative effects on Eastern Spadefoots. This is ­because their unprotected ephemeral breeding sites are particularly prone to destruction, and terrestrial uplands are prime sites for industrial and residential development. Near Tampa, Florida, for example, Eastern Spadefoots ­were eliminated following the development of a large housing proj­ect, even as they remained abundant in adjacent undeveloped areas (Delis, 1993; Delis et al., 1996). This may be ­because of their inability to burrow into the unnatural substrates often used by landscapers, or ­because urban landscapes are heavi­ly watered ( Jansen et al., 2001) and saturated with pesticides.

Spadefoots may persist in suburban areas, but only if breeding and nonbreeding habitats are pre­sent, and friable soils are available for burrowing. On Kiawah Island, South Carolina, Eastern Spadefoots also are found only in low to moderate development areas (Hanson and McElroy, 2015). Roads undoubtedly take a serious toll on some populations (Sutherland et al., 2010). Creating wetlands to replace destroyed habitats may not be a successful conservation strategy for this species. Mitchell (2016) found this species at only 9% of agriculturally restored ponds in the mid-­Atlantic region. Although adults may visit created ponds, Pechmann et al. (2001) found no successful breeding for 8 yrs following pond creation at a South Carolina restoration site. In Mary­land, S. holbrookii did not colonize newly created ponds over a 2 yr period following construction (Merovich and Howard, 2000). However, Eastern Spadefoots ­were found around a number of created wetlands along the Patuxent River in Mary­land (Touré and Middendorf, 2002). A total of 1,916 S. holbrookii ­were exported from Florida for the pet trade from 1990 to 1994 (Enge, 2005a). This species is considered Endangered in Connecticut (Klemens et al., 2021), Ohio, Pennsylvania, and Rhode Island, Threatened in Mas­sa­chu­setts, Declining in New Jersey, Rare in West ­Virginia, and of Special Concern in New York. In Mas­sa­chu­setts, a proj­ect is underway to create potential breeding habitat for this species through translocation and the construction of temporary ponds in the Southwick Wildlife Management Area (see https://­www​.­mass​.­gov​ /­service-­details/taking-­action-­a-­new-­conservation-­effort-­for-­ the-­threatened-­eastern-­spadefoot). Pennsylvania developed a species action plan aimed at Eastern Spadefoot conservation (https://­www​.­amphibians​.­org​/­wp​-­content​/­uploads​/­2019​/­04​ /­species​-­plan​-­spadefoot​.­pdf).

Scaphiopus hurterii Strecker, 1910 Hurter’s Spadefoot

IDENTIFICATION

ETYMOLOGY

hurterii: a patronym honoring Julius Hurter (1842–1917), Missouri herpetologist and author of the monograph Herpetology of Missouri, published in 1911. NOMENCLATURE

Conant and Collins (1998): Scaphiopus holbrookii hurterii Synonyms: Scaphiopus holbrookii hurterii

Dodd_Canada_int_5pgs_B4.indd 725

Adults. The dorsal coloration is greenish brown with pale yellowish lines extending from each eye and converging posteriorly. This species is similar to S. holbrookii but has a narrower head and smaller tympanum. The parotoid glands are round, high, and con­spic­u­ous. Venters are yellowish white. As in all spadefoots, ­there is a distinct digging spade on each rear hind foot. Based on museum specimens, Goldberg (2019f) recorded Oklahoma males from 48 to 67 mm SUL (mean 60 mm) and females from 55 to 73 mm SUL (mean 60.1 mm). Sexual size dimorphism is not apparent. Wright and Wright (1949) give an adult size range of 43–73 mm SUL. A leucistic metamorph was reported by McKnight and Ligon (2013) from Oklahoma.

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Larvae. At hatching, tadpoles are black but quickly become dark gray. Chromatophores give the appearance of an hourglass shape when viewed dorsally. Heads are narrow with the small eyes close to one another. Venters are gray but lighter than the dorsum. Tail fins are generally clear and low, and the tail tip is rounded. Tails are longer than the body. Larvae reach a maximum size of 21–27 mm TL. Bragg (1944b) and Bragg et al. (1964) described the tadpole and its mouthparts. Eggs. Eggs are black dorsally and light gray to white ventrally. ­There is a single gelatinous capsule surrounding the egg that is ca. 6.7 mm in dia­meter; the vitellus is ca. 2.3 mm in dia­meter (Bragg, 1944b). Trauth and Holt (1993) gave a much smaller size range for recently oviposited eggs (mean 0.7 mm dia­meter; mean 2.01 mm with gelatinous membranes). Gelatinous coats are sticky and easily acquire mud or debris, making them difficult to see in muddy ­water. The gelatinous eggs tend to stick together in vari­ous patterns, although they often appear strung together. ­These egg strings can be short or extend many cm in length. DISTRIBUTION

Hurter’s Spadefoot occurs from eastern Oklahoma and western Arkansas south to northwest Louisiana and the Texas Gulf Coast. The species occurs east of the Balcones Escarpment in Texas (Smith and Buechner, 1947). An isolated population occurs in central Arkansas, but the ranges of this species and the Eastern Spadefoot do not overlap. Zweifel (1956a) indicated overlap in Louisiana, but this was based on an erroneous rec­ord in the Tulane collection (Wasserman,

1958). This species is l­imited to the eastern deciduous forest biome and does not occur in prairies. Impor­tant distributional references include: Arkansas (Trauth et al., 2004), northern Louisiana (Dundee and Rossman, 1989; Boundy and Carr, 2017), Oklahoma (Sievert and Sievert, 2006, 2021), and Texas (Hardy, 1995; Dixon, 2000, 2013; Tipton et al., 2012). FOSSIL REC­O RD

Pleistocene fossils of this species ­were reported by Davis (1973) from Peccary Cave in Arkansas. Although referred to S. holbrookii, the cave location is within the range of S. hurterii. Holman (1969) noted that 2 bones found in a cave in Texas could be referable to ­either this species or S. couchii. SYSTEMATICS AND GEOGRAPHIC VARIATION

Hurter’s Spadefoot was described from a single individual collected in Texas (Strecker, 1910a). Scaphiopus hurterii is closely related to S. holbrookii and S. couchii and is only distantly related to the western Spadefoot genus Spea (Tanner, 1939; Sattler, 1980; García-­París et al., 2003). Scaphiopus hurterii is often included as a subspecies of S. holbrookii. The 2 are geo­graph­i­cally separate, but their calls are essentially the same (Blair, 1958b), and they readily hybridize ­under laboratory conditions (Wasserman, 1958). Fertility of hybrid backcrosses is high. However, ­there are differences between the skulls of S. hurterii and S. hol­ brookii, especially in the fronto-­parietal bone (Smith, 1937; Tanner, 1939; Zweifel, 1956a). In addition, ­there are size differences, differences in the presence or absence of a boss (a raised area) between the eyes, and minor tadpole prey differences. García-­París et al. (2003) provided ge­ne­tic evidence supporting recognition of 2 distinct species. The 2 spadefoots also are treated as separate species by Lannoo (2005), Crother (2017), and Elliott et al. (2009). Natu­ral hybridization between S. couchii and S. hurterii has been reported in Texas (Wasserman, 1957). In laboratory crosses, ♀ S. couchii × ♂ S. hurterii pairings produced a high percentage of hybrids. However, ♀ S. hurterii × ♂ S. couchii crosses ­were inviable (Wasserman, 1957). Crosses between S. hurterii and S. holbrookii ­were fully successful regardless of which species was the parent (Wasserman, 1958). ADULT HABITAT

Distribution of Scaphiopus hurterii

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Scaphiopus hurterii is a woodland and savanna species that is found in oak-­hickory and oak-­pine associations. They also may be found in river floodplains that extend into grassland habitats. They prefer sand, sandy-­clay, or calcareous soils that afford them the opportunity to burrow. In Oklahoma, they are more likely to be associated with calcareous soils than they are in Texas.

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Scaphiopus hurterii 727

TERRESTRIAL ECOLOGY

Hurter’s Spadefoot is a nocturnal species that spends most of its time in well-­concealed burrows, emerging only to breed and occasionally forage. The terrestrial ecol­ogy and be­hav­ior of adult S. hurterii likely is similar to S. holbrookii, although specific information is unavailable. Recent metamorphs emerge en masse and move across the landscape in waves of thousands of tiny toadlets. Dispersal occurs both by night and by day ­until the tiny spadefoots reach their destination. Young S. hurterii seek shade and protection ­under debris and vegetation rather than by burrowing into the mud in order to avoid desiccation (Bragg, 1945b). Mortality in direct sunlight may be high. Dispersing toadlets may emerge from cover at night or ­after rain showers to feed. CALLING ACTIVITY AND MATE SE­L ECTION

Males enter breeding pools during or shortly ­after heavy rainfall and begin calling immediately. Males call mostly while floating in the deeper ­water of a temporary pool, although they may call from the bank or at the very edge of the pool (Bragg and Smith, 1942). Calling males on the bank tend to remain stationary, whereas ­those calling from the ­water move around a ­great deal. Females move to pools si­mul­ta­neously with males. Mate se­lection is by scramble competition; that is, males ­will move ­toward and attempt to amplex any spadefoot moving in the pond. However, a female must touch a calling male for him to amplex her (Axtell, 1958). Inguinal amplexus and oviposition occur quickly. Males frequently wrestle with other males in clasping attempts, but the clasped male gives a warning vibration and call, which ­causes the amplexing male to release its hold. The male advertisement call is a harsh, guttural, loud noise that can be heard at some distance and directs both males and females to the location of a breeding site. The call of S. hurterii is characterized by a minimum frequency range of 164–225 cps and a maximum frequency range of 245–375 cps, depending on temperature. Calls last from 0.3 to 0.6 sec (Blair, 1955c, 1958b) with a trill rate of 185 per sec (McAlister, 1959). The dominant frequency is 1,325–1,710 cps (Blair, 1958b). McAlister (1959) described the morphology of the vocal structures. Wiest (1982) noted calling at 13–18°C over a 5 day period in Texas.

calling from permanent, beaver-­formed lakes that contained predatory fish; ­later captures of metamorphs suggested the spadefoots bred at the site. Bragg (1964a) speculated that chemoreception might be impor­tant in breeding pool se­lection by this species but offered no empirical data. REPRODUCTION

Spadefoots are the epitome of explosive breeders. Reproduction occurs only ­after heavy rainfall and for a very short time. For example, large numbers of toadlets ­were captured following heavy rains (2.8 and 7.8 cm) on 2 days in mid-­ March in central Texas (Brown et al., 2014). Most eggs are oviposited the first night following torrential rains. Breeding may occur at any time of the year that has a favorable combination of warm temperatures and heavy precipitation. Based on histology, Goldberg (2019f) noted that males and females breed from February to September in Oklahoma, corresponding with lit­er­a­ture reports. The smallest mature male was 48 mm SUL, whereas the smallest mature female was 55 mm SUL in his museum-­based sample. In Oklahoma, breeding has been reported from March to September (Bragg and Smith, 1942; Bragg, 1944b, 1950a; Black, 1973b). Females tend to stagger breeding, such that not all females breed at the same time, regardless of favorable conditions. Other rec­ords of breeding dates include mid-­March to June in Arkansas (Trauth et al., 1990) and March–­April in Louisiana (Connior et al., 2015b). The mean clutch size, based on internal examination, of 8 females in Arkansas was 2,494 eggs (range 1,961–4,847) (Trauth et al., 1990). Counts for egg clusters of 6 females in Louisiana ­were 49–131 (mean 100.8) (Connior et al., 2015b). Another Arkansas rec­ord gave a mean of 111.8 eggs (range 35–216) per cluster (Trauth and Holt, 1993). It would appear that this species is capable of ovipositing multiple clutches/

BREEDING SITES

This species breeds in shallow temporary pools, puddles, gravel pits, muddy pools in woodlands, ponds, and ditches (Bragg and Smith, 1942; Bragg, 1944b). ­These pools may be devoid of vegetation and exposed to direct sunlight. Algae are usually pre­sent and serve as a food source. In contrast, McKnight and Ligon (2016) reported ­these spadefoots

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Adult Scaphiopus hurterii, amplexus with eggs. Photo: Stan Trauth

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clusters of eggs, perhaps allowing for opportunistic deposition over an extended period of time. Oviposition occurs in shallow ­water near the shore, with eggs being placed over detritus and vegetation. Hatching occurs from 16 hrs to 2 days (Black, 1973b); Bragg (1944b) described the hatching pro­cess in detail. LARVAL ECOLOGY

The larval period is extremely short, since spadefoots must complete metamorphosis in temporary pools that may dry very rapidly. Developmental periods last from 12–15°C (Preston, 2009); for example, spadefoots are observed from mid-­January through mid-­October in Kansas and Iowa, depending upon the weather, although peak activity is from late April to early July (Heinrich and Kaufman, 1985; Mabry and Christiansen, 1991; Collins et al., 2010). Plains Spadefoots may be active ­after light rains, but rain is not necessary for nocturnal activity. Adults cease activity prior to juveniles; juveniles presumably are active ­later in the year in order to maximize foraging and growth prior to dormancy. Spadefoots usually spend their days in under­ground retreats and emerge only at night to forage. They dig their own burrows backward using their keratinized digging spades, and they leave no trace of the burrow opening on the surface to indicate their presence. They also are frequently found in burrows of Black-­tailed Prairie Dogs (Cynomys ludovicianus) (Kretzer and Cully, 2001; Lomolino and Smith, 2003), gophers, or Spotted Ground Squirrels. Emergence from burrows occurs at temperatures >11°C (Bragg and Smith, 1942). Overwintering spadefoots dig deep into the soil and have been found 90 cm below the ground surface (Black, 1970). Juvenile (and

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732  Scaphiopodidae

presumably adult) S. bombifrons are freeze intolerant, but some individuals have been reported to be able to supercool to -4°C (Swanson and Graves, 1995). This could be impor­ tant for juveniles that might not be able to dig as deep as adults to avoid freezing temperatures. In the spring, S. bombifrons may be active several days prior to the ­actual arrival of a rain event that ­will initiate breeding. They can move distances of at least 1 km to a breeding site within a single night (Landreth and Christensen, 1971). In experimental ­trials, Plains Spadefoots ­were able to use the sun’s position for orientation ­toward a breeding site and to use celestial cues during postbreeding activities (Landreth and Christensen, 1971). Since they generally move at night to the breeding ponds, prebreeding surface activity may allow them to use celestial cues to orient in the right direction. Once a few individuals reach a breeding site and begin calling, auditory and perhaps celestial cues are used as a guide for ­later arriving animals. Upon metamorphosis, young S. bombifrons burrow rapidly into the mud in order to avoid desiccation (Bragg, 1945b). They emerge at night or ­after rain showers to feed but other­wise remain in the burrow. ­After about a week, the toadlets begin to disperse away from the breeding site, making a new burrow each night. According to Werner et al. (2004), adults may disperse up to 2.3 km from breeding ponds, although most spadefoots remain within 400 m. Hammerson (1999) recorded movements of 60–150 m per night over 1–2 nights of observation. Plains Spadefoots are photopositive in their phototactic response, suggesting they use both sunlight and moonlight when feeding and ­going about their activities ( Jaeger and Hailman, 1973). They are sensitive to light in the blue spectrum (“blue-­mode response”) (Hailman and Jaeger, 1974). Plains Spadefoots likely have true color vision. CALLING ACTIVITY AND MATE SE­L ECTION

Spea bombifrons is an explosive breeder, with choruses forming only ­after torrential rainstorms in spring and summer, although Gilmore (1924) reported the initiation of breeding ­after several days of drizzling rain. For example, Harings and Boeing (2014) found that S. bombifrons ­were detected calling most often only on the first night ­after heavy rains. Females arrive at pools shortly ­after chorusing has begun. Most breeding occurs at night, although rare breeding choruses form by day, especially in the late after­ noon (Gilmore, 1924; Trowbridge and Trowbridge, 1937; Black, 1970). Calling Plains Spadefoots have been observed at temperatures as low as 9–12°C (Bragg, 1945b; Hammerson, 1999). Males call while floating in the ­water (rarely from the shore), and they tend to stay in 1 location throughout the

Dodd_Canada_int_5pgs_B4.indd 732

calling period (Bragg and Smith, 1942). Creusere and Whitford (1976) recorded them most often in sparse vegetation >12 cm in ­water depth. Females enter the pool and move directly ­toward a calling male. Inguinal amplexus is initiated when the female actually touches the male. Even juveniles (28–38 mm SUL) have a well-­developed clasping reflex (Bragg, 1958b). Amplexed pairs are wary and readily dive below the ­water’s surface when alarmed. As the male tightly clasps his partner, abdominal contractions force the extrusion of a small number of eggs. At the same time, the male arches his back, bringing his vent in position to emit semen and fertilize the eggs. The call of S. bombifrons is a trill (likened to a “waah” or “waac” [Black, 1970] or a “weird plaintive cry” [Gilmore, 1924]) with a frequency of 1,250–1,800 cps, a range of 0.4–0.7 sec in duration, 22–47 trills per call, and 41–74 trills per sec (Blair, 1955c; McAlister, 1959; Northen, 1970; Forester, 1973). Specific values ­will vary with temperature. ­Sullivan (2019) describes the call as a “quack” in southeastern Arizona and notes the presence of both a fast and slow calling type within the state. Other lit­er­a­ture rec­ords of dif­fer­ent trill rates (fast and slow) within this species (e.g., Pierce, 1976) may refer to S. intermontana or S. multiplicata. The call has a long carry­ing capacity and can be heard >1.6 km away. McAlister (1959) described the morphology of the vocal structures. Plains Spadefoots spend only a short period of time at the breeding site. In the Sonoran Desert of Arizona, for example, the mean chorus duration was 1.8 days and the mean chorus size was 27 males (­Sullivan and ­Sullivan, 1985; ­Sullivan, 1985a, 1989). Most reproductive activity occurs during the first night of breeding. Even if calling males persist ­after the first night, ­little if any successful breeding takes place. Other reports (Gilmore, 1924) note chorus sizes in the hundreds. As might be expected, males outnumber females at breeding pools; the latter depart quickly ­after breeding. BREEDING SITES

This species breeds in temporary pools and puddles that form ­after heavy thunderstorms, but Plains Spadefoots may use slightly deeper ­water than other spadefoots. Anderson et al. (1999) recorded presence in pools 1,000 males ( Jennings and Hayes, 1994b). Males call while concealed at the edge of a breeding pool, with only part of the body immersed in ­water (Brown, 1976). They rarely are observed in the open area of the breeding pool. Calling occurs asynchronously, with many males calling at once. The call of S. hammondii is a slow trill with a dominant frequency of 1,370–1,600 cps, a range of 0.32–1.25 sec in duration, 17–45 trills per call, and 28–50 trills per sec (Brown, 1976); specific values vary with temperature. Storer (1925) likened the call to a prolonged, low-­toned “tirr-­r-­r-­r.” The call carries a long distance, and it has a ventriloquist quality that makes it difficult for a ­human observer to locate a calling male. Females make a somewhat muffled sound, especially when handled. BREEDING SITES

This species breeds in unshaded, shallow, temporary pools and puddles with ­water temperatures from 9 to 30°C (Brown, 1967). Eggs also may be deposited in temporary pools along or in stream courses or in road ruts (Ervin et al.,

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2005). Such pools lack predators such as crayfish, American Bullfrogs, and fish. In laboratory ­trials, tadpoles grew faster and had much greater survival through metamorphosis in alkaline ­water than in neutral tap ­water (Burgess, 1950). Many of the natu­ral breeding pools of spadefoots tend to be alkaline, and Storer (1925) noted that high alkalinity did not seem to be a deterrent to Western Spadefoots. REPRODUCTION

Eggs are deposited in shallow ­water (1,200 m3 of ­water. Both temporary and permanent ­water sources are used, especially man-­made reservoirs (Hovingh et al., 1985). ­Great Basin Spadefoots readily breed in very muddy and polluted ponds (from livestock manure), in stock tanks, and along both intermittent and permanent desert streams (Hovingh et al., 1985; Morey and Reznick, 2004; Buseck et al, 2005). Breeding sites often contain no vegetation. In Nevada, Linsdale (1938) reported breeding in a large field of shallow ­water formed by excessive snow meltwater along a river. In this case, no rain had fallen for a long period of time, but spadefoots still bred by the hundreds in the flooded pastures. Hovingh et al. (1985) concluded that any available standing ­water might be used as long as the total dissolved solids ­were 50 ng/L of azinphos-­methyl, S. inter­ montana experience a significant decrease in survivorship. Survivorship also significantly decreases as concentrations of diazinon increase. Significant mortality occurs at the highest lethal concentrations of endosulfan and diazinon (Westman et al., 2010). pH. In Utah, breeding occurs in pools with a pH of 7.2–10.4 (Hovingh et al., 1985). Breeding pools in California had a pH of 7.1–9.0 (Morey and Reznick, 2004). ­There are no reports for lethal pH values.

PREDATION AND DEFENSE

The secretive habits and cryptic coloration of this species make it difficult to locate. When in chorus, males w ­ ill cease calling at the approach of an observer. They burrow rapidly into the soil when disturbed, and the openings of the burrows are closed and difficult to locate, even where a toad was pre­sent a few seconds before. When handled, S. intermontana has a distinctive odor similar to that of peanuts (Waye and Shewchuk, 1995). It exudes a mucous secretion that serves to make the spadefoot unpalatable to predators. This secretion may cause allergic reactions similar to ­those of hay fever in the mucous membranes of some ­people. Predators of larvae include the larvae of the fly Tabanus punctifer ( Jackman et al., 1983) and possibly other aquatic invertebrates. Adults are eaten by snakes (Thamnophis sp.), birds (burrowing owls, crows), and mammals (coyotes), but ­little information is available (Wood, 1935; Harestad, 1985; Hammerson, 1999; in Buseck et al., 2005). POPULATION BIOLOGY

STATUS AND CONSERVATION

The G ­ reat Basin Spadefoot undoubtedly ­faces threats from habitat loss, alteration, and fragmentation. Other threats include the effects of toxic chemicals and impacts from invasive species, such as introduced fish (Buseck et al., 2005). High levels of road mortality of dispersing ­Great Basin Spadefoots have been reported (Logier, 1932: Crosby, 2014). Crosby (2014) reported substantial occurrence of spadefoots on roads in the Okanagan Valley of British Columbia, where ­Great Basin Spadefoots comprised 46.5% of observed amphibian mortality. Installing fences, however, reduced road occurrence considerably (by 89.6% in double-­fenced sections).

According to Nussbaum et al. (1983), the age at first breeding is 2–3 yrs (i.e., during their third summer). In males, maturity occurs ca. 40 mm SUL, whereas females mature at ca. 45 mm SUL. DISEASES, PARASITES, AND MALFORMATIONS

The biflagellated alga Chlorogonium has been observed as an epizoic symbiont of S. intermontana larvae in Utah (Drake and Trauth, 2010). The parasitic trematode Ribeiroia ondatrae has been reported from a single individual S. inter­ montana ( Johnson et al., 2002). Other helminths include the trematode Polystoma nearcticum, the cestode Distoichome­ tra bufonis, and the nematodes Aplectana incerta, Physalop­ tera sp., and Acuariidea gen. sp. (Goldberg and Bursey, 2002b). No parasites ­were observed in 5 S. intermontana from Idaho (Waitz, 1961).

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Adult Spea intermontana. Photo: David Dennis

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748  Scaphiopodidae

Concern has been expressed about how climate change ­will affect many frog species. In experimental ­trials, O’Regan et al. (2014) showed that S. multiplicata displayed ­great phenotypic plasticity in response to warming and artificial drying (altering ­water depth) in mesocosms. Warming has a negative effect on time to metamorphosis (i.e., the larval period decreases), but survivorship increased, and mass and length ­were only slightly reduced at metamorphosis. Tadpoles from permanent pools ­were larger and weighed more than larvae from temporary pools, but larvae from temporary pools had greater survival rates and shorter larval periods. ­These mixed results suggest that warming and drying effects are additive or antagonistic, not synergistic. Pool warming and drying may not always be harmful to this species, but their interacting effects suggest that antagonistic responses are difficult to predict. Throughout much of its range, the ­Great Basin Spadefoot is relatively common (e.g., Hammerson, 1999), but declines in populations in British Columbia have been noted (Weller and Green, 1997). The species is considered Threatened by COSEWIC. In areas where ­Great Basin Spadefoots are threatened, such as the Okanagan Valley of British Columbia, constructing breeding ponds and restoring wetlands allows for per­sis­tence and connectivity among isolated populations, where spadefoots colonized 18 of 21 ponds and produced metamorphs in 13 from 2007 to 2014 (Ashpole et al., 2018). In the United States, Buseck et al. (2005) discussed conserva-

Spea multiplicata (Cope, 1863) Mexican Spadefoot ETYMOLOGY

multiplicata: from the Latin multi meaning ‘many’ and plicatus meaning ‘folded’ or ‘braided.’ NOMENCLATURE

Synonyms: Scaphiopus dugesi, Scaphiopus hammondii multiplicatus, Scaphiopus multiplicatus, Spea hammondii stagnalis, Spea multiplicata stagnalis, Spea stagnalis Much of the lit­er­a­ture on “Scaphiopus hammondii” actually refers to Spea multiplicata (e.g., all of A.N. Bragg’s work in Oklahoma; Cowles, 1924; Ortenburger, 1924; Slevin, 1928; Wright, 1929; Blair, 1955c; Brown, 1967 [in part]; Ruibal et al., 1969; Aitchison and Tomko, 1974; Whitaker et al., 1977). It is necessary to check collecting localities in original citations in order to determine which species is being discussed.

Dodd_Canada_int_5pgs_B4.indd 748

Breeding habitat of Spea intermontana. Photo: Dana Drake

tion mea­sures necessary to ensure the species’ survival in Wyoming. ­Because of the stochastic nature of breeding, status surveys frequently fail to detect the presence of ­Great Basin Spadefoots, even though historical rec­ords may be available (e.g., Bradford et al., 2005a).

IDENTIFICATION

Adults. The background coloration is gray, tan, or light brown, with color changes occurring depending on ­whether the spadefoot is observed by day or by night. For example, an individual might appear light brown at night and grayish brown during daylight. Dark pigmentation surrounds the dorsal warts and is pre­sent on the hind legs. Small dark spots or blotches and red-­tipped tubercles may be pre­sent dorsally. ­There is no distinct boss between the eyes, and the pupils are vertical, indicating nocturnal activity. The iris is pale copper. No dorsolateral stripes are pre­sent. As in other spadefoots, ­there is a black, wedge-­shaped, keratinized spade on each hind foot. Males have a dark, heavi­ly pigmented vocal sac and keratinized patches on the first 3 digits of the forelimbs. The vocal sac is slightly bilobed when expanded. Sexual size dimorphism is absent. Brown (1976) reported a mean SUL for Arizona males of 46.9 mm (range 41–51.9 mm). Mean SULs of adult males in other Arizona populations ­were 48–51.3 mm SUL (­Sullivan and ­Sullivan, 1985). Based on museum specimens, Goldberg (2019b) reported New Mexico males from 34 to 56 mm SUL (mean 46.6 mm) and females

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Spea multiplicata 749

from 40 to 58 mm SUL (mean 47.2 mm). Degenhardt et al. (1996) reported a maximum size of 65 mm SUL. Larvae. The body is olive to gray and somewhat transparent. It tends to be globular to depressed and wider anteriorly than posteriorly when viewed from above. Eyes are close together and dorsally positioned. The jaws are wide and frequently cuspate. In carnivorous tadpoles, ­there is a distinctive beak on the upper jaw and a notch into which it fits in the lower jaw; omnivorous tadpoles have a more “typical” oral apparatus. The tail fin does not arise abruptly from the body. Larvae reach a maximum size of 74 mm TL (Cowles, 1924). Altig (1970) included this species in his tadpole key. Detailed descriptions are in Wright (1929) ­under the name Scaphiopus hammondii. Larvae of S. multiplicata are nearly impossible to discriminate from S. bombifrons larvae, particularly in shallow-­water playa wetlands. The carnivorous larval morph of S. bombi­ frons is generally larger than the carnivorous morph of S. multiplicata in deep-­water playa wetlands, but omnivorous larvae of ­these species cannot be phenotypically differentiated in shallow-­water playa wetlands (Ghioca-­Robrecht and Smith, 2013). Body mass also decreases for S. multiplicata omnivorous larvae as ­water recedes, but not for the omnivorous morph of S. bombifrons. Tadpoles of ­these species also tend to be larger in cropland wetlands than in grassland wetlands. Eggs. Spea multiplicata eggs range in color from light cream to gray black, although most are brown to dark brown. A single clutch may have dif­fer­ent colored eggs. The eggs are small with a thin gelatinous capsule; their dia­meter is 1.16–2 mm with a mean of 1.52–1.8 mm (Ortenburger, 1924; Pomeroy, 1981). The smallest eggs are gray whereas the largest eggs are red brown (Pomeroy, 1981). The eggs are attached to one another via a thin stalk 5–10 mm in length. Egg complements are oviposited in small masses which, according to Ortenburger (1924), are arranged spirally around vegetation.

Holycross, 2006; Murphy, 2019; Holycross et al., 2021), Colorado (Hammerson, 1999), Oklahoma (Sievert and Sievert, 2006), New Mexico (Van Denburgh, 1924; Degenhardt et al., 1996), and Texas (Dixon, 2000, 2013; Tipton et al., 2012; Davis and LaDuc, 2018). FOSSIL REC­O RD

Fossils of the Mexican Spadefoot are known from Pleistocene deposits in New Mexico (Holman, 2003). Skeletal features are essentially identical with S. hammondii and S. intermontana. SYSTEMATICS AND GEOGRAPHIC VARIATION

The phyloge­ne­tic relationships within the genus Spea historically have been unclear, and ­there have been differing interpretations of relationships (Tanner, 1939; Bragg, 1945b; Wiens and Titus, 1991; García-­París et al., 2003; Holman, 2003). Brown (1976) noted differences in the mating calls and physiology between California S. hammondii and Arizona “S. hammondii” (i.e., S. multiplicata), and separated ­these former subspecies into full species. Tanner (1989) considered S. multiplicata to be a subspecies of S. hammon­ dii and relegated Spea stagnalis to subspecific status, Spea hammondii stagnalis, based on osteological characters. However, Wiens and Titus (1991) considered S. h. stagnalis to be conspecific with S. multiplicata. Based on fossils, Holman (2003) identified 2 clades within the monophyletic genus Spea, an eastern clade consisting of S. bombifrons and a western clade consisting of S. hammondii, S. intermontana, and S. multiplicata. In Holman’s (2003) scenario, the eastern clade appeared by the ­middle Miocene, whereas the western clade first

DISTRIBUTION

The Mexican Spadefoot occurs from southeastern Utah and southwestern Colorado south throughout eastern and central Arizona to Guerrero and Oaxaca, México. Populations are found from southeastern Colorado and the Oklahoma and Texas panhandles southward throughout much of central (west of the Balcones Escarpment; Smith and Buechner, 1947) and western Texas. Murphy (2019) reported an apparently isolated population north of the ­Grand Canyon based on museum specimens listed in VertNet, but ­these specimens are most likely misidentified S. intermontana. Hammerson (1999) noted a number of instances of misidentification, leading to confusion about distribution. Impor­tant distributional references include: Arizona (Brennan and

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Distribution of Spea multiplicata

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750  Scaphiopodidae

appeared in the late Pliocene. This would make S. hammon­ dii the oldest member of the western clade, with the other members branching in the Pleistocene. Molecular data, however, do not support this scenario. Instead, S. multipli­ cata was likely the first to branch from the ancestral Spea (Sattler, 1980; Wiens and Titus, 1991; García-­París et al., 2003). In addition, ­there appear to be 2 separate lineages (Arizona, México) within S. multiplicata, which indicates the presence of additional ge­ne­tic diversity and could lead to the recognition of cryptic species (García-­París et al., 2003). Natu­ral hybridization between S. multiplicata and S. bombifrons has been reported in southeastern Arizona (Brown, 1976; Simovich, 1985, 1994; Simovich et al., 1991; Pfennig, 2003), the Four Corners region of the Colorado Plateau (Northen, 1970), and from southwestern New Mexico and Texas (Forester, 1973; Sattler, 1985). Northen (1970) noted intermediate calls of natu­ral hybrids between S. multiplicata and S. intermontana. Experimental hybridization between S. multiplicata and S. hammondii indicates a high level of ge­ne­tic compatibility (Brown, 1967; Forester, 1975), although hybrids involving ♀ S. multiplicata and ♂ S. bombifrons have lower survival and longer larval periods than nonhybrid S. multiplicata (Pfennig and Simovich, 2002). Ge­ne­tic compatibility is also high for crosses of S. multiplicata with S. intermontana and S. bombifrons (Littlejohn, 1959; Brown, 1967). ADULT HABITAT

This species occurs in plains and desert grasslands, pinyon-­ juniper woodlands, sagebrush flats, river valleys, semiarid shrublands, creosote-­mesquite landscapes, succulent desert scrub, and even in the open ponderosa pine forest savanna of north central Arizona ( Jameson and Flury, 1949; Aitchison and Tomko, 1974; Boeing et al., 2014). It is found in short-­grass prairies as well as in agricultural pastures and croplands. The species occurs to 2,134 m in Arizona (Aitchison and Tomko, 1974) and 900–2,600 m in New Mexico (Degenhardt et al., 1996). TERRESTRIAL ECOLOGY

Activity may occur at any time during the warm season when temperatures are 10–24°C (Hammerson, 1999). The Mexican Spadefoot is a nocturnal species that spends most of its time in well-­concealed burrows, emerging only to breed and occasionally forage. They have been known to selectively forage on ­cattle dung, where presumably insects gather for food and moisture (Barragán-­Ramírez et al., 2014). Spea multiplicata enter burrows in the fall (e.g., September in southeastern Arizona; Ruibal et al., 1969) and do not emerge in numbers ­until heavy summer

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rains fall the next July. Thus, they spend at least 9 months of the year within the burrow. Feeding occurs only rarely once a spadefoot has entered its cool-­season burrow. Nonbreeding habitats are in sparse to dense grasslands within 300 m of the breeding site (Creusere and Whitford, 1976). Positions within the burrow change seasonally according to temperature and moisture conditions. Mean depths early in the season are shallow (ca. 24 cm below the surface), but by midwinter are a mean of 54 cm below the surface. With the coming of warmer weather in June, spadefoots again move ­toward the surface at a mean depth of 42 cm. The maximum depth at which Ruibal et al. (1969) recorded a burrowed Mexican Spadefoot was 91 cm below the surface. During the rainy reason, burrows are very shallow (ca. 4 cm). The spadefoot digs the burrow using its hind foot spade, and the burrow is backfilled so that the vertically positioned spadefoot is enclosed within a small chamber in the tunnel. Ruibal et al. (1969) recorded temperatures of 17–22.4°C within the burrows they examined early in the dormancy season, and 5–15°C in winter. In summer, the shallow burrows can reach 27–39°C. Emergence occurs in late summer, although a few Mexican Spadefoots may emerge in June or early July ­after light rain. It appears likely that some individuals are active nocturnally at the mouths of their burrows prior to breeding emergence. Most animals, however, emerge in large numbers to breed only ­after very heavy (>5 mm) rainfall, such as occurs during desert thunderstorms. Mexican Spadefoots emerge in response to the low frequency sounds (12 cm (Creusere and Whitford, 1976). Most Mexican Spadefoots call while floating in the ­water with only their heads above the surface, and they tend to move around frequently throughout the calling period. In the Sonoran Desert of Arizona, the mean chorus duration was 1.6 days, and the mean chorus size was 44 males (­Sullivan and ­Sullivan, 1985; ­Sullivan, 1985a, 1989). Males outnumber females at breeding pools. ­Sullivan (1989) suggested that skewed sex ratios resulted from male attempts to breed at ­every opportunity, whereas females breed only once, regardless of how many favorable opportunities arise throughout the breeding season. Males associate with conspecific males and are attracted to a chorus by the advertisement call of conspecifics (Pfennig et al., 2000). In Arizona, for example, Pfennig et al. (2000) found a mean nearest neighbor distance of 6.6 m, with most spadefoots 3,000/m3 early in the larval period that gradually decreased in volume to ca. 1,000/m3 as development proceeded. Tadpole aggregations usually remain together throughout the developmental period ­until about 4 days prior to metamorphosis. The SUL of metamorphs was 12–29 mm in Arizona (Pfennig et al., 1991). Pomeroy (1981) found that carnivorous morph tadpoles ­were 19–31 mm at metamorphosis, whereas omnivore morph tadpoles ­were 16–26 mm at metamorphosis. DIET

Spea multiplicata are generalists whose diet includes many types of beetles, lepidopteran larvae, ants, crickets, flies, true bugs, spiders, pillbugs, cicadas, leafhoppers, aphids, and termites (Ruibal et al., 1969; Whitaker et al., 1977; Dimmitt and Ruibal, 1980a; Punzo, 1991a). Termites in par­tic­u­lar made up a large part of the diet of Mexican Spadefoots in southeastern Arizona. Assuming termites and beetles ­were the primary prey, and that a spadefoot could eat a large number in a single feeding, Dimmitt and Ruibal (1980a) estimated that an adult would need 6.8 feedings to

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obtain the amount of fat metabolized in a single year. A single spadefoot could eat 36% of its body weight in a single feeding. Newly metamorphosed juveniles consume beetles (many taxa, especially click and leaf beetles), spiders, hemipterans, ants, and lepidopteran larvae (Smith et al., 2004). Some vegetation also may be ingested while feeding. PREDATION AND DEFENSE

Calling males readily dive underwater when alarmed. This species possesses a noxious skin secretion. To a ­human observer, the secretion has an odor of raw peanuts, buttered popcorn, or garlic, is irritating to the eyes and nose, and produces an allergic reaction similar to hay fever. ­Others have reported the secretion to be actually painful. When handled or confronted by a predator, Mexican Spadefoots puff up and vocalize “eh, eh, eh, eh, eh” (Hammerson, 1999). Both sexes produce this sound. They also arch their bodies and close their eyes, extend the front limbs, and tightly tuck the rear limbs into the body (Livo et al., 1997). In experimental ­trials, larvae are eaten by a wide variety of potential predators, indicating no chemically based defensive mechanisms. In addition, they do not change their levels of activity in the presence of predators. A lack of larval antipredator mechanisms may partially explain why this species breeds in temporary rather than permanent pools (Woodward, 1983). Reported predators include garter snakes (Thamnophis cyrtopsis, T. eques, T. marcianus) and Loggerhead Shrikes (Woodward and Mitchell, 1990; Seiler et al., 2017; Ford, 2020; Jones and Hensley, 2020; Jones et al., 2020). POPULATION BIOLOGY

In a study of ge­ne­tic population structure, Mims et al. (2015) found that ­there is greater ge­ne­tic structuring of populations with increasing ­water requirements in desert anurans, and this pattern is illustrated in the population ge­ne­tic structure of Mexican Spadefoots. For S. multiplicata, the scale of population structure was relatively consistent throughout the Madrean Sky Islands of southeastern Arizona, with relatively low levels of differentiation and diffuse spatial structuring by mountain range, perhaps enhanced by this species’ mobility. Ge­ne­tic distances ­were most correlated with uniform landscape re­sis­tance, with most support for the isolation by distance hypothesis. Mobility coupled with an ability to use ephemeral breeding sites makes this species more genet­ically uniform than species with dif­fer­ent life history characteristics, such as Canyon Treefrogs and Red-­spotted Toads. Still, inasmuch as aquatic habitats are threatened throughout the Desert Southwest, ­there is a potential threat of loss of ge­ne­tic diversity, especially since ge­ne­tic connectivity is highly corelated with terrestrial and aquatic connectivity.

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Spea multiplicata 755

Adult Spea multiplicata. Photo: Rob Schell COMMUNITY ECOLOGY

Two or 3 dif­fer­ent species of spadefoot may breed in close proximity to one another (e.g., Spea multiplicata, S. bombi­ frons, Scaphiopus couchii), yet the species often do not breed in the same temporary pool. One species ­will breed in one pool, while a dif­fer­ent species breeds in a pool only a short distance away. On the infrequent occasions when they do breed together, they often occur in rather equal numbers (Bragg, 1945b). Differences in mating calls help keep the species from interbreeding (Forester, 1973). It also seems likely that a priority effect occurs; that is, the species that arrives at a pool first claims that pool, and the pool subsequently is avoided by other spadefoots. Conspecifics may cluster together when calling from the same pool. For example, S. couchii may call from the same pools as Spea multiplicata and S. bombi­ frons, with Scaphiopus couchii preferring the deeper parts of the pool. ­There appears to be no real macrohabitat segregation among species, however (e.g., McAlister, 1958). Natu­ral hybridization between S. multiplicata and S. bombifrons has been reported in southeastern Arizona, where it tends to be more prevalent at ephemeral pools at lower elevations. Hybridization also occurs more frequently in smaller pools, where the dif­fer­ent species are likely to come into close contact (Simovich, 1994). Large numbers of F1 hybrids and the offspring of hybrid backcrosses occur in the area. Spea bombifrons females are more likely to hybridize than S. multiplicata females or S. bombifrons males (Simovich, 1985; Pfennig and Simovich, 2002). Hybrid males are sterile, and hybrid females produce about half the eggs of the parentals. In laboratory experiments, Simovich et al. (1991) found that hybrid tadpoles developed faster and to a larger size than the tadpoles of ­either parental species; they also had higher survivorship. However, they tended to

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be intermediate in size at metamorphosis when compared to the parentals. Thus, the life history characteristics of the hybrid offspring may afford them some advantages over the parentals, especially when exposed to conditions of short hydroperiods, despite prob­lems with fertility and fecundity. Not surprisingly, hybridization tends to occur most often in the most temporary ponds, and the duration of hydroperiod is inversely correlated with hybridization. Pfennig (2003), however, found that the rates of hybridization between ­these species had declined over a 27 yr period in this region. ­After testing a number of hypotheses, Pfennig (2003) concluded that despite the larval advantages in short hydroperiod ponds, the hybrids are less fit than the parental species, and thus reproductive isolation as a result of the evolution of premating isolating mechanisms (termed reinforcement) best explained decreasing hybridization. In addition to hybridization, the presence of Plains Spadefoots has specific life history consequences for the less competitive Mexican Spadefoot in areas of sympatry. Spea multiplicata females tend to be in poorer body condition when they are in sympatry with S. bombifrons than they are in allopatry. Since the abundance of S. bombifrons is inversely correlated with mean body condition of female S. multiplicata, this suggests that severe competition is correlated with poor body condition in the latter species (Pfennig and Martin, 2009). Mean egg size is larger in allopatric S. multiplicata females than when in sympatry with S. bombifrons, and females with the best body conditions produce the most carnivorous larvae that are the most ­adept at foraging, regardless of ­whether in sympatry or not. Taken together, ­these results indicate that ­there is a maternal effect that mediates population divergence and character displacement when faced by severe competition (Pfennig and Martin, 2009), and that competition plays an impor­tant role in divergent trait evolution in spadefoots (Rice et al., 2009). When S. multiplicata and S. bombifrons breed in the same pool, the frequency of the dif­fer­ent tadpole morphotypes is very dif­fer­ent than when only 1 species is found in a pool. When sympatric, S. multiplicata larvae nearly all take on the omnivorous morphotype, whereas S. bombifrons larvae assume the carnivorous morphotype. This is not surprising, since S. multiplicata is a superior competitor on detritus, whereas S. bombifrons is a superior competitor on fairy shrimp. When allopatric, however, both species’ larvae develop both carnivorous and omnivorous morphotypes. The differences in the frequency of the morphotypes appear to have evolved in sympatry in order to reduce competition for scarce food resources. The manner in which ­these desert species partition resources by changing larval morphology is an example of character displacement (Pfennig and Murphy, 2000, 2002, 2003).

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756  Scaphiopodidae

DISEASES, PARASITES, AND MALFORMATIONS

Bd has been detected on this species along the Mexican–­US border (Sigafus et al., 2014). The Mexican Spadefoot is parasitized by metacercariae of the trematode Clinostomum attenuatum (Miller et al., 2004). Adult and juvenile spadefoots living in playa grasslands tend to have much smaller parasite loads of C. attenuatum than ­those living in agricultural areas. This may be ­because of the differences in hydroperiods (short in agricultural areas, long in playa grasslands) that cause disruptions in the parasite’s life cycle (Gray et al., 2007c). Yearly variation in parasite load also is prob­ably related to hydroperiod. Other parasites include the cestode Distoichometra bufonis, the nematodes Aplectana incerta, A. itzocanensis, and Physaloptera sp., and the protozoans Opalina sp. and Nycto­ therus cordiformis (Parry and Grundman, 1965; Goldberg et al., 1995). The mite Eutrombicula alfreddugesi is known from Mexican Spadefoots in Texas (Mertins et al., 2011).

Breeding habitat of Spea multiplicata. Southeast Arizona. Photo: David Pfennig

SUSCEPTIBILITY TO POTENTIAL STRESSORS

Chemicals. Spea multiplicata was not affected by the herbicides Roundup WeatherMAX® or Ignite® 280SL in experimental ­trials. However, juvenile survivorship was reduced when exposed to Roundup Weed and Grass Killer Ready-­to-­use Plus® (both at 1.33 ml glyphosphate/m2) for 48 hrs (Dinehart et al., 2009). STATUS AND CONSERVATION

Undoubtedly, Mexican Spadefoots face the usual threats from habitat loss, alteration, and fragmentation. ­There are no indications, however, that this species has declined throughout much of its range (e.g., Hammerson, 1999). The effects of habitat alteration on this species may be unclear or even misleading. For example, larval densities at playa breeding sites are not dif­fer­ent among playa habitats with dif­fer­ent adjacent land uses, suggesting widespread accommodation to ­human activity (Anderson et al., 1999; Ghioca, 2005). Abundance may actually appear high in agricultural areas when compared to other land uses (Gray et al., 2004b). On a landscape scale, occupancy of playa wetlands is positively associated with decreasing distance between pools and with increasing interplaya landscape complexity. Such a relationship may be of recent occurrence, as such nestedness patterns may result from the inability of this small species to disperse across the geometrically complex agricultural landscapes that are found in the Texas Panhandle (Gray, 2002; Gray et al., 2004a, 2004b). In addition, a major predator of spadefoot larvae, Ambystoma mavortium, tends to be less common in agricultural settings. Thus, habitat fragmentation historically might have affected this species positively and negatively

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despite the species’ current abundance and ability to persist in agricultural landscapes. Habitat alteration also may have increased the likelihood of hybridization in areas of contact, such as in southeastern Arizona. Simovich (1994) noted that vegetation and preci­ pitation runoff patterns changed substantially ­after the introduction of ­cattle grazing. Runoff previously allowed to form in scattered pools is now diverted to more permanent stock tanks. Spadefoots now breed in ­these pools, which results in dif­fer­ent species breeding in close proximity. Modification of ­water flow patterns coupled with periodic drought could exacerbate hybridization potential. A threat to Mexican Spadefoots is premature emergence in response to sound vibrations that accompany off-­road vehicle activity in arid habitats. Vibrations from electric motors have been shown to induce 100% emergence from protected burrows (Dimmitt and Ruibal, 1980b). If spadefoots emerged ­under inhospitable conditions due to human-­induced vibrations, they could be exposed to direct harm from vehicles or be unable to rebury themselves in dry desert substrates. ­There are certain trapping biases when sampling for larval S. multiplicata as part of community analyses. Mexican Spadefoots are not readily captured in funnel traps, and are more likely to be captured by seining or dip-­netting (Ghioca and Smith, 2007). Thus, trapping tends to underestimate the proportion of Mexican Spadefoots within a pool. Traps also tend to catch more of the carnivorous morph larvae when compared to the proportion of all morphs within a population. Popu­lar accounts of spadefoot biology are in Bragg (1955) and, especially, Bragg’s (1965) book Gnomes of the Night.

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INTRODUCED SPECIES

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F ­ amily Dendrobatidae

Dendrobates auratus (Girard, 1855) Green and Black Dart-­Poison Frog ETYMOLOGY

auratus: derived from Latin meaning ‘golden.’ The species was described from a population in Panama that has metallic gold spots on a brown background. NOMENCLATURE

Synonyms: Phyllobates auratus. Complete synonymy in Amphibian Species of the World 6.1, an online reference. IDENTIFICATION

Adults. Dendrobates auratus is one of the most beautiful frogs known. It is a small species with a highly variable, bright iridescent green and black coloration. Rarely, some individuals ­will be black and white or purple and white. ­Others may tend ­toward yellow or blue. This frog is small and has a pointed snout and short, delicate limbs. The fin­ger disks are large and ­there is no webbing between the toes. Males are slimmer and have a vocal sac but are other­wise indistinguishable from the females. The normal size range is 25–40 mm SUL for males and 27–42 mm SUL for females (in Savage, 2002). Larvae. Larvae are uniformly colored dark brown to black, including the robust body and tail musculature. No spots or markings are apparent. Snouts are blunt. Tail fins are narrower than the tail musculature. Tails are long and about 1.5 times the length of the body. Maximum size is 30–39 mm TL (but see below). Eggs. The eggs are small and black, with each egg surrounded by a clear gelatinous capsule. The clutch is oviposited on a cushion of clear jelly, which presumably holds moisture and cushions the eggs.

1932 in the upper Manoa Valley and Waiahole Valley on Oahu, ostensibly for the biocontrol of injurious insects (Hunsaker and Breese, 1967; McKeown, 1996; Kraus and Duvall, 2004). The frogs originally released came from Panama. The species occurs only on Oahu in moist leeward and windward valleys. LIFE HISTORY

The Green and Black Dart-­Poison Frog is a terrestrial species of moist and humid leaf litter in shaded forest. They are ­adept at climbing, with reports from Central Amer­i­ca of frogs >13 m above the ground in the tree canopy (Savage, 2002). They may be especially active in the morning and late after­noon, but less so on bright sunny days. They walk or move in short hops, searching for tiny insects. In the Hawaiian winter (November–­April), they are more active than they are during the hottest summer months (May–­October), when they ­will seek refuge ­under debris and sheltering objects. Dendro­ bates auratus eats small insects, mites, and collembolans, with a preference for ants. Sexual maturity is reached in 15 months in Central Amer­i­ca (Savage, 2002). The bright coloration indicates that the frog is highly toxic. Although not particularly dangerous to ­humans, care should be taken when ­handling them. ­People should wash their hands immediately and never touch their eyes or mucous membranes ­after touching one of ­these frogs. It is likely that few predators bother this species. Dendrobates auratus is photopositive in their phototactic response, suggesting that they use sunlight when feeding and ­going

DISTRIBUTION

This species is native to Central Amer­i­ca (Nicaragua to northwestern Colombia). It was introduced to Hawai’i in

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Distribution of Dendrobates auratus

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760  Introduced Species

about their daily activities ( Jaeger and Hailman, 1973). They have a flattened spectral curve that is unclassifiable (Hailman and Jaeger, 1974). BREEDING SITES AND REPRODUCTION

Males call diurnally to attract females, usually in the late after­noon, and they are aggressive in defense of a calling location. Calls are made from holes or hollows at the bases of trees. Both males and females are aggressive ­toward conspecifics. Intruding males are met with a challenge, and likely a wrestling match ­will ensue. The call is a low, slurred buzz (“cheez-­cheez-­cheez”) lasting 2–4 sec, with 3–5 notes followed by a 5 sec pause before the call is repeated. The dominant frequency is 3.5 kHz (Savage, 2002). Females actually court the males ­after they reach the males’ calling position, and they ­will chase other females from the territory of a prospective mate. Although a male can mate with many females, a territorial female ­will attack an intruder female. Courtship lasts several hours, with males and females touching and hopping around one another. Mating occurs in a protected and secluded terrestrial location. Amplexus does not occur. The female deposits her eggs (normally 4–6, but up to 13) in a moist location, and the male then fertilizes them. The female then departs, and the male takes up a position to guard the nest from predators and desiccation. Males ­will guard multiple clutches and continue to call for females ­after mating. Females are capable of laying multiple clutches at 8–10 day intervals throughout the breeding season. The eggs hatch in 10–16 days, and the male then transports the larvae on his back to standing ­water (e.g., shallow pools, treehole cavities) and releases them. The male also may pick up tadpoles and carry them between sites. The tadpoles are omnivorous, but they readily eat other animals, particularly mosquito larvae, daphnia (­water fleas), and drowned insects. For this reason, ­there is usually only 1 tadpole in each small, water-­filled pocket or treehole, inasmuch as larvae ­will readily attack and consume conspecifics. They also eat algae and moss. Tadpoles are active by day but ­will rest in a vertical position at night with the head pointing upward at the ­water’s surface. In Hawai’i, tadpoles normally grow to at least 22 mm TL, but McKeown (1996) reported larvae as large as 45 mm TL. The duration of the larval period is 39–102 days in Central Amer­i­ca. DISEASES, PARASITES, AND MALFORMATIONS

Three species of nematodes have been reported from this species in Hawai’i: Cosmocerca sp., Physaloptera sp., and

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Adult Dendrobates auratus carry­ing tadpoles. Costa Rica. Photo: Brian Kubicki

Adult Dendrobates auratus. Oahu, Hawaii. Photo: Fred Kraus

Acuariid larvae. One species of Acanthocephala is also reported as a cystacanth (Goldberg et al., 2018). Ticks are reported as ectoparasites in Central Amer­i­ca. Neither Bd nor ranavirus was found on this species on the island of Hawai’i (Goodman et al., 2019). IMPACT ON NATIVE SPECIES

This species has a l­imited distribution, and ­there is no evidence it has had a detrimental effect on native Hawaiian fauna. ­There is nothing reported in the lit­er­a­ture on the life history of this species in Hawai’i. Indeed, some information in the primary lit­er­a­ture appears contradictory and may reflect geographic variation in be­hav­ior and life history characteristics.

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F ­ amily Eleutherodactylidae

Eleutherodactylus coqui Thomas, 1966 Coquí ETYMOLOGY

coqui: the name refers to the male’s advertisement call, a loud CO-­QUI (KO-­KEE) IDENTIFICATION

Adults. The ground color of Eleutherodactylus coqui is brown to grayish brown, with the dorsum quite variable in pattern, from uniform to mottled or freckled. A broad, light dorsolateral band may extend down each side with the dorsum dark and the lateral colors much lighter below the band. Some Coquis have a vertebral stripe of varying width extending down the ­middle of the back. A dark line may be pre­sent from the nostril through the eye extending to the front limb. Eyes are large, and the snout is pointed. The thighs are sometimes brown with yellowish-­green mottling and are darker than the body. Venters are variable in the amount of pigmentation, from a light, salt-­and-­pepper pattern to a nearly even, darker granular coloration. Males are smaller than females. In Puerto Rico, males are 29.5– 37 mm SUL, whereas females are 35.5–52 mm SUL (Townsend and Stewart, 1994), but they are smaller in Hawai’i. Males average 27.6–30.8 mm SUL (range 25– 34.6 mm) at Hilo and Kona, Hawai’i, and females 33.2– 36 mm SUL (range 27.8–41 mm) (O’Neill and Beard, 2011; O’Neill et al., 2018). Body size of both sexes increases with elevation and, as such, a decrease in ambient temperature (O’Neill et al., 2018). Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg and froglets hatch as miniature adults. Eggs. Eggs are white and unpigmented and tend to stick together in a clump. They are 3.5 mm in dia­meter and surrounded by a tough egg capsule (4.8–5.0 mm dia­meter with capsule). Egg size is negatively correlated with temperature (O’Neill et al., 2018).

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DISTRIBUTION

This species is native to Puerto Rico. Established populations are reported from Hawai’i (Hawai’i, Kauai, Maui, and Oahu; Kraus et al., 1999), but the frogs have been nearly eradicated from Maui and completely removed from Kauai (Beard et al., 2018). Only 1 population occurs on Maui at Maliko Gulch, but frogs are occasionally heard elsewhere. Individuals have been found infrequently in Florida [Dade and Suwannee counties] (Krysko et al., 2019; Somma, 2021), California, and Louisiana; no breeding populations are known from the mainland. Reports of Eleutherodactylus marticensis introduced to Hawai’i with nursery stock at Kokomo and Omaopio on Maui (Kraus et al., 1999) ­were misidentified E. coqui (Kraus and Campbell, 2002). The Hawaiian populations originated from northern Puerto Rico, based on an analy­sis of genomic DNA (Velo-­ Antón et al., 2007). Two separate introductions (one to Maui and the other to Hawai’i) occurred, and ­there is low ge­ne­tic diversity among populations throughout the islands (Peacock et al., 2009). ­There is no evidence of isolation by distance in the range of the species in Hawai’i, suggesting an extensive mixing among populations via human-­mediated dispersal. Dispersal has been facilitated through the nursery trade, with the first reports of Coquis occurring in the late 1980s (Kraus and Campbell, 2002; Kalnicky et al., 2014), and through vehicle transport around the islands (Everman and Klawinski, 2013). Frogs also have been intentionally released in order to establish new populations. The species is found associated with nurseries, residential areas, resort

Distribution of Eleutherodactylus coqui in Hawaii

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762  Introduced Species

­ otels, ornamental parks, and natu­ral forests. The species is h rapidly expanding its range and, ­because of low ge­ne­tic diversity, a single female with eggs may be able to establish a new population (Beard et al., 2018). LIFE HISTORY

The life history traits of Hawaiian populations are similar to low elevation populations in Puerto Rico (O’Neill et al., 2018). This species is very common and widespread in Puerto Rico, both in humid native forest and in human-­ disturbed areas. Coquis appear to be occupying the same type of habitats in Hawai’i, where populations have been reported from 0 to 1,200 m in elevation (1,158 m in Hawai’i Volcanoes National Park). Populations can be very large. For example, Kraus et al. (1999) reported collecting 105 adult Coquis in 50 minutes from an area 50 m2. Populations are small for about a year following introduction but rapidly increase afterward. Woolbright et al. (2006) reported densities 3 times greater in Hawai’i than in Puerto Rico, with adult estimates ranging from means of 3,413–11,800/ha depending upon population and season. ­There was a mean of 0.7–7.5 juveniles per adult, with ratios again changing seasonally. The life history of this species is reviewed by Rivero (1998) and Joglar (1998). Sex ratios are biased ­toward males, with mark-­recapture estimates ranging from 2.2 to 4.5 males per female (Woolbright et al., 2006). Sexual maturity is reached in about 8 months to 1 yr; populations are capable of rapid turnover, but Coquis may live 4–5 yrs (Rivero, 1998). Coquis are largely opportunistic leaf litter feeders that consume prey in proportion to availability. The diet varies among populations in accordance with invertebrate abundance. Coquis tend to eat more and a greater volume of prey in the morning than at night, but the diet composition is similar (Wallis et al., 2016). Prey consists of spiders, roaches, beetles, collembolans, flies, ants, lepidopteran larvae, termites, amphipods, isopods, mites, and lesser numbers of other invertebrates including native species (Beard and Pitt, 2005; Beard, 2007; Wallis et al., 2016; Smith et al., 2017). Most prey comprise nonnative species, especially ants (30%) and amphipods (22%) (Beard, 2007; Wallis et al., 2016); mites, amphipods, and cockroaches make up > 90% of the diet (Smith et al., 2017). Endemic invertebrates (beetles, springtails, flies) and pest species (mosquitoes, termites) do not make up a significant portion of the diet. In turn, Coquis made up 6.6% of the prey by weight of mongooses (Her­ pestes javanicus) in 1 Hawaiian study, but neither rats (Rattus exulans, R. rattus) nor Cane Toads (Rhinella marina) consumed Coquis (Beard and Pitt, 2006). Smith et al. (2017) speculated that birds would prey on Coquis, but no direct observations ­were provided.

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BREEDING SITES AND REPRODUCTION

Reproduction may occur year-­round, but peaks during the wet season. The call is a very loud (80–90 dB at 0.5 m) “ko-­kee” that is unmistakable and makes this species easy to detect during call surveys (Olson et al., 2012). The ko part of the call establishes territorial bound­aries with other males, whereas the kee is the advertisement call to attract females. Calls may be heard at any time, but most calling occurs from dusk to midnight, with an increase in intensity at dawn. Males call from the ground, short bushes and branches, and well into the tree canopy. Females approach the calling male, who then “leads” her to the area where eggs ­will be deposited. ­There is a certain degree of variation in call characteristics (fundamental frequency of each call syllable and call duration, and the number of calls per minute), which is correlated positively or negatively with elevation (O’Neill and Beard, 2011). Some of the variation can be explained by differences in male body size, which is greater at higher elevations. Amplexus lasts a considerable period of time (to 12 hrs). Fertilization is internal via cloacal apposition, with the male remaining in amplexus as the eggs are extruded (Rivero,

Male Eleutherodactylus coqui with eggs. Hawaii. Photo: Jesse Poulos

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Eleutherodactylus coqui 763

IMPACT ON NATIVE SPECIES

Adult Eleutherodactylus coqui. Hawaii. Photo: Steve Johnson

1998). Eggs are deposited on soil or in leaf axils (see Joglar, 1998) in a moist location in terrestrial litter or ­under surface debris. In Puerto Rico, females deposit 4–6 clutches per year with a mean clutch size of 28 eggs per clutch (range 16–41) (Townsend and Stewart, 1994). O’Neill et al. (2018) gave a clutch size of 30 for Coquis from Hawai’i that oviposited in the laboratory. Clutch size is positively correlated with female body size, and larger clutches are oviposited in the wet season than in the dry season. A female can oviposit multiple clutches at about 8 week intervals, and males can mate with multiple females. Male mating success is not correlated with SUL. The male guards the eggs from predators and desiccation for the 2–3 weeks (17–26 days) required for development. In ­doing this, he ­will position himself directly over the clutch with his arms folded around the eggs ( Joglar, 1998). ­Under laboratory conditions, hatchlings from Hawaiian frogs are 6.8– 7.4 mm SUL, with a size of ca. 7.4–8.2 mm SUL at 1 month, depending on temperature (O’Neill et al., 2018). In Puerto Rico, hatchlings are ca. 5 mm SUL ­under field conditions.

As noted above, populations of E. coqui can be very large in Hawai’i, prob­ably due to a lack of predators and an abundance of retreat sites. Much concern has been expressed about the potential effects of E. coqui as it expands its range into native rainforest and mesic forest at higher elevations. ­These areas have a very diverse insect and spider fauna, and authors have voiced concern that Coquis could impact invertebrate biodiversity (Kraus et al., 1999; Kraus and Campbell, 2002; Beard and Pitt, 2005). Choi and Beard (2012) found that Coquis had significant effects (via predation) on the leaf litter invertebrate community, reducing it by 27%, especially among the Acari (mites and ticks) by 36%. Other taxa ­were affected at par­tic­u­lar sites, but not everywhere. The presence of Coquis actually increased flies by 19% across sites, presumably by providing food via excrement and carcasses. Such changes occur on a landscape scale. Rödder (2009) noted that Coquis could spread into higher elevations on Hawai’i as a result of climate change as well as to many other habitats around the world, including Florida. Increased predation on native insects could have detrimental impacts on Hawai’i’s endangered avifauna, although ­there is ­little overlap in habitats used by endemic birds and Coquis. However, ­there does not appear to be any competition between Coquis and native birds based on isotopic analyses, as ­these species occupy dif­fer­ent trophic positions in the forest (Smith et al., 2017). In studies of leaf production, Sin et al. (2008) found that the presence of Coquis significantly reduced the invertebrate communities, especially aerial species, herbivores, and leaf litter species, in some habitats. This in turn resulted in

DISEASES, PARASITES, AND MALFORMATIONS

The amphibian chytrid fungus Batrachochytrium dendroba­ tidis has been reported from E. coqui on Maui and Hawai’i (Beard and O’Neill, 2005). The infection rate was 2.4% of the frogs tested. The rat lungworm, Angiostrongylus cantonensis, was found in 21 of 24 of the Coquis examined on the island of Hawai’i (Niebuhr et al., 2020).

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Habitat of Eleutherodactylus coqui. Oahu, Hawaii. Photo: Steve Johnson

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764  Introduced Species

lowered herbivory rates, increases in nutrients (NH4, magnesium, nitrogen, phosphorus, potassium), increased leaf production by some plants, and increased decomposition rates in other plants. As such, Coquis could inadvertently make ecosystems more vulnerable to invasive plants by making nutrients more available while at the same time decreasing native invertebrate populations. Hawaiians have often reported the loud advertisement call to be annoying, and the annoyance ­factor plus concerns about decreased property values due to loud frogs may have spurred more efforts at eradication than concern for biodiversity (e.g., Raloff, 2003; Kaiser and Burnett, 2006; Beard et al., 2018). The State of Hawai’i bans the sale of this species and has forbidden intentional introduction since 1998. Kraus and Campbell (2002) advocated removal programs but noted a ­great amount of indifference to the prob­lem, an inability or unwillingness to control introductions, public misperceptions of beneficial aspects, and outright defiant ignorance (Singer and Grismaijer, 2005). In 2012, the state spent $4 million to control the frog.

Many control ideas have been considered, but some are logistically impractical, work only ­under certain environmen­ thers ­were tal conditions, or are effective only in small areas; o not publicly acceptable. Hand capture does appear to be efficient in some small areas (Pitt et al., 2012). Research has been undertaken to test the effects of weak detergents, vegetation management (locally effective -­Kalnicky et al., 2013; Klawinski et al., 2014), hydrated lime (too caustic), spraying 16% citric acid (the only approved chemical method), caffeine application (effective, but not ­adopted ­because of ­human health concerns and public opposition), the use of hot ­water (steam kills frogs and their eggs; useful in nursery control), and even the introduction of frog parasites to eradicate the species (not likely to be successful; Marr et al., 2010) (options reviewed by Pitt et al., 2012; Beard et al., 2018). The National Park Ser­vice has an eradication program at Hawai’i Volcanoes National Park that combines spraying citric acid with the placement of a frog barrier fence (Tavares, 2008). Despite ­these efforts, it appears E. coqui has become another member of Hawai’i’s nonindigenous invaders.

Eleutherodactylus planirostris (Cope, 1862) Green­house Frog

continues laterally but tends to fade out ventrally. The venter is light gray to white and speckled with scattered, dark brown spots, as is the throat and undersides of the thighs. Eyes are sometimes reddish. Mottled frogs are similar in coloration but lack the dorsolateral striping. Instead, the dorsum is a uniform light to dark brown, but often mottled with light brown coloration. Dorsal coloration is not correlated with light intensity or with sex. An unusually colored Eleutherodactylus planirostris (partial albino or “pinto”) was reported from Miami–­Dade County, Florida (Petrovic, 1973). ­There is considerable variation in morph frequency among populations as noted by Goin (1947b), based on frequency counts from throughout the species’ range. In north central Florida, the striped form constituted up to 75% of the patterns observed by Goin (1947b). In south Florida, however, striped individuals comprised only 28.4–44.1% of individuals examined (Duellman and Schwartz, 1958). In Louisiana, the mottled morph is more common than the striped morph (Meshaka et al., 2009c). Coloration appears to result from a ­simple dominant/recessive gene, with the striped morph dominant (Goin, 1947b). Males are smaller than females. For example, in Louisiana, males are 16–25 mm SUL (mean 19.8 mm) and females are 18–29 mm SUL (mean 24.1 mm) (Meshaka et al., 2009c). In the Florida Keys, males are 15–17.5 mm SUL (mean 16.6 mm) and females are 19.5–25 mm SUL (mean 22.6 mm) (Duellman and Schwartz, 1958).

ETYMOLOGY

planirostris: from the Latin prefix plani-­meaning ‘flat’ and rostris meaning ‘nose.’ The name refers to the flat area anterior to the eyes. NOMENCLATURE

Frost et al. (2006a): Euhyas planirostris Fouquette and Dubois (2014): Eleutherodactylus (Euhyas) planirostris Synonyms: Eleutherodactylus ricordii planirostris, Euhyas planirostris, Hylodes planirostris, Lithodytes planirostris Hedges et al. (2008) placed E. planirostris within the Eleutherodactylus (Euhyas) planirostris Species Series. IDENTIFICATION

Adults. This is a small frog with granular skin. ­There are 2 phenotypes of Green­house Frogs, a mottled morph and a striped morph. Striped frogs are light to dark brown, reddish brown, or bronze with a light orange to tan stripe ­running from the nostril dorsolaterally down the body. A triangular area of light orange to tan is pre­sent on top of the head, and a dark bar is pre­sent between the eyes. The brown coloration

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Eleutherodactylus planirostris 765

Aransas County, Texas (Simpson et al., 2019). The species is found throughout the Florida Keys and on St. George Island, Florida (Duellman and Schwartz, 1958; Lazell, 1989; Irwin et al., 2001). Subsequent range expansion has occurred both naturally in Florida and through the nursery trade elsewhere. The Green­house Frog has been reported from Alabama, Georgia, Hawai’i (Hawai’i, Oahu; introduced in 1994), Louisiana, Mississippi, South Carolina, and Texas (Plotkin and Atkinson, 1979; Carey, 1982; Kraus et al., 1999; Winn et al., 1999; Kraus and Campbell, 2002; Dinsmore, 2004; Liner, 2007; Kraus, 2009; Meshaka et al., 2009c; Alix et al., 2014a; Mann et al., 2015; Boundy and Carr, 2017; Lieto and Burke, 2019; Simpson et al., 2019; Selman and Garza, 2020). LIFE HISTORY

Distribution of Eleutherodactylus planirostris on the mainland (upper) and Hawai'i (lower). This species is expanding its range around the northern Gulf Coast.

Larvae. ­There is no free-­swimming larva. The larval period is passed within the egg and froglets hatch as miniature adults. Eggs. The eggs are white with a vitellus ca. 2 mm in dia­meter. ­There is a single jelly envelope 3–4 mm in dia­meter that encloses a narrow layer of fluid, a wide layer of jelly, and a tough jelly envelope on the outside (Goin, 1947b). Just prior to hatching, the eggs are 5–6 mm in dia­meter.

The Green­house Frog is common in humid and mesic habitats throughout Florida and adjacent states in natu­ral, suburban, silvicultural, and agricultural settings (Goin, 1947b; Duellman and Schwartz, 1958; Dodd, 1992; Delis et al., 1996; Enge, 1998a; Surdick, 2005; Alix et al., 2014b; Erwin et al., 2016). This is a species of the leaf litter, although Harper (1935a) recorded it ­under tree bark on wild tamarind trees in south Florida hardwood hammocks, and Fonseca et al. (2020) noted arboreal be­hav­ior to escape rising ­water levels. It hides ­under surface debris, rocks, woodpiles, and rotting logs, and in burrows, leaf mold, and residential gardens within the soil. In cold and dry weather, it occupies burrows in the soil. Similar microhabitats are used in Hawai’i, as are subterranean la­va tubes (Olson and Beard, 2012). In Hawai’i, the species occurs up to 1,115 m in elevation. Activity occurs year-­round, weather permitting, in Florida and Hawai’i. In Louisiana, activity has been reported in all months except December, with most activity from May to October (Meshaka et al., 2009c). Most activity occurs at

DISTRIBUTION

The native range of E. planirostris is the Bahamas, the Cayman Islands, and Cuba. The species was first reported by Cope (1863) from Key West (often incorrectly attributed to Barbour, 1910) and south Florida, where it may have arrived on ships coming from Cuba. It now occurs throughout urban and natu­ral habitats in peninsular Florida, including some islands along the west coast (in north central Florida by 1933, Van Hyning, 1933; Meshaka, 2011b; Dodd and Barichivich, 2017; Mendelson and Kelly, 2017; Krysko et al., 2019), in southern Georgia ( Jensen et al., 2008), and in scattered locations along the Gulf of Mexico (Alix et al., 2014a, 2014b) as far as west of Dallas and southward in

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Adult Eleutherodactylus planirostris, striped pattern. Photo: Alan Cressler

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766  Introduced Species

Adult Eleutherodactylus planirostris, non-­striped pattern. Photo: David Dennis

night, especially during warm weather, but Green­house Frogs may be active diurnally ­after rain and on cloudy days. They are rapid jumpers and are readily able to escape into the leaf litter; however, they are more sluggish in winter at temperatures 120 yrs in the islands.

Neither Bd nor ranavirus was found on this species on the island of Hawai’i (Goodman et al., 2019).

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F ­ amily Hylidae

Litoria caerulea (White, 1790) Green Tree Frog, White’s Green Tree Frog

Wales–­Victoria border. In Florida, individuals have been found along canals in Broward, Collier, Lee, Miami-­Dade, and Sarasota counties (Butterfield et al., 1997; Meshaka et al., 2004; Krysko et al., 2011, 2019).

ETYMOLOGY

LIFE HISTORY

caerulea: from the Latin caeruleus meaning ‘greenish-­blue.’ This species appears bluish green in its native New South Wales, hence its name. IDENTIFICATION

Litoria caerulea is a large, bright green or bluish-­green frog with markedly expanded toe pads, large eyes, and a distinctive thick fold of skin that extends in a curve from the back of the eye at midlevel across the top of a distinct tympanum (hence, a supratympanic fold) ­toward the top of the forelimb. Very small white dots may be pre­sent dorsally. At night, the frog may appear olive green. The snout is distinctly rounded when viewed dorsally. The skin is smooth and moist, and ­there is a prominent parotoid gland above the supratympanic fold. Venters are white to lemon yellow, but pinkish on the undersides of the hind limbs. The webs between the fin­gers are less in extent than ­those between the toes. Males grow to 77 mm SUL and females to 160 mm SUL, but size varies substantially geo­graph­i­cally in Australia. Juveniles are similar to adults except for a dark lateral (canthal) stripe through the eye from the snout across the eye to the forelimb. This stripe dis­appears ­after 2 weeks. Tadpoles are large, from 50 to 68 mm TL. The body is plump, dark brown to olive or golden brown. Eyes are golden. Venters are coppery in color. The dorsal tail fin is translucent with some very small dark stippling. A distinct dark lateral stripe is pre­sent on the tail musculature, bordered above and below by mostly pale muscle. Anstis (2017) provides additional descriptions and color photo­ graphs of all life stages of this species.

Litoria caerulea lives on trees in terrestrial habitats and breeds in ephemeral ponds, wetlands, and ditches. They are often found along watercourses. They are not uncommon in residential and urban habitats, although populations in some of Australia’s cities have declined. Adults are insectivorous, and tadpoles eat algae and other protein rich sources in ­water. BREEDING AND REPRODUCTION

In Australia, males call in large choruses, often in response to rainfall, during the spring and summer (austral November–­ February). Up to 2,000 eggs are deposited in floating clusters, and hatching occurs in 2–4 days depending on ­water temperature. Larvae develop rapidly, and metamorphosis

DISTRIBUTION

This species is native to Australia from the northwest across the top end and as far south as near the New South

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Distribution of Litoria caerulea. The map represents sightings or museum specimens, and may not be indicative of established populations.

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772  Introduced Species

Adult Litoria caerulea. New South Wales, Australia. Photo: Kevin Enge Tadpole of Litoria caerulea. Australia. Photo: Marion Anstis IMPACT ON NATIVE SPECIES

occurs from 27 to 48 days in Australia (Anstis, 2017). However, tadpoles are capable of overwintering if eggs hatch late in the season. Metamorphs are 15–26 mm. DISEASES, PARASITES, AND MALFORMATIONS

Nothing known for Florida.

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Inasmuch as L. caerulea in Florida is not known to have established a breeding population (Krysko et al., 2019), the impact on native species is likely negligible at pre­sent. This species is popu­lar in the pet trade, and individuals have been bred commercially in Florida. Given south Florida’s climate and the presence of this species in urban areas, L. caerulea is likely to become established somewhere in Florida.

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F ­ amily Hylidae

Osteopilus septentrionalis (Duméril and Bibron, 1841) Cuban Treefrog ETYMOLOGY

external nuptial pads by 26 mm SUL (McGarrity and Johnson, 2009). In south Florida, males average 46.1– 52.9 mm SUL, whereas females average 57.3–71.3 SUL, depending upon population. Waddle et al. (2010) reported a mean length of 38.6 mm SUL from southwest Florida. In the Everglades, males averaged 46.1 mm SUL (range 28.9– 59.8 mm), whereas females averaged 64.2 mm SUL (range

septentrionalis: Latin for ‘northern.’ Duméril and Bibron thought that the frog came from Norway. NOMENCLATURE

Synonyms: Hyla dominicensis [in part], H. septentrionalis. Complete synonymy in Amphibian Species of the World 6.1, an online reference. IDENTIFICATION

Adults and juveniles. This is the largest treefrog in Florida, particularly the adult females. The ground color is gray to light brown; some frogs are uniform in coloration, whereas ­others have brown or green mottling or lichen-­like patterns dorsally. The skin is bumpy, and the skin on the head is fused to the skull. This allows Cuban Treefrogs to back into tight places with the head blocking the entrance. Since the skin is fused to the skull, the frog is difficult to ­handle, and the bony skull prevents ­water loss. The eyes and toepads are exceptionally large and con­spic­u­ous. A yellow wash may be pre­sent in the axil of the forelegs. Venters are white to off-­white. The male has bilateral vocal sacs. Juveniles have a pale green to light tan ground color with a broad, dorsolateral, cream to yellow stripe. They are sometimes confused with Dryophytes cinereus, but the expanded toe disks are much larger in O. septentrionalis. In addition, juvenile Cuban Treefrogs often have red eyes, and they have blue bones that are especially evident in the long hind limbs. Females are substantially larger than males, but the sexual size dimorphism decreases with increasing latitude; females tend to be smaller in the north, but male SULs remain about the same. In central Florida, females are 25–79 mm SUL (mean 53 mm), whereas males are 25–62 mm SUL (mean 43.1 mm). Females have eggs by 53 mm SUL and males have

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Leg of juvenile Osteopilus septentrionalis showing blue bones. Photo: Steve Johnson

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774  Introduced Species

44.5–99 mm) (Meshaka, 2001). Meshaka (2001) and Meshaka et al. (2015c) provided additional information on body size for populations in the lower Keys, Palmdale, Lake Placid, Okeechobee, Fakahatchee Strand, and Tampa, Florida. In contrast, Duellman and Schwartz (1958) recorded males averaging 53–57.5 mm SUL and females averaging 71.7–78.6 mm SUL from Miami and Key West, with the largest female mea­sured at 96.5 mm SUL and the largest male at 61.6 mm SUL. Cuban Treefrogs within their native range are larger, with males averaging 44–56.2 mm and females averaging 68.5–77.3 mm SUL. The largest Florida male is 85 mm SUL, and the largest Florida female is 165 mm SUL (McGarrity and Johnson, 2009; Krysko et al., 2019). In Louisiana, males averaged 49.4 mm SUL (range 38–61 mm) and females 54.9 mm SUL (range 40–88 mm) (Glorioso et al., 2018). Larvae. Larvae retain yolk for several days ­after hatching. Mature tadpoles are black dorsally and around the eyes with grayish-­brown tail musculature. This grayish-­brown color extends onto the body laterally. They have a round body and a wide, transparent tail fin that may contain small, brown pigment spots. Venters are light. Tadpoles reach at least 32 mm TL. Duellman and Schwartz (1958) provided a figure of the tadpole and its mouthparts. Eggs. Dark-­colored eggs are oviposited in a surface film (see below). Eggs average 1.2–1.3 mm in dia­meter (range 1–1.6 mm) at oviposition. The outer envelope is 3 mm in dia­meter, the inner envelope is 2.4 mm in dia­meter, and the vitellus is 2 mm in dia­meter within 24 hrs of deposition (Duellman and Schwartz, 1958). Ovum size is not correlated with clutch size or female body size (Meshaka et al., 2015c). DISTRIBUTION

Cuban Treefrogs are native to Cuba, the Cayman Islands, and the Bahamas (e.g., Smith and Iverson, 2016). Florida individuals are derived from 2 locations in Cuba (Heinicke et al., 2011). The first rec­ord in Florida was by Barbour (1931), although the species was observed before 1928 and likely arrived ­earlier in shipping crates. Duellman and Schwartz (1958) speculated that the species could be considered native to Florida. Osteopilus septentrionalis has spread from the Florida Keys throughout peninsular Florida (with primary distribution roughly along a line extending from Duval to Alachua to Levy counties), with individuals reported in the Florida Panhandle (Krysko et al., 2019) and coastal Georgia (Cumberland Island, Brunswick, Savannah). The frogs are established in New Orleans, Lafayette, and north of Lake Pontchartrain in Louisiana (Glorioso et al., 2018; Car­ter et al., 2021) and ­were pre­sent in Southern California as of 2015 (https://­nhm​.­org​/­stories​/­los​-­angeles​

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Distribution of Osteopilus septentrionalis

-­being​-­invaded​-­frogs). Additional reports occur along transportation corridors (Interstate Highways 75 and 95) as far north as Ontario, Mas­sa­chu­setts ( Jarboe et al., 2019), Minnesota, and ­Virginia (Mitchell, 1999), and west to Colorado (Livo et al., 1998). ­There are unconfirmed reports of an established population in the Houston, Texas, area (M. McGarrity, personal communication). Another isolated rec­ord is from Alabama. Other distribution references include: Goin (1944), Trapido (1947), Schwartz (1952), Allen and Neill (1953), Duellman and Schwartz (1958), Austin (1973), Meshaka (2001), Johnson and McGarrity (2010), and Lieto and Burke (2019). Dispersal is aided by the tendency of Cuban Treefrogs to hide in closed spaces such as in car doors and engines, boats, motor homes, and camping trailers, and in nursery stock, for example, within palm trees used for landscaping (Meshaka, 1996b; Herman, 2014; Glorioso et al., 2018). The species occurs in both urbanized and natu­ral habitats. The northward spread of O. septentrionalis ­will likely be ­limited by minimum ambient temperature. However, the species could easily spread along the northern Gulf Coast all the way through southeast Texas. Although Wilson (2010) estimated the CTmin at only 7.8°C for frogs acclimated at 15°C, a more comprehensive study by Simpson (2013) showed that the CTmin was actually -1.01°C for north Florida frogs acclimated at 13°C. In addition, this temperature was less for north Florida frogs than for south Florida frogs, indicating a degree of adaptation to cooler temperatures. As climate warms along the Gulf and Atlantic Coasts, this species can be expected to expand its range.

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Osteopilus septentrionalis 775

LIFE HISTORY

The life history of O. septentrionalis in Florida is reviewed by Meshaka (2001) and Krysko et al. (2019). The Cuban Treefrog is especially common in human-­disturbed habitats such as urban and suburban residential developments and agriculture, including orange groves and nurseries (Surdick, 2005). It also has moved into pine rockland, mesic hardwood forests, scrubby and mesic pine flatwoods, cypress ponds and domes, prairie mangroves, wet prairies, hydric hammocks, bottomland forests, swamps, and even into xeric sandhill Florida habitats (McGarrity and Johnson, 2009; Krysko et al., 2019). Cuban Treefrogs are often pre­sent in ­great abundance. Females grow faster than males and mature ­later. Meshaka (2001) recorded differences in the age and sex structure among populations. Osteopilus septentrionalis is active mostly during warm weather (>15°C), as the CTmin is only 6.4°C; the CTmax is 39°C ( John-­Alder et al., 1988). Freezing temperatures (-4°C) cause mortality, but populations can recover from short-­term freeze events (Haggerty and Crisman, 2015). In the winter during the dry season and in the heat of the day, the frogs take refuge in shelters offering confined spaces and cavities. On Egmont Key, for example, I have found them in arboreal retreats within dead palms and sheltered ­under oil drums. Meshaka (1996c) reported them using bird boxes, PVC pipe, air conditioning units, burrows in sphagnum, drainpipes, tree holes in a variety of species, palm axils, and in bark and sheet metal folds. As with other arboreal hylids, evaporative ­water loss is lower than in more terrestrial and aquatic species (Wygoda, 1984), which likely also aids in human-­mediated dispersal. In natu­ral habitats, Cuban Treefrogs ­favor forested habitats, where they are found most often in tree canopies. If displaced into old-­field habitats, they ­will occupy the shrub and subcan­ ill rapidly move to opy layers if trees are not available but w adjacent forested habitats. Most movements occur at night, when humidity and temperature is high, wind speed is low, and moonlight is subdued. If displaced 200 m to other forested habitats, movements occur more randomly, at least over a short time, with frogs tending to ­settle into the new area. In old-­field to forest movements, frogs averaged 21 m/day (range 11–47 m), whereas within forest, movements only averaged 8 m/day (range 4–19 m) (McGarrity and Johnson, 2010). Cuban Treefrogs eat just about anything they can stuff into their mouths, including insects, spiders, snails, small frogs, lizards, and even snakes. Heflick (2001), for example, found 11 ­orders of prey in O. septentrionalis guts, including beetles, cockroaches, moths, spiders, and the treefrog Dryophytes cinereus. Meshaka (1996a) found roaches to be the main prey from a small Florida sample, but in a much larger sample (Meshaka, 2001), he found 20 ­orders of prey in stomachs,

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including a substantial number of frogs and even lizards, as well as shed skin, stones, and vegetation. Beetles, isopods, lepidopterans, and cockroaches seem particularly favored, with females eating roaches, beetles, earwigs, and spiders, and males and juveniles concentrating on beetles, moths, and spiders. Glorioso et al. (2012) also reported a wide variety of invertebrate prey that varied depending on habitat, with frogs in only 3.5% of the stomachs examined. Other prey includes the Florida Striped Scorpion Centruroides hentzi (Granatosky et al., 2011) and the snake Storeria victa (Maskell et al., 2003). Duellman and Schwartz (1958) and Wilson (2010) also noted beetles and other small invertebrates in the diet. Osteopilus septentrionalis does not necessarily prefer Green Treefrogs (D. cinereus) in prey choice ­trials, but it readily consumes this and other hylids (Wyatt and Forys, 2004). Osteopilus septentrionalis is eaten by ­giant ­water bugs (Lethocerus americanus), conspecifics, turtles (Chelydra serpentina), lizards (Anolis equestris), snakes (Agkistrodon piscivorus, Coluber constrictor, Nerodia clarkii, N. fasciata, Pantherophis alleghaniensis, P. guttatus, Thamnophis

Adult Osteopilus septentrionalis, green phase. Alachua County, Florida. Photo: Dale Marcum

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sauritus, T. sirtalis), and birds (Egretta caerulea, Corvus brachyrynchus, Quiscalus quiscala, Strix varia, Tyto alba) (Meshaka and Ferster, 1995; Meshaka, 2001; Mitchell and Johnston, 2013; Herman et al., 2015; Nikolakis et al., 2016; Krysko et al., 2019). It is also attacked by the mosquito Culex iolambdis (Blosser and Lounibos, 2012). Tadpoles are consumed by turtles (C. serpentina), snakes (N. fasciata, T. sauritus), and birds (E. caerulea). Osteopilus septentriona­ lis blends in well with its background and wedges itself into hiding places, both of which help it avoid detection and predation. If attacked, the frogs kick, scream, inflate their bodies with air, and exude a smelly, sticky secretion that is noxious and prob­ably toxic. Some ­people that ­handle Cuban Treefrogs report a mild allergic reaction (sneezing, runny nose) to the secretions or a burning of the mucous membranes if contacted. Although the secretion may be considered toxic, it does not deter predation by native garter snakes (Thamnophis) (Goetz et al., 2018).

Tadpoles can be observed year-­round in south Florida. Foraging occurs at night and on cloudy days in open ­water. Development occurs in ­water 10–41°C. Growth is rapid, with transformation occurring in ca. 27 days at 25°C. However, metamorphosis can be delayed in cool weather, and Meshaka (2001) reported overwintering from October to March. In cold weather, tadpoles form aggregations and appear to bask. Newly metamorphosed froglets are 10.5– 16 mm TL (Duellman and Schwartz, 1958; Meshaka, 2001). Maturity is reached within 7–9 months of transformation, with males growing slightly faster than females. According to Meshaka (2001), longevity is short, with males usually living only about 1 yr. In experimental ­trials, larval O. septentrionalis had 100% survival at salinities ≤5 ppt. However, no larvae survived at salinities of 14 or 16 ppt over a 72 hr period (Brown and Walls, 2013). Larvae are able to survive exposure to 10–12 ppt at > 80%. The ability of larvae to survive greater exposure to salt than native species may enhance this species’ colonizing ability. BREEDING SITES, REPRODUCTION, AND AGE STRUCTURE

Eggs of Osteopilus septentrionalis. Photo: Steve Johnson

Tadpole of Osteopilus septentrionalis. Alachua County, Florida. Photo: Kevin Enge

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The breeding season is primarily during warm weather and long daylight hours from March to October in peninsular Florida (e.g., Meshaka et al., 2015c), but may be year-­round in extreme south Florida and the Everglades. Although ­there is a distinct testicular cycle in males, they are capable of breeding year-­round, and amplexed pairs have been observed in winter (Meshaka, 2001). Males with nuptial pads can be found in any month of the year. Gravid females are also found year-­round, but most gravid females are observed during the summer wet season. Cuban Treefrogs use just about any standing ­water for breeding, including temporary wetlands, cypress swamps, marshes, wet flatwoods, pools in bottomland forests, urban ­water catchments, retention ponds, garden ponds, swimming pools, roadside ditches, and open containers holding ­water. On Egmont Key, for example, they breed in old sewer system catchments. On Key Vaca, Peterson et al. (1952) reported them breeding in brackish ­water. However, reproduction is not successful in ­water with predaceous fish, although they may breed in the grassy shallows of large permanent ­water bodies containing fish. The species is an explosive breeder, with large numbers breeding si­mul­ta­neously. Calling occurs from the substrate to about 3 m in height on vegetation along the shoreline. They do not float in ­water but ­will call from debris in shallow ­water or from near the shore in very shallow ­water. Males have a loud advertisement call described as a “hoarse mraaaak!” ( Johnson and

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EFFECTS OF STRESSORS

Adult Osteopilus septentrionalis, amplexus. Photo: C.K. Dodd, Jr.

McGarrity, 2010) or “waarh-­waarh” (Lee, 1969b). Calls consist of 2 ele­ments and are described in detail by Blair (1958a); the frequency ranges from 300 to 5,100 cps over a duration of 0.35 sec. In south Florida, calls occur throughout the year, but a­ fter dark only from March to October (Meshaka, 2001). Calls also occur diurnally during heavy thunderstorms or during tropical cyclones. Partial egg clutches are deposited in a large surface film of 200–300 (up to 1,000) eggs at a time, although the total egg complement averages 3,000–16,000 eggs. The film may sink, however, during hard rain. The egg film is sticky, especially ­after deposition. Total clutch size and ovum dia­meter are correlated with female SUL, but the relationship between SUL and ovum size varies among populations. In south Florida, clutches averaged 3,961 eggs (range 1,177–16,371) (Meshaka, 2001) and 3,298 eggs (range 1,814–5,141) (Meshaka et al. (2015c). Hatching occurs in ca. 27.5–30 hrs. Tadpoles are 5.9 mm TL 48 hrs ­after hatching. The larval period is 2 months in north Florida. Tadpoles range in size up to 3.2 cm TL, with metamorphs averaging 1.2–1.7 cm SUL. In the Everglades, females mature in 7–8 months at 4.5 cm SUL, and males at 3 months at 2.7 mm SUL (Meshaka, 2001). Meshaka (2001) reported that wild males only live about 1 yr, whereas females live 2–3 yrs; captives have lived much longer. DISEASES, PARASITES, AND MALFORMATIONS

Parasites of the introduced population in Florida have not yet been examined. Amphibian chytrid (Bd) has been reported from this species in Florida (Wilson, 2010; Hughey et al., 2014), as has Perkinsea, an alveolate pathogen (Karwacki et al., 2018). In New Orleans, ranavirus and Perkinsea have been reported in larvae and postmetamorphs, although Bd was not detected (Galt et al., 2021).

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This species is sensitive to the herbicide atrazine. At concentrations of 10.2 and 50.6 µg/L, corticosterone levels decrease, but at low concentrations (0.1 µg/L) and high concentrations (102 µg/L), levels actually increase (Mc­ Mahon et al., 2017). ­These results suggest that atrazine might dysregulate corticosterone in native frog species. Traffic noise may affect tadpole be­hav­ior since tadpoles detect sound through vibrations in the ­water. In a series of experiments, larvae increased their activity and reduced the amount of food consumed when exposed to real-­world simulated traffic noise, but ­there was no effect on growth rate or subsequent size at metamorphosis (Castaneda et al., 2020). Other potential effects of loud anthropogenic noise are unknown. IMPACT ON NATIVE SPECIES

­ ecause of the Cuban Treefrog’s large size and appetite, B ­there is much concern about its effects on native treefrog populations in Florida. In areas where O. septentrionalis populations are large, both the abundance and survivorship of native species decrease ­after habitats are invaded by O. septentrionalis. In southwest Florida, D. cinereus and D. squirellus ­were much less likely to occur at sites where Cuban Treefrogs ­were pre­sent (Waddle et al., 2010; Schuman et al., 2021), and in the latter study, it appeared clear that invasive Cuban Treefrogs had displaced native species over the period 2005–2017 at both natu­ral and restored sites in protected areas. In central Florida, Hoffman (2007) found a negative correlation between O. septentrionalis abundance and the abundance of D. femoralis. The good news is that removing Cuban Treefrogs results in an increase in native treefrog (D. cinereus, D. squirellus) abundance (Rice et al., 2011). Competition may occur for food resources among postmetamorphs, and Cuban Treefrogs readily eat smaller hylids. For example, Meshaka (2001) reported significant dietary overlap between O. septentrionalis and D. cinereus and D. squirellus. Competition also may occur for breeding and retreat sites, and Cuban Treefrogs may interfere with the reproduction of native species. However, Hoffman (2007) was unable to demonstrate interference competition for retreat sites in laboratory ­trials or inhibition against occupancy by D. cinereus and D. femoralis in refugia previously occupied by O. septentrionalis. Cuban Treefrog tadpoles are superior competitors to some native species, and larvae are known to eat the larvae of other species, such as D. squirellus, at least in captivity (Smith, 2005b). This is of concern, since Cuban Treefrog tadpoles often outnumber the tadpoles of native species ­because of the high fecundity of O. septentrionalis. In

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778  Introduced Species

Osteopilus septentrionalis breeding habitat. Alachua County, Florida. Photo: Marian Griffey

laboratory ­trials, the presence of O. septentrionalis larvae resulted in decreased growth rates and delayed metamorphosis of Anaxyrus terrestris and D. cinereus larvae. In addition, the mass at metamorphosis was less than normal in A. terrestris. Once O. septentrionalis larvae had metamorphosed, the mass of longer-­developing D. cinereus larvae actually increased at metamorphosis (Smith, 2005a). Similar results ­were obtained by Knight et al. (2009), although ­these latter authors could not detect an effect on A. terrestris larval survivorship. ­These results suggest that priority at breeding pools can significantly impact the larval development of Southern Toads. In a series of similar experiments, Knight et al. (2009) could not demonstrate significant impacts by O. septentrio­ nalis on the growth, size, mass, or duration of the larval period of D. squirellus. However, survivorship of D. squire­ llus larvae decreased in both sunny and shaded mesocosms in the presence of Cuban Treefrog larvae. This result occurred particularly in the presence of older Cuban Treefrog larvae, but not young larvae, again implicating a priority effect of the Cuban Treefrog larvae on larval native species. Interspecific competition appears to have a much stronger effect on this species than it does for A. terrestris, perhaps ­because the larval period of A. terrestris is so much shorter than ­either O. septentrionalis or D. squirellus (Knight et al., 2009).

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In south Florida, the presence of Osteopilus ameliorates interactions between the exotic geckos Hemidactylus mabouia and H. garnotii. With Osteopilus absent, H. mabouia outcompetes H. garnotii on buildings, but with Osteopilus pre­sent, ­these species reach a stabile equilibrium allowing the species to coexist. The Cuban Treefrog is a better competitor around lights than H. mabouia, whereas the be­hav­ior (particularly in wariness and flight distance) of H. garnotii makes it less susceptible to predation by the treefrog (Meshaka et al., 2020b). Although Cuban Treefrogs are eaten by native garter snakes, the snakes do not appear to metabolize the species very effectively and do not gain weight in the same proportion they would by eating native treefrogs. This creates a potential evolutionary trap, whereby snakes essentially waste energy in pursuing and eating a substandard prey, which in turn reduces their fitness (Goetz et al., 2018). Temperature ­will likely play an impor­tant role in determin­ ill disperse in the southern ing how far O. septentrionalis w United States. In addition, the small female size in northern dispersers suggests decreased fecundity among colonizing females compared with ­those farther south (McGarrity and Johnson, 2009). Smaller female size could also mean less probability of large females having a significant impact on native hylids through predation or competition. Goetz et al. (2017) have also shown that the immune response of invading frogs at the northern limit of their range in Florida is not as effective as that of frogs that have been established in southern Florida for ca. 70 yrs. Taken together, ­these ­factors may restrict expansion to some extent in northern populations. It may be impossible to completely eliminate Cuban Treefrogs from Florida’s fauna. The University of Florida Extension Ser­vice recommends euthanizing Cuban Treefrogs using 20% benzocaine. An alternative humane method of euthanizing Cuban Treefrogs is to cool and then freeze them (Lillywhite et al., 2017; Shine et al., 2019). Johnson et al. (2010) reported that a product called “Stiff ‘n’ Stop” (IFOAM Specialty Products, Sanford, Florida) is effective at deterring Cuban Treefrogs from taking up residence on buildings and in electrical switchgear boxes. Avoidance is based on time-­release of the chemical isophorone, which many animals find unpleasant. This is impor­tant, since electrical outages caused by Cuban Treefrogs are common in south Florida, causing thousands of dollars in damage. In real­ity, Cuban Treefrogs likely are ­here to stay.

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F ­ amily Pipidae

Xenopus laevis (Daudin, 1802) African Clawed Frog

laevis: from Latin meaning ‘smooth.’

are absent. Viscera are vis­i­ble through the skin. In high concentrations of algae, the tadpole can take on a green appearance. Eyes are black. Larvae have a very long tail with a whip-­like tip and high tail fin. Maximum length is 80 mm. Eggs. The eggs are pale brown and very small, averaging 1.1–1.2 mm in dia­meter inside a firm capsule 1.6 mm in dia­meter. Hatching occurs in 24 hrs.

NOMENCLATURE

DISTRIBUTION

ETYMOLOGY

Synonyms: Complete synonymies in Amphibian Species of the World 6.0, an online reference, and Fouquette and Dubois (2014). IDENTIFICATION

Adults. This species is streamlined for living in the w ­ ater, with a narrow, dorsoventrally flattened head and smooth, globular, gray body. The ground color is olive to brown, finely spotted to marbled or with large, round or irregular spots in yellow or dark tints. A lateral line system is obvious on the sides of the body, looking like a line of stitches. The eyes face upward. ­There are no eyelids, nor does it possess a tongue or vis­i­ble tympanum. The front legs are small in relation to the rear legs; front toes are not webbed. The hind legs are long and muscular, and the hind toes are fully webbed with claws on the tips, hence its common name. Venters are immaculate yellowish white to densely spotted. The body is covered with mucous, making it difficult to ­handle. Males have black, roughened areas on the ventral surfaces of the forelimbs during the breeding season; females have large vent lips, whereas ­those of males are barely discernible. Females are larger than males, normally averaging 90– 100 mm SUL, whereas males average 60–80 mm SUL. In California, McCoid and Fritts (1989, 1995) reported males to 80 mm SUL and females 65–119 mm SUL. Larvae. The larvae are easily recognized by their tentacles and transparent body. They actually look more like young catfish than tadpoles. Heads are flattened and nearly translucent, and the body is almost completely transparent with only slight indications of pigment (transparent in daylight and darker at night). The oral disc and keratinized mouthparts

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The African Clawed Frog is indigenous to central and southern Africa, from Uganda and adjacent Sudan to South Africa. A variety of subspecies are recognized, with ­those in the United States most likely X. l. laevis. The species was introduced into the United States in large numbers beginning in the 1930s to 1940s in connection with ­human pregnancy tests. ­Others ­were sold as pets. In 1970–1971, more than 200,000 African Clawed Frogs ­were imported into the United States. However, this number increased to ca. 1.83 million from 1999 to 2015, most of which originated from Asian frog farms rather than from Africa (Measey, 2017). Populations ­were first reported in California in 1968 (St. Amant and Hoover, 1969; Mahrdt and Knefler, 1972, 1973; St. Amant et al., 1973; Bury and Luckenbach, 1976; Lemm, 2006; Measey et al., 2012), with periodic accounts of African Clawed Frogs being captured elsewhere across the continent (Colorado, Mas­sa­chu­setts, Nevada, New Mexico, North Carolina, Texas, Utah, ­Virginia, Wisconsin, Wyoming; Zell, 1986; in Tinsley and Kobel, 1996; Kraus, 2009; Measey et al., 2012). They are now established in most drainages of southern California, near San Francisco, in several cities in Washington state (Bothell, Issaquah, Lacey), and in the Tucson and Phoenix areas of Arizona (Brennan and Holycross, 2006; Measey et al., 2012; Murphy, 2019; Flaxington, 2021). The presence of Clawed Frogs has been confirmed in Florida in Brevard and Miami-­Dade counties (Krysko et al., 2011, 2016, 2019), although breeding populations have not been reliably confirmed. Previous reports of a breeding population of this species in Hillsborough County (Krysko et al., 2016, 2019; Hill et al., 2017) actually refer to X. tropicalis, a West African species (Goodman et al., 2021).

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Tadpole of Xenopus laevis, dorsal view. Photo: Chris Brown

Tadpole of Xenopus laevis, lateral view. Photo: Chris Brown

Distribution of Xenopus laevis in the western United States.

Distribution of Xenopus laevis and X. tropicalis (star) in Florida.

No other breeding populations have been confirmed in the United States. McCoid et al. (1993) reported on the apparent extirpation of a number of populations in California, possibly due to prolonged drought. LIFE HISTORY

Xenopus laevis is an entirely aquatic species with a highly developed sense of smell and an ability to remove oxygen from ­water via the skin. As such, it can remain submerged for long periods. The short front limbs help push food into the mouth and circulate ­water across the lateral line system, which detects both vibrations and chemicals in the ­water. The rear legs are used for locomotion. In its native range, it occupies ponds and lakes in a wide variety of climatic conditions, from low tropics to high mountains (to 2,300 m).

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Feral populations in southern California inhabit both lentic and lotic ­waters, especially man-­made impoundments and canals in highly disturbed areas. The biology of Xenopus is reviewed by Tinsley and Kobel (1996) and du Preez and Carruthers (2009). Populations of African Clawed Frogs can be small to large. For example, McCoid and Fritts (1980b) estimated population size at 2 ponds as 602 and 494 adult frogs. Dispersal may occur downstream within a stream system or during sheet-­ flooding events, but most dispersal is prob­ably ­human mediated. African Clawed Frogs can move overland to the next available ­water when ponds dry (Measey, 2016), but normally they do not leave the ­water. Frogs in drying ponds are capable of excavating deeper holes to maintain ­water longer and to lower the surface ­water temperature (McCoid and Fritts, 1980b). Activity occurs year-­round, although aestivation can be induced in the laboratory. When aestivating, frogs maintain a heads-up position within a small vertical chamber that they dig; the chamber retains a connection to the surface of the pond bottom, which allows them to breathe. Feeding occurs entirely in ­water, and adults capture a wide variety of prey and are known to scavenge. Juveniles eat crustaceans and mosquito larvae, whereas adults eat about anything they can shove into their mouths. Lenaker (1972) reported mosquito larvae, dragonfly larvae, aquatic beetles and their larvae, earthworms, leeches, sow bugs, snails, a moth, the fish Gambusia, toad tadpoles (Anaxyrus sp.), and unidentified adult amphibians in stomach contents from southern California frogs. McCoid and Fritts (1980a) reported the diet as consisting of slow-­moving invertebrates, including amphipods, cladocerans, copepods, ostracods, vari­ous aquatic insect larvae, spiders, and snails. Lafferty

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Adult Xenopus laevis. Photo: Ronn Altig

and Page (1997) reported predation on the Tidewater Goby (Eucyclogobius newberryi) and an Arroyo Chub (Gila orcutti). Xenopus laevis also eats conspecific eggs and tadpoles as well as crayfish. Growth rates are unequal, with females growing faster than males and growth occurring even in winter (McCoid and Fritts, 1995). Sexual maturity in California is reported at 6 months following metamorphosis for females and at 34 months for males (McCoid and Fritts, 1989). In its native range, maturity normally is reached in 2 yrs, although in some instances it occurs much faster. Sex ratios are 1:1. Xenopus can tolerate salinity up to 14‰ (40% sea ­water) for a period of several days and have been found in brackish ­water habitats in California (Munsey, 1972). They occur in ­water over a wide range of temperatures (7.2–32.2°C). The tadpoles are obligate midwater suspension feeders with an efficient entrapment pro­cess for securing food. They do this by secreting mucous on the gills of the buccal cavity, which traps phyto-­and zooplankton. They gulp ­water for this and for respiratory purposes. Tadpoles are sedentary at first, hovering in ­water almost perpendicular to the surface. As they grow, they tend to move around more, often forming large midwater schools and coming to the surface periodically to gulp air. Their long tentacles are extremely sensitive to vibrations in the ­water and alert them to pos­si­ble predators. The preferred ­water temperature is ca. 22°C. As with many other amphibians, X. laevis tadpoles are sensitive to the herbicide atrazine, which significantly affects gonad development (Tavera-­Mendoza et al., 2002).

been reported in all months except December, but most often from March to June (Lenaker, 1972; McCoid, 1985; McCoid and Fritts, 1989, 1993, 1995). It is pos­si­ble that reproduction could occur year-­round. Mature females have large ovarian masses throughout the year, and dif­fer­ent sized tadpoles indicate nonsynchronized breeding over an extended period. Calling and mating occur at dusk or ­after dark, although occasional calls may be heard by day (McCoid, 1985). The male approaches a nearby female and gives chase. If amplexed, the pair swims to vegetation sites, at which time their intertwined feet pump 2–10 times in 2–3 sec as the female ejects eggs and the male fertilizes them. Amplexed pairs remain together throughout the night, periodically depositing eggs in clumps of vegetation. Nonresponsive females have a “rigor stance,” whereby she becomes totally catatonic, and the male soon releases her (McCoid, 1985). Calling occurs from underwater. This species has a complex vocal repertoire involving advertisement calls, intermale communication to establish dominance, and female response calls (Tobias et al., 2004). The advertisement call is a series of very long trills (also likened to a buzz or undulating snore), the first part of which is higher in pulse rate and in the number of pulses; the second part is more rattling. The fundamental frequency is 2 kHz with a mean of 54 pulses/sec, 20–37 pulses/ note, 1–2 notes per sec, and a trill duration lasting 6–28 sec. Eggs are laid singly or in small clusters of 8–10 eggs, each firmly attached to vegetation or other underwater debris. A female can deposit 15,000 eggs or more in a single breeding season, but ­these are oviposited on multiple occasions. McCoid and Fritts (1989) gave egg counts of 2,700–17,000, with the number of eggs correlated with female body size. It is likely that reproductive output is much less per bout (1,000–3,500 eggs) than the number of ova counted within a female at any one time (McCoid and Fritts, 1995). Reproductive bouts can occur at about 1 month intervals. Males are always active and ready to breed. In captivity, hatching success is 74–76%. In South Africa, metamorphosis requires 49–64 days. DISEASES, PARASITES, AND MALFORMATIONS

In California, X. laevis is parasitized by the protozoans Balantidium sp., Nyctotherus sp., and Protoopalina xenopo­ dus; the trematodes Clinostomum sp., Gyrdicotylus gallieni, and Protopolystoma xenopodus; the cestode Cephalochla­ mys namaquensis; the nematodes Contracaecum sp. and Eustrongylides sp.; and the acanthocephalan Acanthocepha­ lus sp. (Lafferty and Page, 1997; Kuperman et al., 2004). Not surprisingly, several of ­these have an African origin.

BREEDING SITES AND REPRODUCTION

IMPACT ON NATIVE SPECIES

Breeding occurs in permanent streams and ponds and continues over an extended period. In California, breeding has

Xenopus laevis could eat substantial numbers of the larvae of native frogs, as they are reported to have a voracious

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782  Introduced Species

Habitat of Xenopus laevis in southern California. Photo: Chris Brown

appetite. ­There has been much speculation about potential negative impacts (e.g., Branson, 1975) but ­little empirical data. McCoid and Fritts (1980a, 1980b) suggested that Xenopus could be a nuisance but not a significant threat to native species. However, Lafferty and Page (1997) reported predation on the endangered Tidewater Goby (Eucy­ clogobius newberryi) and recommended trapping to remove the frog. ­There has been speculation that the virulent

Xenopus tropicalis (Gray, 1864) Tropical Clawed Frog ETYMOLOGY

tropicalis: Latin, referring to the tropical distribution of this species. NOMENCLATURE

Synonyms: Xenopus calcaratus [in part], Silurana tropicalis, Silurana intertropicalis, Xenopus (Silurana) tropicalis IDENTIFICATION

Adults. This species is streamlined (flattened) for living in the ­water, with a narrow, dorsoventrally flattened head and a finely granular, globular, gray or brown body. ­There are small, globular, unpigmented dermal tubercles located linearly dorsolaterally from ­behind the eye to the rear of the body. ­These compose the lateral line system, a sense organ. The ground color is light to dark brown, with fine gray or black spots. The eyes and nostrils are small and face upward; the iris is yellowish. ­There are no eyelids, but ­there

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amphibian chytrid fungus (Batrachochytrium dendrobati­ dis) originated in Africa and has been spread worldwide via trade in X. laevis for medical research and the pet trade (Weldon et al., 2004; Soto-­Azat et al., 2010). This hypothesis is not universally accepted, although it is clear that chytrid has a long evolutionary history in southern Africa. ­There is some concern that the quick-­developing X. tropi­ calis may replace X. laevis in biomedical research and that existing X. laevis colonies may simply be dumped into nearby ­water bodies. Larvae and postmetamorphs are eaten by the Two-­striped Garter Snake Thamnophis hammondii (Ervin and Fisher, 2001, 2007) and Black-­crowned Night Herons (Crayon and Hothem, 1998) in California. The noxious skin secretions of X. laevis induce gaping be­hav­ior in some snakes, which presumably allows the frog to escape (Zielinski and Barthalmus, 1989). However, T. hammondii readily eats African Clawed Frogs with no apparent ill effects, nor do the secretions produce any effects on locomotor per­for­mance (Foster and Mullin, 2008). It is illegal to own, transport, or sell X. laevis in Arizona, California, Hawai’i, Kentucky, Louisiana, Nevada, New Jersey, North Carolina, Oregon, ­Virginia, and Washington, but it is ­legal to own X. laevis in New Brunswick, Canada.

is a small tentacle below the eye. The frog does not possess a tongue or vis­i­ble tympanum. ­There are 4 narrow fin­gers on the front feet and 5 toes on the rear feet. The front legs are small in relation to the power­ful rear legs; the front fin­gers are not webbed. The hind legs are muscular, and the hind toes are nearly fully webbed with 3 of 5 toes with black claws on the tips and another claw on the prehallux. Venters are whitish to yellowish and vaguely mottled black. Black spots may become numerous on the back legs. Breeding males have blackened areas on the outer parts of their fin­gers, whereas the dermal lobes above the vent are better developed in females. In western Africa, females are larger than males, normally averaging 48–55 mm SUL, whereas males average 32–39 mm SUL. Based on 34 museum specimens, males averaged 42.1 mm SUL (to 44.4 mm) and females averaged 50.6 mm SUL (to 59.4 mm) (Evans et al., 2015). This species is diploid in chromosome number (2n = 20) rather than tetraploid as in X. laevis. Evans et al. (2015) noted that clawed frogs of the subgenus Silurana are extremely similar morphologically to one another and may be differentiated only through molecular and internal morphological analy­sis.

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Xenopus tropicalis 783

Larvae. The larvae are easily recognized by their tentacles (barbels), which are long and thin, arising from the upper lip on both sides of the body and extending to the dorsal fin. Tadpoles actually look more like young catfish than tadpoles. The head is flattened. In clear-­water forest pools, larvae are generally pink to orange to off-­white with almost transparent tail fins whose lower margins are often black. In darker habitats, the fin is blackish. The ventral tail fin is much broader than the dorsal tail fin. The oral disc and keratinized mouthparts are absent. Dark pigment surrounds the nostrils, and the area surrounding the mouth is speckled. Paired spiracular openings are located ventrally. Larvae ­were 12–43 mm TL in a random sample of 250 tadpoles in Florida (Hall et al., 2017). Maximum total length is ca. 44–60 mm in Africa (Rödel, 2000). Eggs. The eggs are pale brown and very small. Hatching occurs rapidly. Single eggs are oviposited on stems and leaves of aquatic plants.

Tadpoles of Xenopus tropicalis. Photo: Piotr Naskrecki

DISTRIBUTION

The Tropical Clawed Frog is indigenous to West Africa, from Senegal to Cameroon. Populations of Tropical Clawed Frogs have been confirmed at 26 sites near Riverview, Hillsborough County, Florida, with definite reproduction at 12 sites (Goodman et al., 2021). This species was misidentified as X. laevis in previous publications (Krysko et al., 2016, 2019; Hill et al., 2017). Xenopus tropicalis is widely used in the biomedical and pet trade, and introduced populations likely resulted from escapees from a nearby animal import fa­cil­i­ty. LIFE HISTORY

Xenopus tropicalis appears to be an almost entirely aquatic species. In its native range, the species occupies slow-­ moving streams and both temporary and permanent puddles and wetlands, where they hide ­under nearby debris in times of drought. or. However, they are capable of

Adult Xenopus tropicalis. Photo: Colin Goodman

Distribution of Xenopus laevis and X. tropicalis (star) in Florida.

moving overland to adjacent wetlands and in times of drought (Measey, 2016). The Florida population occupies retention ponds with emergent vegetation. Forested habitats are preferred. The species is a generalist, consuming aquatic invertebrates, earthworms, and tadpoles. As in other clawed frogs, the short front limbs help push food into the mouth and circulate ­water across the lateral line system, which detects both vibrations and chemicals in the ­water. The rear legs are used for locomotion. Nothing is known of its life history in Florida, and much of what is reported in the lit­er­a­ture within its native range is contradictory and may be based in misidentification (see review in Rödel, 2000).

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784  Introduced Species

Africa, breeding occurs in large, clear, temporary pools with abundant aquatic vegetation, and development is rapid. However, other reports have tadpoles inhabiting muddy pools with ­little aquatic vegetation. In Florida, breeding occurs in small, temporary pools and ponds. Pools with sun are preferred. Tadpoles are gregarious and form large aggregations; Hall et al. (2017) provided a crude estimate of 78.6 tadpoles/m2 in a Florida retention pond. Metamorphosis occurs in 2–4 weeks. Recent metamorphs are 11.5– 15.5 mm SUL. DISEASES, PARASITES, AND MALFORMATIONS

Nothing is currently known from the Florida population. Habitat of Xenopus tropicalis in Florida. Photo: Colin Goodman BREEDING SITES AND REPRODUCTION

The breeding call is a low, vibrating, long trill lasting 1–10.5 sec. The dominant frequency is ca. 560–590 Hz; they also have a biphasic call (Tobias et al., 2011). In West

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IMPACT ON NATIVE SPECIES

Nothing is known at pre­sent concerning the effects of this species in its introduced environment. The rapid generation time (maturity is reached in as ­little as 4 months) and its penchant for warm tropical-­like habitats suggest that it could spread throughout much of the Florida peninsula.

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Glossary

Brief definitions of some of the more commonly used words within the text are listed below. For a more comprehensive dictionary of herpetological terms, see H.B. Lillywhite (2008) Dictionary of Herpetology (Krieger Publishing Com­pany, Malabar, FL). Acuminate

Tapering to a point

Aerobic

With oxygen

a.e.

Acid equivalent. The portion of a formulation (as in the case of 2,4-­D for example) that theoretically could be converted back to the corresponding or parent acid

Allele

One of a series of pos­si­ble forms of a gene

Allelochemical

A chemical produced by one organism that has a detrimental effect on another organism; frequently used in reference to plant chemicals that have an effect on other plants and animals

Boss

The raised bony area between the eyes in spadefoots

Buccal pumping

Method of moving air via positive pressure into the lungs by rhythmic movements of the throat or floor of the mouth

Celestial cues

Cues based on the sun, moon, and stars

Chromatophore

A skin cell containing a color pigment

Clade

An evolutionary lineage reflecting monophyly of its derived taxa

Conspecific

Consisting of the same species

Coprophagy

Feces eating. Tadpoles may consume the feces of conspecifics

COSEWIC

Committee on the Status of Endangered Wildlife in Canada

Cryoprotectant

A substance that can be used to depress the freezing point of intracellular fluids and thus allow survival in very cold weather; most amphibians use glucose as a cryoprotectant, although some use glycerol

Cuspate

Possessing serrations

Allozyme

Allelic (dif­fer­ent) forms of enzymes that can be identified using a procedure called electrophoresis

Amelanic

Lacking the pigment melanin that results in black or brown coloration

Amelia

Birth defect involving a missing limb; also a deformed or shrunken limb

Detritivore

An animal feeding on a wide variety of organic plant and animal debris (detritis)

Anaerobic

Without oxygen

Dextral

To the right, on the right side

Animal pole

The dorsal portion of a frog’s egg, usually black or brown in coloration

Distil

Farthest from [the body]

Drink patch

Anosmic

Inability to smell

Anthrogenic

Resulting from ­human activities

Aposematic

Characters that convey warning that the animal may be dangerous, poisonous, or distasteful

An area of high vascularization on the abdomen and ventral part of the thighs that allows ­water uptake directly through the skin. The drink patch is often red in appearance

Ecotone

The abrupt border between two habitat types, such as between a field and forest

Ectromely

A missing limb

Assortive mating

Nonrandom pairing of males and females in reproduction, for example, large females preferably mating with large males

Edema

Swelling due to fluid accumulation

Barrancas

Gully or ravine in the desert Southwest

EPA

U.S. Environmental Protection Agency

Behavioral fever

Raising the body temperature by basking in the sun in response to disease or infection

ESA

United States Endangered Species Act of 1973, as amended

Bicornuate

Literally means having two horns. In the ­Family Ranidae, it refers to the posteriorly bilobed tongue of the frog

Evolution

Organic genet­ically based change through natu­ral se­lection. Evolution is the fundamental unifying princi­ple of biology.

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786  Glossary

Mole

Gape ­limited ­Limited by the extent to which the mouth can be opened

M, a unit of mea­sure­ment of the concentration of a chemical, in par­tic­u­lar of a solute (a dissolved substance) in a solution (a liquid), in terms of the amount of substance per unit volume of solution. For example, the concentration of salt in one liter of ­water. 100% seawater is 35% NaCl, or 600 mM/L

Monophyly

Evolved from a single ancestor

Geotaxis

Myrmecophagy

Ant eating

NaCl

Sodium chloride. Salt.

Nearctic

Biogeographic realm consisting of North Amer­i­ca north of the Isthmus of Tehuantepec

Extirpation

Localized extinction or disappearance of a species

Fossorial

Living beneath the ground

FV-3

One of the six species of viruses in the genus Ranavirus (­Family Iridoviridae)

Directed response to gravity. Geotaxes can be positive (moving ­toward) or negative (moving away from) a source, such as by ­going up or down a slope.

Gosner stages

Graded stages of embryo and tadpole development. See Gosner (1960)

Heliothermic

Controlling body temperature by basking

Necrogamy

Mating with a dead individual

Heliotropic

Moving ­toward (positive) or away from (negative) a heat source

Nekton

Animals within the ­water column capable of in­de­pen­dent movement

Herpetology

The study of amphibians and reptiles

Nestedness

Heterospecific

Consisting of dif­fer­ent species

A mea­sure of interconnectedness among species within a community or region

Hydroperiod

The amount of time a body of ­water holds ­water

Neuston

Biotic community at the ­water’s surface

Hypoxic

Reduced or inadequate oxygen

Operational sex ratio The sex ratio of breeding adults Operculum

Intercalary cartilage A segment of cartilage between the ultimate and penultimate phalanges in the fin­gers of certain frog species

The covering over the gill chamber of tadpoles

Paedomorphosis

Ability to reproduce while retaining larval characteristics

Introgression

Incorporating the genes of one species into the genes of another species via hybridization

Panmictic

A population where mating is completely random

Paraphyletic

Iridophores

A chromatophore containing guanine. Guanine results in a silvery or white coloration or in reflectance.

A taxonomic lineage consisting of two or more species and all their common ancestors, but not all the descendants of the common ancestor

Irvingtonian

North American Land Mammal Age corresponding to 1.9 to 0.15 my BP

Parotoid

IUCN

World Conservation Union, known formerly as the International Union for the Conservation of Nature and Natu­ral Resources

Specialized modified granular (mucous) gland that secretes a toxic or noxious substance to deter predators

PCR

Karyotype

The number and form of the chromosomes

Polymerase chain reaction. A method to rapidly make millions to billions of copies of a specific DNA sample, allowing biologists to take a very small sample of DNA and amplify it to a large enough amount to study in detail

Littoral

Area in shallow ­water adjacent to the shoreline

Pelvic patch

Loess

Fine-­grained soil deposits resulting from wind-­blown silt and clay

A highly vascularized area on a frog’s belly just anterior to the sacrum. This area is pressed to moist soil, and allows the frog to uptake ­water and thus rehydrate

Melanophores

Chromatophore containing melanin, resulting in dark (brown and black) coloration

Periphyton

Metacecariae

The encysted stage of a fluke larva that can produce developmental abnormalities or infection

A complex mixture of algae, cyanobacteria, microbes, and detritus that is attached to submerged surfaces in most aquatic ecosystems

PFAS

Metachrosis

Change in color; the ability to change color

Per-­and polyfluoroalkyl substances. ­These substances are widely used in manufacturing. See following.

Miocene

Geologic epoch extending roughly from 5.3–23 mya

PFHxS

Perfluorohexane sulfonate

PFOA

Perfluorooctanoic acid

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Glossary 787

PFOS

Perfluorooctanesulfonic acid

Phagocytosis

The pro­cess whereby a cell engulfs foreign material

Phenotype

Physical characters of an organism that are observable

Rancholabrean

North American Land Mammal Age corresponding to 150,000 to 10,000  years BP

Recent

Geologic epoch from 12,000 years ago to pre­sent

Phenotypic plasticity Ability to alter the phenotype (morphology, color, life history) in response to environmental or biotic interactions

Riparian

Habitats along the margins of flowing rivers, streams, and creeks

Ruderal

Habitats disturbed by ­human activity

Philopatric

Remaining in the same location where development was completed; literally ‘staying at home’

Silviculture

Photopositive

Orientation ­toward a light source

Management related to the development and use of forests for commercial purposes; associated widely with tree farming

Phototaxis

Moving ­toward (positive) or away from (negative) a light source

Sinistral

To the left, on the left side

Spiracle

Pertaining to the surface features of the habitat

The outside opening to the tadpole’s gill chamber

Subacuminate

Tapering to a point on the lower side

Concentration of osmotically active solutes within the plasma

Sun compass orientation

Geologic epoch from approximately 12,000–2.6 mya

Teratogenic

A firm jelly egg mass that is somewhat rectangular

Refers to an agent causing physical defects in a developing embryo

Thigmotactic

Geologic epoch extending roughly from 2.6–5.3 mya

Crevice-­dwelling or wedging into tight spaces

Titi

A shrubby tree of the Southeast; Cyrilla racemiflora

Toad

A frog in the ­Family Bufonidae; all toads are frogs, but not all frogs are toads

Trophic

Relating to resource (food) use

Tympanum

The sheath of skin covering the opening to the ­middle ear in frogs

USDI

United States Department of Interior

Vegetal pole

The ventral portion of a frog’s egg, usually white or cream colored; contains the yolk for the developing embryo

Vicariance

Speciation due to geographic barriers or isolation

Vitellus

The egg exclusive of its membranes

Xanthophores

Chromatophores containing yellow or orange pigments

Y-­axis orientation

See ‘Sun compass orientation’

Physiographic Plasma osmolality Pleistocene Plinth Pliocene Ploidy

Poikilotherm

The number of sets of chromosomes; haploid is one, diploid is two, and polyploid is more than two Refers to organisms whose body temperature is dependent on ambient temperature; such organisms do not generate a constant body temperature through metabolism

Polyphenic

Having multiple observable patterns, that is, dif­fer­ent phenotypes within a population

Polyploidy

Condition whereby the number of sets of chromosomes is greater than two. Most polyploids are triploids (3) or tetraploids (4)

Proximal

Closer to [the body]

Ranavirus

As a proper noun, a genus of viruses in the ­Family Iridoviridae, i.e., Ranavirus. It is frequently used as a common noun (ranavirus) to include reference to all viruses included in this genus

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The ability to use the position of the sun to orient perpendicularly to the shoreline

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Bibliography

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Zylstra, E.R., and M.K. Ward. 2013. Hyla arenicolor (Canyon Treefrog). Habitat use. Herpetological Review 44:494-495. Zylstra, E.R., R.J. Steidl, D.E. Swann, and K. Ratzlaff. 2015. Hydrologic variability governs population dynamics of a vulnerable amphibian in an arid environment. PLOS One 10:e0125670. Zylstra, E.R., D.E. Swan, B.R. Hossack, E. Muths, and R.J. Steidl. 2019a. Drought-­mediated extinction of an arid-­land amphibian: insights from a spatially explicit dynamic occupancy model. Ecological Applications 29:e01859. Zylstra, E.R., D.E. Swann, and R.J. Steidl. 2019b. Surface-­water availability governs survival of an amphibian in arid mountain streams. Freshwater Biology 64:164–174.

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Index to Scientific and Common Names

Acris, xiii, xiv, xviii, xx, xxiii, xxiv, xxix, xxx barbouri, 215 blanchardi, 84, 193–210, 212, 214, 216, 218–219, 283, 320, 340, 372, 380, 384, 463, 496 crepitans, 193–198, 201, 203, 205–215, 218–219, 236, 283, 340, 372, 463, 640 gryllus, xxiii, 140, 193, 195–196, 206–208, 211–221, 514, 519, 588 g. dorsalis, 215 g. paludicola, 193, 195 Aeromonas, 34, 42, 57, 59, 71, 80, 166, 368, 436, 481, 500, 558, 575, 622, 692 African Clawed Frog, xv, 779–782 alvarius, Incilius, 46, 74, 79, 134, 160, 167–172, 713 Amargosa Toad, 124–127 Amerana, 453, 647–648, 656–657, 664, 670, 672, 678–679, 688–689, 693, 695, 702 American Bullfrog, 84, 89, 114, 164, 174, 306, 450–476, 498–499, 513, 517, 524, 531, 566, 590, 614, 623, 637, 661–662 American Toad, 15–39, 89, 98, 102, 104, 146, 316, 467 Americanus (clade), 40, 62, 92, 105, 112, 118, 148, 160, 162 americanus, Anaxyrus, xxiv, 15–40, 74–76, 80, 89–92, 96–97, 102, 105–106, 110, 112, 118–119, 131, 148–149, 153, 159–161, 165, 172–173, 242, 284, 463, 534, 537, 590, 720, 723 Anaxyrus, xiii, xvii, xxiv, xxv, xxx americanus, xxiv, 15–40, 74–76, 80, 89–92, 96–97, 102, 105–106, 110, 112, 118–119, 131, 148–149, 153, 159–161, 165, 172–173, 242, 284, 463, 534, 537, 590, 720, 723 a. charlesmithi, 15, 18, 31 a. copei, 15, 18 baxteri, xx, 17, 40–44, 105, 110, 112, 160 boreas, xviii, 18, 44–61, 66–68, 74, 85–87, 106, 118, 122, 124–125, 127–128, 131, 156, 161, 555, 652, 670, 675, 698

Dodd_Canada_int_5pgs_BM.indd 947

b. halophilus, 44–46, 48, 50, 66–67, 86, 124 californicus, xx, 17, 61–65, 72, 118–119, 144, 161 canorus, xx, 45–47, 65–72, 86, 124–125, 156, 691, 704 cognatus, 18, 61–62, 72–81, 106, 131, 144–146, 160–162, 165, 170 compactilis, 18, 47, 61, 74, 89, 117, 143–145, 148, 158, 236 debilis, xxix, 18, 47, 74, 81–85, 131, 141, 165, 172 d. insidior, 81–82 defensor, 148 exsul, 46, 66, 85–89, 122, 124, 128, 156 fowleri, xx, 17–18, 24, 40, 89–107, 112, 148–149, 153, 159–161, 165, 172, 174–175, 177, 224, 519 hemiophrys, 17–18, 40–41, 52, 74, 93, 105–110, 112, 131, 148, 160–161, 172 houstonensis, xx, 17, 40, 91–92, 105–106, 111–116, 146, 148, 160–161, 165, 463, 729 kelloggi, 82, 131, 141 microscaphus, 17, 61–62, 92, 106, 117–121, 131, 144–145, 148, 159–161 monfontanus, xx, 44, 46, 66, 86, 122–124, 127, 156 nelsoni, 46, 66–67, 86, 122, 124–128, 156 nevadensis, xx, 44, 46, 66, 86, 122, 124, 127–129 pliocompactilis, 137, 143 punctatus, 18, 47, 74, 82, 106, 118–119, 129–137, 141, 143–144, 148, 160–161, 713 quercicus, 74, 136–140, 179, 588 retiformis, xxix, 82, 131, 141–143 speciosus, 18, 47, 62, 74, 115, 117–118, 131, 143–146, 148, 161, 165, 172, 236, 713, 734 terrestris, 15, 17–18, 39–40, 74, 82, 89, 92, 97, 102, 105–106, 112, 118, 131, 140, 144, 146–156, 160–161, 172, 178–180, 265, 529, 722, 778 tiheni, 137 velatus, 92

williamsi, xx, 44, 46, 66, 86, 122–124, 128, 156–158 woodhousii, 16–18, 23, 40, 61–62, 74, 76, 80, 84, 89–93, 95, 105–106, 112, 115, 117–118, 120–121, 127, 131, 144, 146, 148, 158–168, 172, 174–175, 177, 242, 334, 713 w. australis, 158, 160 w. bexarensis, 158, 160 w. velatus, 89, 91, 158, 160 andersonii, Dryophytes, 102, 221–225, 231, 236, 248, 254, 257, 277, 283, 590 Aquarana, xxv, 450, 453, 483, 486, 512–513, 516–517, 523–524, 544, 568, 570, 596, 599, 635–636 arenicolor, Dryophytes, 225–230, 236, 248, 257, 269, 277, 289, 291–293, 299, 320 areolatus, Lithobates, 424–431, 433, 439, 442, 445, 448, 532–533, 544, 577, 583, 590–591 Arizona Toad, 117–121 Arizona Treefrog, xi, 288–291 Arroyo Toad, xx, 61–65 Ascaphidae, 1 Ascaphus, xiii, xiv, xv, xvii, xviii, xxiv, xxix, xxx montanus, xx, 1–6, 8–13 truei, 1–2, 4–5, 7–14 t. californicus, 7–8 Atlantic Coast Leopard Frog, 520–522 augusti, Craugastor, 182–185 auratus, Dendrobates, xv, xvi, xvii, 759–760 aurora, Rana, 307, 463, 468, 599, 647–655, 664, 670–672, 674–677, 689, 694, 697–700, 702 avivoca, Dryophytes, 222, 230–234, 236, 246, 257, 262, 269, 276–277 Barking Frog, 182–185 Barking Treefrog, 249, 261–267 Batrachochytrium dendrobatidis, xix, 33, 42, 56, 110, 115, 120, 165, 175, 230, 243, 284, 307, 335, 350, 368, 430, 436, 469, 481, 500, 538, 575, 591, 623, 634, 645, 653, 662, 668, 676, 685, 691–692, 699, 705, 724, 763, 782 baudinii, Smilisca, 227, 236, 393–395

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948  Index to Scientific and Common Names

baxteri, Anaxyrus, xx, 17, 40–44, 105, 110, 112, 160 berlandieri, Lithobates, 425, 431–439, 442–443, 463, 478, 528, 533, 541, 544, 580, 582–583, 589, 642, 646 Bird-­voiced Treefrog, 230–234 Black Toad, 85–89 blairi, Lithobates, 84, 417, 424–425, 430–431, 433, 436–444, 478, 528, 533, 541, 544–545, 558, 563, 582–583, 587, 589–590, 593, 642 blanchardi, Acris, 84, 193–210, 212, 214, 216, 218–219, 283, 320, 340, 372, 380, 384, 463, 496 Blanchard’s Cricket Frog, xx, 193–206, 496 bombifrons, Spea, 79, 84, 710, 713, 729–739, 742–744, 749–751, 755 Boreal Chorus Frog, 357–370 Boreas (clade), 66, 86, 124 boreas, Anaxyrus, xviii, 18, 44–61, 66–68, 74, 85–87, 106, 118, 122, 124–125, 127–128, 131, 156, 161, 555, 652, 670, 675, 698 boylii, Rana, 468, 471, 648, 656–663, 665, 672, 679, 688–689, 695, 701–702 brachyphona, Pseudacris, 311–316, 325, 340, 347, 355, 360, 372, 380, 384, 388 brimleyi, Pseudacris, 313, 317–319, 340, 347, 355, 360, 372, 380, 388 Brimley’s Chorus Frog, 317–319 Bronze Frog, 483–507 Bufo antecessor, 158, 160 Bufonidae, xxiv, 15 Bufo planiorum, 158, 160 Burrowing Toad, 421–423 cadaverina, Hyliola, xxv, 225, 227, 291–295, 298–299, 301 caerulea, Litoria, 771–772 California Red-­legged Frog, xx, 649, 670–678 California Toad, 44 California Treefrog, 291–296 californicus, Anaxyrus, xx, 17, 61–65, 72, 118–119, 144, 161 Cambarus diogenes, 426, 616–617 Canadian Toad, 40, 105–110 Cane Toad, 39, 176–181 canorus, Anaxyrus, xx, 45–47, 65–72, 86, 124–125, 156, 691, 704 Canyon Treefrog, 225–230 capito, Lithobates, 140, 425, 444–449, 532, 544, 576–577, 579 carolinensis, Gastrophryne, 103, 140, 179, 402–410, 412, 414–415, 417–418 Carpenter Frog, 635–641 cascadae, Rana, 648, 652, 657, 664–670, 672, 679, 689–690, 694–695, 698, 702 Cascades Frog, 664–670

Dodd_Canada_int_5pgs_BM.indd 948

catesbeianus, Lithobates, xiii, 30, 39, 64, 126–127, 164, 201–202, 283, 305, 316, 417, 425, 442, 450–477, 482, 486–487, 498, 511–513, 515, 517, 519, 524, 528, 531, 544, 568, 570–571, 573, 575, 599, 614, 617, 632, 636, 646, 651, 654–655, 658, 662–663, 671, 675–677, 701, 722, 741, 768 Chiricahua Leopard Frog, xx, 477–483 chiricahuensis, Lithobates, xx, 433, 439, 442, 477–483, 508–511, 541, 544, 582, 634, 642, 646 chrysoscelis, Dryophytes, 32, 146, 165, 174, 202, 227, 231, 234–246, 248, 257, 262, 265, 269, 274–277, 279–280, 282–284, 287, 289, 589, 719, 723, 737 cinereus, Dryophytes, xxi, 179, 222, 224, 227, 231, 236, 246–255, 257, 262, 265–269, 277, 320, 408, 415, 463, 514, 519, 588, 773, 775, 777–778 clamitans, Lithobates, xxx, 33, 202, 204, 335, 425, 430, 445, 453, 463, 468, 483–507, 513, 517, 519, 523–526, 531, 534, 568, 570–571, 573, 575, 582, 599, 616, 640 clarkii, Pseudacris, 196, 227, 236, 248, 277, 299, 313, 319–322, 340, 346–349, 355, 360, 372, 380, 384, 388 Cliff Chirping Frog, 190–192 Coastal Tailed Frog, 7–14 cognatus, Anaxyrus, 18, 61–62, 72–81, 106, 131, 144–146, 160–162, 165, 170 Collinses’ Mountain Chorus Frog, 311–316 collinsorum, Pseudacris, 311–317, 340, 347, 355, 360, 372, 388 Columbia Spotted Frog, 678–687 Cope’s Gray Treefrog, 202, 234–246, 279, 723 Coqui, 761–764 coqui, Eleutherodactylus, 761–764 couchii, Scaphiopus, 135, 146, 170, 174, 418, 708–714, 726, 728, 734–736, 755 Couch’s Spadefoot, 708–714 Cranopsis alvaria, 167 Cranopsis nebulifer, 171 Crapaud d’Amerique, 15 Crapaud de Fowler, 89 Crapaud de l’Ouest, 44 Crapaud de Wood­house, 158 Crapaud des Grandes Plaines, 72 Crapaud du Canada, 105 Crapaud pied-­bêche de Plaines, 729 Crapaud pied-­bêche du ­Grand Bassin, 742 Craugastor, xxiv, xxx augusti, 182–185 a. cactorum, 182–184 a. latrans, 182–184 Craugastoridae, 182

Crawfish Frog, 424–431 crepitans, Acris, 193–198, 201, 203, 205–215, 218–219, 236, 283, 340, 372, 463, 640 crucifer, Pseudacris, 153, 236, 311, 313, 323–338, 340, 356, 376, 380, 382, 384, 388, 391, 463, 496, 534, 616 Cuban Treefrog, 153, 180, 255, 273, 773–778 cystignathoides, Eleutherodactylus, 186–188 Czatkobatrachus, xv debilis, Anaxyrus, xxix, 18, 47, 74, 81–85, 131, 141, 165, 172 Dendrobates, xv, xvi, xvii, xxiv auratus, xv, xvi, xvii, 759–760 Dendrobatidae, 759 Dermocystidium, 500, 559, 575, 591 ­Dixie Valley Toad, xx, 156–158 dorsalis, Rhinophrynus, 421–423 draytonii, Rana, xx, 463, 467, 647–648, 653, 665, 670–678 Dryophytes, xiv, xxi, xxiv, xxv, xxx andersonii, 102, 221–225, 231, 236, 248, 254, 257, 277, 283, 590 arenicolor, 225–230, 236, 248, 257, 269, 277, 289, 291–293, 299, 320 avivoca, 222, 230–234, 236, 246, 257, 262, 269, 276–277 chrysoscelis, 32, 146, 165, 174, 202, 227, 231, 234–246, 248, 257, 262, 265, 269, 274–277, 279–280, 282–284, 287, 289, 589, 719, 723, 737 cinereus, xxi, 179, 222, 224, 227, 231, 236, 246–255, 257, 262, 265–269, 277, 320, 408, 415, 463, 514, 519, 588, 773, 775, 777–778 femoralis, 140, 221–222, 224, 227, 231, 234, 246, 256–262, 266, 268–269, 777 gratiosus, 153, 222, 231, 236, 246, 248, 250, 255, 257, 259–267, 269, 463, 589 squirellus, 179, 223, 227, 231, 236, 246, 248, 257, 262, 267–274, 277, 289–290, 298–299, 408, 415, 588, 777–778 versicolor, 32, 146, 224, 227, 230–231, 234–241, 243–244, 246, 248, 268, 274–289, 320, 372, 380, 384, 463, 640 wrightorum, 226, 228–229, 269, 288–291, 298, 463 Dusky Gopher Frog, xx, 576–580 Dwarf American Toad, 18 Eastern Narrow-­mouthed Frog, xvii, xviii, 402–410 Eastern Spadefoot, xvii, 714–725, 726, 728 East Texas Toad, 92

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Index to Scientific and Common Names 949

Eleutherodactylidae, 186 Eleutherodactylus, xiii, xv, xvi, xvii, xxiv, xxx coqui, 761–764 cystignathoides, 186–188 guttilatus, 188–190 marnockii, 188–192 planirostris, ix, 764–767 exsul, Anaxyrus, 46, 66, 85–89, 122, 124, 128, 156 femoralis, Dryophytes, 140, 221–222, 224, 227, 231, 234, 246, 256–262, 266, 268–269, 777 feriarum, Pseudacris, 311, 313, 316, 318, 320, 325, 334, 338–348, 354–356, 358, 360, 362, 370–374, 380, 387–389 fisheri, Lithobates, x, 477–478, 508–511, 527, 530, 543 fodiens, Smilisca, 395–398 Foothill Yellow-­legged Frog, 656–663 fouquettei, Pseudacris, 277, 313, 320–322, 338–340, 346–350, 355–356, 358, 360, 371–373, 384, 386–388 fowleri, Anaxyrus, xx, 17–18, 24, 40, 89–107, 112, 148–149, 153, 159–161, 165, 172, 174–175, 177, 224, 519 Fowler’s Toad, xx, 89–104, 146, 165, 174 fragilis, Leptodactylus, xv, 399–401 Gastrophryne, xxiv, xxx carolinensis, 103, 140, 179, 282, 402–410, 412, 414–415, 417–418 mazatlanensis, 410–412, 415 olivacea, 146, 165, 236, 402, 404, 409–419, 713 Glandirana, xxiv, xxix rugosa, 768–770 Gopher Frog, 139, 444–449, 532 gratiosus, Dryophytes, 153, 222, 231, 236, 246, 248, 250, 255, 257, 259–267, 269, 463, 589 Gray Treefrog, 234, 239, 244, 279–288 Gray Treefrog complex, xxxii, 230–231, 235, 237–238, 243, 245, 275–278, 283–284, 287, 289 ­Great Basin Spadefoot, xx, 128, 742–748 ­Great Plains Toad, 72–81 Green and Black Dart-­Poison Frog, 759–760 Green Frog, 202, 335, 483–507, 531 Green­house Frog, 764–767 Green Toad, 81–85 Green Treefrog (Dryophytes), xxi, 246–255, 266 Green Tree Frog (Litoria), 771–772 Grenouille à pattes rouges du Nord, 647 Grenouille-­à-­queue côtière, 7 Grenouille-­à-­queue des Rocheuses, 1

Dodd_Canada_int_5pgs_BM.indd 949

Grenouille des bois, 576 Grenouille des marais, 531 Grenouille du nord, 568 Grenouille léopard du Nord, 540 Grenouille maculée de Columbia, 678 Grenouille maculée d’Orégon, 693 Grenouille verte, 483 grylio, Lithobates, 216, 453, 468, 486, 512–517, 519, 524, 599, 635 gryllus, Acris, xxiii, 140, 193, 195–196, 206–208, 211–221, 514, 519, 588 Gulf Coast Toad, 102, 171–175 guttilatus, Eleutherodactylus, 188–190 hammondii, Spea, 79, 145, 729, 731, 737–744, 748–750, 754 heckscheri, Lithobates, xvii, 152, 453, 486, 513, 516–519, 524, 599 hemiophrys, Anaxyrus, 17–18, 40–41, 52, 74, 93, 105–110, 112, 131, 148, 160–161, 172 holbrookii, Scaphiopus, 32, 140, 179, 242, 265, 335, 463, 590, 709, 714–727 Hot Creek Toad, xx, 122–123 houstonensis, Anaxyrus, xx, 17, 40, 91–92, 105–106, 111–116, 146, 148, 160–161, 165, 463, 729 Houston Toad, xx, 111–116 hurterii, Scaphiopus, 709, 713–714, 716, 719, 725–729, 735 Hurter’s Spadefoot, 725–729 Hyla, xxiv, xxv affinis, 225, 291 avivoca ogechiensis, 230–231 cinerea evittata, 246, 248 eximia, 288 nebulosa, 291, 296 versicolor sandersi, 234 Hylidae, 193 Hyliola, xiii, xiv, xxiv, xxv, xxix, xxx cadaverina, 225, 227, 291–295, 298–299, 301 regilla, xxv, 56, 227, 269, 277, 288–294, 296–310, 320, 325, 463, 468, 668, 675, 691, 698, 700 Hypopachus, xxiv, xxx, 404, 414 variolosus, 418–420 Ichthyophonus, 335, 500, 538, 559, 622 Ichthyostega, xiv illinoensis, Pseudacris, xx, 351–354, 379, 384 Illinois Chorus Frog, xx, 351–354 Incilius, xiii, xxiv, xxix alvarius, 46, 74, 79, 134, 160, 167–172, 713 canaliferus, 47, 82, 131 coccifer, 148

nebulifer, 18, 74, 92, 93, 102, 112, 113, 114, 115, 144, 148, 160, 161, 162, 171–175, 415, 434, 713 intermontana, Spea, xx, 119, 729, 731–732, 738, 742–750 kalmi, Pseudacris, 313, 318, 325–326, 338–340, 346–347, 350, 354–358, 360, 371–372, 387–388 kauffeldi, Lithobates, 520–523, 540, 580, 582, 587, 596 laevis, Xenopus, xiv, 392, 779–783 Leptodactylidae, 399 Leptodactylus, xxiv, xxx fragilis, xv, 399–401 Leptolegnia, 335 Lithobates, xiii, xvii, xviii, xxv, xxix, xxx areolatus, 424–431, 433, 439, 442, 445, 448, 532–533, 544, 577, 583, 590–591 a. circulosus, 425, 427, 429 berlandieri, 425, 431–439, 442–443, 463, 478, 528, 533, 541, 544, 580, 582–583, 589, 642, 646 blairi, 84, 417, 424–425, 430–431, 433, 436–444, 478, 528, 533, 541, 544–545, 558, 563, 582–583, 587, 589–590, 593, 642 capito, 140, 425, 444–449, 532, 544, 576–577, 579 c. aesopus, 444–445 c. stertens, 444–445 catesbeianus, xiii, 30, 39, 64, 126–127, 164, 201–202, 283, 305, 316, 417, 425, 442, 450–477, 482, 486–487, 498, 511–513, 515, 517, 519, 524, 528, 531, 544, 568, 570–571, 573, 575, 599, 614, 617, 632, 636, 646, 651, 654–655, 658, 662–663, 671, 675–677, 701, 722, 741, 768 chiricahuensis, xx, 433, 439, 442, 477–483, 508–511, 541, 544, 582, 634, 642, 646 clamitans, xxx, 33, 202, 204, 335, 425, 430, 445, 453, 463, 468, 483–507, 513, 517, 519, 523–526, 531, 534, 568, 570–571, 573, 575, 582, 599, 616, 640 c. melanota, 483, 485–486 fisheri, x, 477–478, 508–511, 527, 530, 543 grylio, 216, 453, 468, 486, 512–517, 519, 524, 599, 635 heckscheri, xvii, 152, 453, 486, 513, 516–519, 524, 599 kauffeldi, 520–523, 540, 580, 582, 587, 596 maslini, 596, 599 okaloosae, 453, 486–487, 498, 513, 517, 523–527, 599

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950  Index to Scientific and Common Names

Lithobates (cont.) onca, xx, 433, 439, 508–509, 511, 527–530, 543, 582, 641–643 palustris, xvii, 31–32, 425, 433, 439, 491–492, 522, 531–541, 544, 577, 580, 582–583 p. mansuetii, 531, 533 pipiens, xx, xxvi, 32, 36, 174, 242, 283, 365, 385, 424–425, 431, 433, 435, 437–443, 453, 463, 474, 477–478, 485, 491–492, 498, 501, 508–509, 520–521, 527–529, 531–532, 534, 540–568, 572, 580–583, 587–589, 592–593, 596, 600, 624, 636, 641–642, 669 “Burnsi,” 540–541, 545, 548, 556, 560, 582 “Kandiyohi,” 540–541, 545, 560 septentrionalis, 453, 483, 486–487, 497–498, 513, 517, 524, 568–576, 599 sevosus, xx, 425, 445, 532, 544, 576–580 sphenocephalus, 31–32, 140, 179, 224, 242, 265, 335, 415, 424–425, 430, 433–434, 436–437, 439, 442–444, 514, 519–522, 528, 532–534, 542, 544, 563, 577, 580–596, 604, 640, 642, 723 s. utricularius, 520, 580, 582 subaquavocalis, 477–478, 482 sylvaticus, xv, 369, 425, 491–492, 541, 596–631, 648, 695 tarahumarae, 544, 631–635 virgatipes, 453, 486, 498, 513, 517, 524, 571, 599, 635–641 yavapaiensis, 432–433, 439, 480, 483, 528, 541, 544, 582, 634, 641–646 Litoria caerulea, 771–772 ­Little Grass Frog, 375–382 Lowland Burrowing Treefrog, 395–398 Lowland Leopard Frog, 641–646 luteiventris, Rana, 57, 365, 396, 657, 665, 678–687, 693–695, 700 maculata, Pseudacris, xv, 84, 196, 313, 320, 338–340, 346–348, 354–355, 357–370, 372, 387–389, 391, 428, 433, 561, 629, 737 marina, Rhinella, xvii, 39, 168, 176–181, 254, 762 marnockii, Eleutherodactylus, 188–192 mazatlanensis, Gastrophryne, 410–412, 415 Mexican Spadefoot, 736, 748–756 Mexican Treefrog, 393–398 Mexican White-­lipped Frog, 399–401 Microhyla areolata, 402, 404, 412, 414 Microhylidae, 402 microscaphus, Anaxyrus, 17, 61–62, 92, 106, 117–121, 131, 144–145, 148, 159–161 Mink Frog, 568–576

Dodd_Canada_int_5pgs_BM.indd 950

Mogollon Rim Leopard Frog, 508–511 monfontanus, Anaxyrus, xx, 44, 46, 66, 86, 122–124, 127, 156 montanus, Ascaphus, xx, 1–6, 8–13 Mountain Chorus Frog, 311–316 multiplicata, Spea, 79, 145, 713, 729–732, 735–740, 742–745, 748–756 muscosa, Rana, xx, 71, 305, 648, 657, 664–665, 672, 679, 688–693, 695, 701–703, 706 nebulifer, Incilius, 18, 74, 92, 93, 102, 112, 113, 114, 115, 144, 148, 160, 161, 162, 171–175, 415, 434, 713 nelsoni, Anaxyrus, 46, 66–67, 86, 122, 124–128, 156 nevadensis, Anaxyrus, xx, 44, 46, 66, 86, 122, 124, 127–129 New Jersey Chorus Frog, 354–357 Nigrita (clade), 313–314, 318 nigrita, Pseudacris, 236, 313, 318–320, 322, 325, 334, 338, 340–341, 345–348, 354–357, 360, 370–375, 380, 387–388 Northern Crawfish Frog, 425 Northern Cricket Frog, 206–213 Northern Green Frog, 483–484, 486 Northern Red-­legged Frog, 307, 647–655, 675 Novirana, xxv, 433, 439, 453, 478, 486, 509, 528, 533, 544, 570, 582, 599, 632, 636, 642 Oak Toad, 136–140 ocularis, Pseudacris, 140, 325, 334, 375–378, 588 okaloosae, Lithobates, 453, 486–487, 498, 513, 517, 523–527, 599 olivacea, Gastrophryne, 146, 165, 236, 402, 404, 409–419, 713 Ollotis alvaria, 167 Ollotis nebulifer, 171 onca, Lithobates, xx, 433, 439, 508–509, 511, 527–530, 543, 582, 641–643 Oregon Spotted Frog, xx, 365, 693–701 ornata, Pseudacris, 236, 313, 318, 320, 325, 334, 340, 371–372, 379–382, 384 Ornate Chorus Frog, 379–382 Osteopilus, xiv, xxiv, xxx septentrionalis, 153, 179, 255, 273, 773–778 Ouaouaron, 450 palustris, Lithobates, xvii, 31–32, 425, 433, 439, 491–492, 522, 531–541, 544, 577, 580, 582–583 Pantherana, 433, 533, 540 Parapseudacris, 323, 325 Pickerel Frog, 531–540 Pig Frog, 450, 468, 512–516

Pine Barrens Treefrog, 221–225 Pine Woods Treefrog, 256–261 Pipidae, 779 pipiens, Lithobates, xx, xxvi, 32, 36, 174, 242, 283, 365, 385, 424–425, 431, 433, 435, 437–443, 453, 463, 474, 477–478, 485, 491–492, 498, 501, 508–509, 520–521, 527–529, 531–532, 534, 540–568, 572, 580–583, 587–589, 592–593, 596, 600, 624, 636, 641–642, 669 Plains Leopard Frog, 437–444 Plains Spadefoot, 729–737 planirostris, Eleutherodactylus, ix, 764–767 Poloka, 176 Poloka lana, 450 pretiosa, Rana, xx, 55, 468, 599, 651–652, 657, 664–665, 678–679, 693–701 Procambarus acutus, 616 Procambarus clarkii, 64, 127, 307, 463, 499, 514, 588, 676 Prosalirus bitis, xv Pseudacris, xiii, xiv, xv, xxiv, xxv, xxix, xxx brachyphona, 311–316, 325, 340, 347, 355, 360, 372, 380, 384, 388 brimleyi, 313, 317–319, 340, 347, 355, 360, 372, 380, 388 clarkii, 196, 227, 236, 248, 277, 299, 313, 319–322, 340, 346–349, 355, 360, 372, 380, 384, 388 collinsorum, 311–316, 317, 340, 347, 355, 360, 372, 388 crucifer, 153, 236, 311, 313, 323–338, 340, 356, 376, 380, 382, 384, 388, 391, 463, 496, 534, 616 c. bartramiana, 323–324 feriarum, 311, 313, 316, 318, 320, 325, 334, 338–348, 354–356, 358, 360, 362, 370–374, 380, 387–389 fouquettei, 277, 313, 320–322, 338–340, 346–350, 355–356, 358, 360, 371–373, 384, 386–388 hypochondriaca, 296, 298–299 illinoensis, xx, 351–354, 379, 384 kalmi, 313, 318, 325–326, 338–340, 346–347, 350, 354–358, 360, 371–372, 387–388 maculata, xv, 84, 196, 313, 320, 338–340, 346–348, 354–355, 357–370, 372, 387–389, 391, 428, 433, 561, 629, 737 nigrita, 236, 313, 318–320, 322, 325, 334, 338, 340–341, 345–348, 354–357, 360, 370–375, 380, 387–388 ocularis, 140, 325, 334, 375–378, 588 ornata, 236, 313, 318, 320, 325, 334, 340, 371–372, 379–382, 384 sierrae, 296, 298–299 streckeri, 196, 236, 277, 320, 322, 325, 351, 379–380, 383–386

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Index to Scientific and Common Names 951

triseriata, xx, 236, 313–314, 318–320, 322, 325, 334, 338–340, 346–347, 351, 354–355, 357–360, 362, 368, 370–372, 387–392 Pternohyla, 396 punctatus, Anaxyrus, 18, 47, 74, 82, 106, 118–119, 129–137, 141, 143–144, 148, 160–161, 713 quercicus, Anaxyrus, 74, 136–140, 179, 588 Railroad Valley Toad, xx, 127–129 Rainette crucifere, 323 Rainette du Pacifique, 296 Rainette faux-­grillon boréale, 357 Rainette faux-­grillon de Blanchard, 193 Rainette faux-­grillon de l’Ouest, 387 Rainette versicolore, 274 Rana, xiii, xvii, xviii, xx, xxiv, xxv, xxix, xxx aurora, 307, 463, 468, 599, 647–655, 664, 670–672, 674–677, 689, 694, 697–700, 702 boylii, 468, 471, 648, 656–663, 665, 672, 679, 688–689, 695, 701–702 cascadae, 648, 652, 657, 664–670, 672, 679, 689–690, 694–695, 698, 702 draytonii, xx, 463, 467, 647–648, 653, 665, 670–678 luteiventris, 57, 365, 396, 657, 665, 678–687, 693–695, 700 muscosa, xx, 71, 305, 648, 657, 664–665, 672, 679, 688–693, 695, 701–703, 706 pretiosa, xx, 55, 468, 599, 651–652, 657, 664–665, 678–679, 693–701 sierrae, xx, 71, 648, 657, 664, 672, 688–692, 701–707 Ranavirus (ranavirus), 34, 57, 102, 153, 166, 180, 203, 212, 219, 243, 254, 267, 273, 284–285, 307, 316, 335–336, 345, 350, 368, 378, 386, 391, 409, 418, 430, 443, 449, 469, 481, 500–501, 515, 519, 522, 527, 538, 558, 560, 575, 579, 591, 622–623, 625, 630, 640, 685, 692, 706, 724, 736, 760, 766, 770, 777, 786–787 Ranidae, xxiv, xxx, 424 Red-­spotted Toad, 129–136 regilla, Hyliola, xxv, 56, 227, 269, 277, 288–294, 296–310, 320, 325, 463, 468, 668, 675, 691, 698, 700 Relict Leopard Frog, xx, 527–530 retiformis, Anaxyrus, xxix, 82, 131, 141–143 Rhacophorus, ix Rhinella, xiii, xvii, xxiv, xxix arenarum, 177 cerradensis, 177 jimi, 177 marina, xvii, 39, 168, 176–181, 254, 762 paracnemis, 177

Dodd_Canada_int_5pgs_BM.indd 951

poeppigii, 177 schneideri, 177 veredas, 177 Rhinophrynidae, 421 Rhinophrynus, xxiv, xxx dorsalis, 421–423 Rio Grande Chirping Frog, 186–188 Rio Grande Leopard Frog, 431–437, 646 River Frog, xvii, 451, 516–519 Rocky Mountain Tailed Frog, xx, 1–6 rugosa, Glandirana, 768–770 Scaphiopodidae, 708 Scaphiopus, xiv, xvii, xxiv, xxx alexanderi, 709 couchii, 135, 146, 170, 174, 418, 708–714, 726, 728, 734–736, 755 holbrookii, 32, 140, 179, 242, 265, 335, 463, 590, 709, 714–727 h. albus, 714, 717 hurterii, 709, 713–714, 716, 719, 725–729, 735 Scurrilirana, 425, 433, 439, 445, 528, 577, 582, 642 Scutiger, ix septentrionalis, Lithobates, 453, 483, 486–487, 497–498, 513, 517, 524, 568–576, 599 septentrionalis, Osteopilus, 153, 179, 255, 273, 773–778 sevosus, Lithobates, xx, 425, 445, 532, 544, 576–580 Sheep Frog, 418–420 sierrae, Rana, xx, 71, 648, 657, 664, 672, 688–692, 701–707 Sierra Nevada Yellow-­legged Frog, xx, 701–707 Smilisca, xiv, xxiv, xxx baudinii, 227, 236, 393–395 fodiens, 395–398 Sonoran Desert Toad, 167–170 Sonoran Green Toad, 141–143 Southern Chorus Frog, 355, 370–375 Southern Crawfish Frog, 425 Southern Cricket Frog, 213–220 Southern Leopard Frog, 439, 467, 580–596 Southern Toad, 146–156 Spea, xiii, xiv, xx, xxiv, xxx bombifrons, 79, 84, 710, 713, 729–739, 742–744, 749–751, 755 hammondii, 79, 145, 729, 731, 737–744, 748–750, 754 hardeni, 716 intermontana, xx, 119, 729, 731–732, 738, 742–750 multiplicata, 79, 145, 713, 729–732, 735–740, 742–745, 748–756 stagnalis, 738, 748–749 studeri, 731

speciosus, Anaxyrus, 18, 47, 62, 74, 115, 117–118, 131, 143–146, 148, 161, 165, 172, 236, 713, 734 sphenocephalus, Lithobates, 31–32, 140, 179, 224, 242, 265, 335, 415, 424–425, 430, 433–434, 436–437, 439, 442–444, 514, 519–522, 528, 532–534, 542, 544, 563, 577, 580–596, 604, 640, 642, 723 Spotted Chirping Frog, 188–190 Spotted Chorus Frog, 319–322 Spring Peeper, 102, 316, 323, 338, 382, 391, 555, 616 squirellus, Dryophytes, 179, 223, 227, 231, 236, 246, 248, 257, 262, 267–274, 277, 289–290, 298–299, 408, 415, 588, 777–778 Squirrel Treefrog, 267–274 streckeri, Pseudacris, 196, 236, 277, 320, 322, 325, 351, 379–380, 383–386 Strecker’s Chorus Frog, 383–386 Sterirana, 544, 582 sylvaticus, Lithobates, xv, 369, 425, 491–492, 541, 596–631, 648, 695 Syrrhophus campi, 186–187 Syrrhophus gaigae, 189 tarahumarae, Lithobates, 544, 631–635 Tarahumara Frog, 631–635 terrestris, Anaxyrus, 15, 17–18, 39–40, 74, 82, 89, 92, 97, 102, 105–106, 112, 118, 131, 140, 144, 146–156, 160–161, 172, 178–180, 265, 529, 722, 778 Texas Toad, 143–146 Triadobatrachus massinoti, xv triseriata, Pseudacris, xx, 236, 313–314, 318–320, 322, 325, 334, 338–340, 346–347, 351, 354–355, 357–360, 362, 368, 370–372, 387–392 Tropical Clawed Frog, xv, 782–784 tropicalis, Xenopus, 782–784 truei, Ascaphus, 1–2, 4–5, 7–14 Upland Chorus Frog, 316, 338–345 variolosus, Hypopachus, 418–420 Vegas Valley Leopard Frog, 508, 530 versicolor, Dryophytes, 32, 146, 224, 227, 230–231, 234–241, 243–244, 246, 248, 268, 274–289, 320, 372, 380, 384, 463, 640 virgatipes, Lithobates, 453, 486, 498, 513, 517, 524, 571, 599, 635–641 Western Chorus Frog, xx, 354, 387–392 Western Narrow-­mouthed Frog, 412–418 Western Spadefoot, 737–742

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952  Index to Scientific and Common Names

Western Toad, 44–61, 122, 128 White’s Green Tree Frog, 771 williamsi, Anaxyrus, xx, 44, 46, 66, 86, 122–124, 128, 156–158 Wood Frog, 29, 316, 552, 555, 596–631 Wood­house’s Toad, 80, 158–167 woodhousii, Anaxyrus, 16–18, 23, 40, 61–62, 74, 76, 80, 84, 89–93, 95,

Dodd_Canada_int_5pgs_BM.indd 952

105–106, 112, 115, 117–118, 120–121, 127, 131, 144, 146, 148, 158–168, 172, 174–175, 177, 242, 334, 713 wrightorum, Dryophytes, 226, 228–229, 269, 288–291, 298, 463 Wrinkled Frog, 768–770 Wyoming Toad, xx, 40–44

Xenopus, xiv, xxiv, xxix laevis, xiv, 392, 779–783 tropicalis, 782–784 yavapaiensis, Lithobates, 432–433, 439, 480, 483, 528, 541, 544, 582, 634, 641–646 Yosemite Toad, xx, 65–72

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Index to Potential Stressors

17β-­estradiol, 505 2-­chloroethanol, 473 2,2,2-­trichloroethanol, 473 2,4-­D, 33, 36, 57, 286, 307, 335, 336, 559, 563, 627, 668, 785 3-­trifluoromethyl-4-­nitrophenol, 564. See also TFM 6-­chloro-2-­picolinic acid, 369 Abate®, 58, 504 acetochlor, 33, 57, 307, 335, 473, 559 acridine, 437 alachlor, 37, 562 aldrin, 36, 103, 212, 219, 472, 593 alkalinity, xxvii, 5, 108, 502, 572, 605, 740 Altosid®, 58 aluminum, 34, 35, 71, 212, 254, 285, 311, 321, 336, 411, 416, 502, 503, 561, 624, 625 ammonia, xvi, 36, 308, 502, 561, 594, 654 aniline, 564 arsenic, 154, 336, 410, 437, 472, 502, 561, 624, 634 atrazine, 33, 37, 57, 204, 244, 285, 307, 335, 336, 443, 473, 499, 503, 504, 539, 559, 560, 562, 566, 593, 627, 668, 777, 781 azinphos-­methyl, 472, 562, 747 barium, 212, 336, 538 Basudin®, 504 Bayer37289®, 58, 741 Bayer38920®, 472 benzene, 564 benzene hexachloride, 103, 593 beryllium, 212, 336, 538 bifenthrin, 308, 503, 562 B-­naphtal, 473 boron, 35, 103, 561, 624 bromoxynil, 504 cadmium, 34, 35, 154, 243, 336, 409, 410, 436, 472, 502, 538, 592, 624, 634, 686 cadmium chloride, 560 carbaryl, 33, 36, 57, 166, 255, 285, 286, 307, 309, 335, 336, 430, 443, 472, 503–505, 539, 559, 563, 593, 594, 626, 653, 654, 662, 668, 700

Dodd_Canada_int_5pgs_BM.indd 953

carbon tetrachloride, 103, 473, 539, 564 carbophenothion, 369 chloroform, 103, 336, 539, 564 chlorpyrifos, 33, 57, 244, 307–309, 335, 504, 559, 563, 626, 662, 668, 692 chlorpyrifos-­ethyl, 58 chlorpyrifos-­methyl, 58 chromium, 154, 409, 472 coal ash, 154, 472, 501, 502, 624 coal oil, 57 cobalt, 389, 409 conductivity, xxvii, 41, 138, 237, 239, 251, 271, 280, 288, 321, 331, 406, 503, 546, 572, 585, 600, 605, 609, 625, 626, 632, 637 copper, 36, 57, 154, 286, 336, 337, 409, 410, 472, 473, 502, 503, 538, 539, 561, 592, 624 copper sulfate, 451 crankcase oil, 255 crude oil, 473 daidzein, 505 DDE, 36, 204, 308, 309, 336, 369, 503, 562, 654, 692 DDT, 58, 103, 204, 205, 212, 308, 309, 336, 369, 472, 474, 503, 504, 561, 564, 626, 700 DEF, 303 diazinon, 308, 309, 504, 563, 662, 692, 747 dichlorobenzene, 564 dieldrin, 36, 103, 204, 212, 219, 336, 369, 472, 564, 593 dioxins, 504, 564 diquat, 563, 592 diquat dibromide (Reward®), 36, 286, 473, 503, 539 dissolved oxygen content (DOC), 21, 416, 455, 491, 552, 575 Dithane®, 504 endosulfan, 33, 36, 57, 58, 286, 307, 309, 335, 336, 472, 473, 504, 559, 563, 626, 668, 669, 747 endrin, 103, 212, 219, 369, 472, 593, 626 equilenin, 505 equilin, 505

esfenvalerate, 443, 563, 593 estradiol, 505 fenbuconazole, 562 fenitrothion, 36, 472, 504, 563 fenvalerate, 103, 255, 504, 563, 593 fire retardants, 594 fluorine, 592 fluoxastrobin, 369, 562 furans, 504 G-30493 (an insecticide), 58 G-30494 (an insecticide), 58 gallium, 336 GC-3582, 472 germanium, 409 glyphosate, 33, 36, 57, 80, 204, 286, 307, 335, 336, 443, 473, 503, 505, 539, 559, 562, 563, 592, 626, 627, 668 Guthion®, 472 hardness, 21, 625. See also conductivity Headline®, 80 heated ­water (nuclear reservoir), 154 heptachlor, 103, 204, 472 hexachlorobenzene, 204 hexachloroethane, 473 hexazinone, 473, 504, 563 hydrothol, 103 Ignite 280SL®, 80, 756 imidacloprid, 212, 503–505 Imidan®, 504 iron, 57, 212, 285, 336, 502, 538, 624 lead, 35, 154, 212, 336, 409, 410, 437, 471, 501, 502, 504, 560, 624, 686 leptophos-­com tri-­o-­totyl phosphate, 244 lindane, 103, 369 magnesium, 212, 285, 336, 502, 538, 572, 626, 764 malathion, 36, 103, 166, 244, 286, 308, 309, 336, 369, 472, 503, 563, 564, 593, 623, 626, 662 Mancozeb®, 37, 562 manganese, 212, 285, 502

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954  Index to Potential Stressors

mercuric chloride, 244, 519 mercury, 35, 103, 212, 244, 308, 336, 409, 410, 436, 472, 502, 503, 515, 519, 561, 624, 662 metalloestrogens, 505 methoxychlor, 36, 103, 369, 593 methylene chloride, 103, 473, 539, 564 methylmercury, 244, 472, 502, 624 methyl parathion, 58, 309, 741 metolachlor, 285, 369, 443, 504, 562 molinate, 103 m-­xylene, 564 naled, 369 nickel, 35, 212, 336, 409, 561 nitrates, xxvii, 36, 58, 153, 154, 244, 308, 392, 502, 561, 625, 653, 669, 699 nitrites, xxvii, 308, 625, 699 nitrobenzene, 564 Nova®, 504 octamethyl pyrophosphoramide, 593 ozone, 396 paraoxon, 244 paraquat, 103, 369, 436, 473, 592 parathion, 58, 309, 369, 472, 593, 741 PCB 126, 502, 504 pentachlorophenol, 473 perchlorate, 204 perfluorohexanesulphonic acid (PFHxS), 473, 505, 564, 565 perfluorooctanesulfonate (PFOS), 37, 287, 473, 505, 564, 627 perfluorooctanoic acid (PFOA), 37, 287, 473, 505, 564, 627 permethrin, 33, 57, 307, 335, 473, 503–505, 559, 563, 668 pH (acidity), xxvii, 205, 212, 244, 336, 369, 502, 561, 572, 594, 624, 637, 640, 706

Dodd_Canada_int_5pgs_BM.indd 954

phenol, 103, 539, 564 phenyl saliginen cyclic phosphate, 244, 410, 593 Phos-­Chek D75F, 594 Phos-­Chek D75R, 594 phosdrin, 593 phosphate, 244, 321, 410, 593 phosphorus, 621, 625, 654, 764 phytoestrogens, 505 piperonyl butoxide, 369 polybrominated diphenyl ethers, 504 polychlorinated dibenzofuran, 204 polyethoxylated tallowamine (POEA), 36, 503, 626 pyraclostrobin, 308, 562 radiocesium, 254 Release®, 563, 592 Rodeo®, 204, 626 Ronnel®, 58, 741 rotenone, 564, 594 Roundup®, 36, 58, 286, 309, 336, 473, 503, 509, 563, 626, 669, 685 Roundup Concentrate®, 80 Roundup Ready-­to-­Use Plus®, 80, 756 Roundup WeatherMAX®, 80, 626, 627, 756 salinity, 103, 154, 156, 175, 204, 220, 252, 254, 256, 261, 267, 272, 273, 309, 375, 410, 437, 473, 516, 561, 594, 622, 625, 674, 781 salts, 288, 503, 625, 626, 628. See also salinity selenium, 35, 154, 285, 409, 410, 472, 502, 538, 624 Sertraline (Zoloft®), 627 Sevin®, 36, 563, 626. See also carbaryl silver, 409, 410, 538, 561 silvex, 103, 369 Stratego®, 80

strobilurin fungicides, 80 strontium, 154, 212, 472, 502, 624 strychnine, 180 sulfate, 35, 308, 451, 561, 572, 653 TDE, 103, 369 tebuconazole, 308, 562 temephos, 309, 504 tetra-­ethyl pyrophosphate, 593 TFM (lampricide), 286, 472, 564 thiodan, 58, 472 thiosemicarbazide (TSC), 627 tin, 336 tire debris, 624 titanium, 336 toluene, 564 toxaphene, 58, 103, 212, 309, 369, 472, 593 trans-­nonachlor, 204, 692 triallate, 504 triclopyr, 473, 504, 563, 592 trifluralin, 103, 504 trisodium nitrilotriacetic acid (NTA), 103, 140, 473, 539, 564 urea, 58, 308, 669 UVA radiation, 505, 564, 627 UVB radiation, 37, 58, 60, 71, 205, 244, 287, 295, 296, 309, 310, 336, 337, 369, 443, 473, 505, 565, 575, 593, 594, 627, 652, 654, 668, 669, 676, 677, 686, 692, 700 vanadium, 154, 285, 472, 502 Vision®, 36, 503, 562, 592, 626 zinc, 57, 154, 409, 410, 502, 538, 561, 592, 624, 634, 686 zirconium, 336 Zoloft®, 627. See also Sertraline

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