Invertebrate-Microbial Interactions: Ingested Fungal Enzymes in Arthropod Biology 9781501737138

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Explorations in Chemical Ecology Edited by Thomas Eisner and Jerrold Meinwald

Invertebrate-Microbial Interactions: Ingested Fungal Enzymes in Arthropod Biology by Michael M. Martin

INVERTEBRATEMICROBIAL INTERACTIONS Ingested Fungal Enzymes in Arthropod Biology

Michael M. Martin Department of Biology The University of Michigan

Comstock Publishing Associates a division of Cornell University Press ITHACA AND LONDON

Library of Congress Cataloging-in-Publication Data Martin, Michael M., 1935Invertebrate-microbial interactions. (Explorations in chemical ecology) Bibliography: p. 127 Includes index. 1. Insects—Physiology. 2. Fungal enzymes. 3. Digestion. I. Title. II. Series. QL495.M36 1987 595.7'0132 87-47549 ISBN 0-8014-2055-1 (alk. paper) ISBN 0-8014-9459-1 (pbk.: alk. paper)

Copyright © 1987 by Cornell University All rights reserved. Except for brief quotations in a review, this book, or parts thereof, must not be reproduced in any form without permission in writing from the publisher. For information, address Cornell University Press, 124 Roberts Place, Ithaca, New York 14850. First published 1987 by Cornell University Press. Printed in the United States of America The paper in this book is acid-free and meets the guidelines for permanence and durability of the Committee on Production Guidelines for Book Longevity of the Council on Library Resources.

Contents

Foreword

by Thomas Eisner and Jerrold Meinwald

Preface

1 The Digestion of Plant Cell Wall Polysaccharides; InsectMicrobial Interactions; and Symbiosis The Digestion of Cellulose 2 The Digestion of Other Plant Cell Wall Polysaccharides 11 The Study of Insect-Microbial Interactions 13

2 Acquired Enzymes in the Fungus-Growing Termite Macrotermes natalensis The Fungus-Growing Termites 17 The Enzymes of Macrotermes natalensis Gut Fluid 25 Conclusion 32 Epilogue 33

3 Acquired Enzymes in the Siricid Woodwasp Sirey cyaneus The Siricid Woodwasps 37 The Enzymes in the Woodwasps’ Gut Fluid 42 Conclusion 48

Contents

vi

4 Acquired Enzymes in Detritivores

49

Detritus as Food 50 Tipula abdominalis 54 Tracheoniscus rathkei 59 Prerequisites for Deriving Benefits from Ingested Enzymes 67 Studies from Other Laboratories 68 Conclusion 72

5 Acquired Enzymes in Cerambycid Beetles

73

Wood-Feeding Beetles 73 Monochamus marmorator 77 Saperda calcarata 85 Conclusion 89

6 The Symbiosis between the Attine Ants and the Fungi They

91

Culture in Their Nests The Ants and Their Associated Fungi 92 Host Plant Selection by Leaf-Cutting Ants 100 The Role of Antibiotics 103 The Role of Growth-Promoting Substances 105 The Role of Substrate Preparation 105 The Role of the Ants’ Fecal Material 107 The Origin of the Attine Ants’ Fecal Proteases 118 The Origin of Other Enzymes in the Ants’ Fecal Fluid 124 Summary 125

References

127

Index

143

Foreword

All organisms have chemical needs and sensitivities; and each is the source of substances of potential use or signal value to others. In the course of evolution; this potential for interaction has been thoroughly exploited; and organisms of the most diverse kinds have entered into chemical interdependencies; both mutualistic and antagonistic; which are central to the fabric of life. Chemical ecology focuses on such inter¬ dependencies. It brings the molecular dimension to our understanding of biological relationships; the relationships between animal and plant; the multicellular and unicellular; social and nonsocial; and kin and non-kin. It deals with chemical utilization and usurpation; chemical attraction and deterrence; as well as chemical defense; offense; and communication; and it has served to bring the biologist and chemist together in an area of dynamic and diverse investigation. The series Explorations in Chemical Ecology focuses on seminal studies by re¬ searchers who are expanding our knowledge in this field. The purpose is not only to summarize what they have learned but to convey a feeling for what it is like to venture into an unknown area of promise. One might think that chemical ecology considers only those chemi¬ cal events transpiring in the surroundings of an organism. Yet the in¬ sides of an organism, to the extent that they harbor parasites, sym¬ bionts, or exogenous chemical agents, may also be the province of chemical-ecological phenomena. Enzymes in the gut of some inverte¬ brates, as Michael Martin shows in this book, can come from a surpris¬ ing source: ingested fungal tissue. The resulting augmentation of diges••

Vll

••• viii

Foreword

live capacity may have been a key to evolutionary success in certain termites, woodwasps, and beetles. Michael Martin and his coworkers, more than any other contemporary group, deserve credit for document¬ ing this phenomenon, which has considerably enhanced our under¬ standing of cellulose degradation in nature. One particular facet of his work, the elucidation of how attine ants culture their fungal gardens by fecal administration of some of the fungus’s own digestive enzymes, is among the more exciting discoveries in recent chemical-ecological in¬ vestigation. Michael Martin is a chemist and biologist rolled into one. Given the nature of his research, this dual capacity has served him well. There is elegance to his work, and a very special style, qualities that come through in his writing. Thomas Eisner Jerrold Meinwald

Ithaca, New York

Preface

Those who study mutualism (or symbiosis in the narrow definition of the term), commensalism, and parasitism must inevitably consider the chemically mediated interactions between organisms. Indeed, I believe it would not be too much of an exaggeration to state that the study of invertebrate-microbial interactions is the study of biochemical alliances and conflicts between microorganisms (here defined to include bacteria, protozoa, yeasts, and filamentous fungi) and their metazoan hosts. Thus anyone who investigates invertebrate-microbial interaction is unavoida¬ bly a chemical ecologist. The pioneers of symbiosis research—P. Buch¬ ner, L. R. Cleveland, R. F. Hungate, W. Trager—were practicing chemical ecologists 50 years before the field had acquired its current label. Since the late 1960s, my colleagues and I have explored the biochemi¬ cal implications of invertebrate mycophagy, the consumption of fungal tissue. Fungal tissue is the food of many invertebrates from phylogenetically and ecologically diverse taxa. The fruiting bodies of higher fungi harbor numerous obligate fungivores. In addition, many species of inver¬ tebrates feed exclusively or predominantly on fungal tissue by grazing selectively on the vegetative portions and microscopic reproductive structures of filamentous fungi that permeate the habitats in which they live. Still other species ingest relatively small quantities of fungal tissue along with larger amounts of other materials, most often the substrate on which the fungus is growing. The most exciting finding of our research on insect mycophagy is that fungal enzymes, acquired by the ingestion of live fungal tissue or ix

X

Preface

substrate into which fungal enzymes have been secreted, can remain active in an arthropod’s gut and contribute to the digestion of structural polysaccharides. Because we have detected this phenomenon in ar¬ thropods from quite diverse taxa, I propose that the augmentation of digestive capacity through the ingestion of active fungal or bacterial enzymes may be widespread among invertebrates. The present book describes the series of investigations that has led to this idea. The research described herein has three main themes: cellulose digestion in insects, insect-microbial interactions, and the biochemical bases for symbiosis. The first chapter provides a general background for the work described in subsequent chapters. The next two chapters deal with the role of fungi in the nutrition of two groups of wood feeders, the fungus-growing termites and the siricid woodwasps. These insects are involved in complex, highly coevolved mutualistic associations with fungi. The fourth and fifth chapters discuss the importance of fungi in the diets of detritus feeders and in wood-feeding cerambycid beetles. The fungi consumed by the detritus feeders and the cerambycid beetles are casual associates of the organisms, not obligate symbionts. The book concludes with an account of the fungus-growing ants, another group that is involved in a spectacular mutualism with fungi. This book has two objectives. The first is to describe and discuss an interesting biological phenomenon, that is, the augmentation of diges¬ tive capacity through the ingestion of microbial enzymes. Although this phenomenon may not prove to be as widespread as I now believe it to be, it is nonetheless an important mechanism by which some insects digest the structural polysaccharides of higher plants. Biologists should be familiar with the phenomenon and the evidence that led to its dis¬ covery. The second objective is to illustrate the evolution of a research pro¬ gram. The direction of this research was not set by some grand design conceived in the late 1960s. Rather, the program has advanced one step at a time, with much improvisation along the way. With each new study that my colleagues and I have undertaken, we have introduced more sophisticated techniques into our experimental repertoire, and we have increased the number of procedures we use to test the validity of our hypothesis that ingested microbial enzymes augment digestive capac¬ ity. This progression should be evident to the reader. I would like to believe that the changes and refinements have brought a steady im¬ provement in the quality of our work. At times the directions we took were logical and defensible. At other times directions were determined by biases and hunches. In writing the chapters that follow, I have tried

Preface

xi

to reveal both my thought processes and my biases, including some that turned out to be wrong. Because I was trained as an organic chemist, it is often assumed that I recognized early in my career that my knowledge of chemistry and my methodological expertise would be valuable assets in research on chemi¬ cally mediated interactions between organisms and that I made a calcu¬ lated decision to enter the expanding field of chemical ecology. The truth is very different. I drifted into chemical ecology when a rather casually planned research project on the attine ants did not work out. I had learned about the attine ants in 1964 while reading the volume on insects in the Time-Life Nature Library. From the description of the natural his¬ tory of the ant-fungus symbiosis, I thought it quite possible that the ants might be a source of fungicidal chemicals. I had no plan to study the interactions between the ants and their mutualistic fungus and certainly no intent of ever examining other insect-microbial interactions. As it turned out, I found no interesting chemicals in the ants, but I found the ants themselves utterly fascinating. In spite of myself I began reading more and more about ants, ecology, and evolution and less and less about nonclassical carbonium ions, free radicals, and peroxyesters. Eventually I could resist no longer, and I undertook some research that did not use ants simply as a source of chemicals but rather had the goal of understanding something about the biology of the ants. Since then I have done no research of a purely chemical nature. Three coworkers have played particularly crucial roles in the work described in this book, and their contributions deserve special note. The first is Joan S. Martin. Joan and I spent our honeymoon on Barro Colorado Island in Panama, acquainting ourselves with the attine ants and planning the initial stages of our work on these insects. Joan was my tutor in biology, and her expertise in insect physiology was instru¬ mental in launching our research. The second is Norman D. Boyd, who was my tutor in biochemistry. His work on the attine ants finally led us to realize the potential significance of ingested fungal enzymes. The third is Jerome J. Kukor, my tutor in mycology, whose broad knowledge of biology and incredible versatility as an experimentalist made it pos¬ sible for us to jump from woodwasps to crane flies to woodlice to longhorned beetles in our quest for additional examples of invertebrates that benefit from the ingestion of microbial enzymes. Without the con¬ tributions of these three colleagues, I would have had nothing to write about. I am also deeply grateful to Neal Weber for introducing me to the

•• xn

Preface

attine ants and encouraging me to pursue the chemical studies that ultimately changed the entire direction of my professional career. Finally; this book has benefited from the contributions and criti¬ cisms of several colleagues. David Bignell and Reinhard Leuthold sup¬ plied me with data, electron micrographs; and suggestions for Chapter 2. Carole Chamier provided valuable criticisms of Chapter 4, and Jerome Howard and David Wiemer contributed data for inclusion in Chapter 6. Robin Crewe; Stephen Hubbell; and Debra Hoffmaster graciously lent me photographs for Chapter 2 and Chapter 6. Michael M. Martin

Ann Arbor, Michigan

Chapter 1 The Digestion of Plant Cell Wall Polysaccharides, Insect-Microbial Interactions, and Symbiosis

Plant cell walls are constructed of biopolymers that are potential sources of metabolic fuels and biosynthetic precursors for those organ¬ isms that can degrade them. The major chemical constituents of plant cell walls are polysaccharides; but lignin; a polymer of high molecular weight in which the repeating unit is the phenylpropane moiety; is also present in many plant tissues. Plant cell walls are made up of layers of highly crystalline aggregations of cellulose (microfibrils) embedded in a noncrystalline matrix of other complex noncellulosic substances. Of these the most important are the hemicelluloses; the pectic substances; and lignin. The layers that compose the cell wall differ in thickness; ratio of microfibrillar components to matrix components; orientation of microfibrils within the matrix; the nature of the matrix polysaccharides; the degree of lignification; and water content. In higher plants it is possible to recognize three distinct layers: the middle lamella; the pri¬ mary cell wall; and the secondary cell wall; listed in the order in which they are formed. Young growing cells; storage cells, and the photosynthesizing cells of leaves have only primary cell walls, whereas ma¬ ture cells, especially those that provide support or are involved in fluid transport, develop thick secondary cell walls toward the end of their growth.

1

2

Invertebrate-Microbial Interactions

The Digestion of Cellulose The Structure of Native Cellulose

Cellulose molecules are unbranched polymers of glucose residues linked by p-l,4-glycosidic bonds. Within the plant cell walk cellulose molecules are organized in bundles with their long axes parallel. These structural units are called microfibrils. The core of the microfibril is a region of high three-dimensional order and therefore high crystallinity. Surrounding this crystalline core is the paraciystalline cortex, a region in which the cellulose molecules are parallel to those in the core but are not so organized as to be part of the crystalline lattice, perhaps because of admixture with hemicellulose or other matrix polymers. Because aqueous solutions cannot penetrate regions of high crystallinity, dis¬ solved cellulases have much less access to cellulose molecules in the crystalline core than to those in the paracrystalline cortex. Crystallinity is therefore an important determinant of digestibility. Another factor that influences the digestibility of cellulose is the presence of lignin. When present, lignin forms a network that perme¬ ates the matrix of the cell wall. It is intimately associated with the other components and restricts the access of enzymes to susceptible bonds in the cellulose. Thus natural cellulose associated with lignin (lignocellulose) is much less easily degraded than purified crystalline cel¬ lulose. The Enzymes of Cellulose Digestion in Microorganisms

Microorganisms that degrade cellulose are abundant in nature. They include aerobic and anaerobic fungi, bacteria, and protozoa. In organ¬ isms that have been thoroughly studied, the digestion of cellulose has been shown to involve the synergistic action of a complex of enzymes (Wood and McCrae 1979; Ljungdahl and Eriksson 1985). The white rot fungi produce a soluble cellulase complex composed of three distinct classes of enzymes (Table 1.1): (1) endo-p-l,4-glucanases (also called Cx-cellulases), which randomly cleave p-l,4-glucosidic linkages along a cellulose chain, (2) eyo-(3-l,4-glucanases (also called Cacellulases), which cleave cellobiose or glucose units from the nonreduc¬ ing end of a cellulose chain, and (3) p-l,4-glucosidases, which hydrolyze cellobiose and water-soluble cellodextrins (oligosaccharides of glucose) to glucose. The presence of a complete cellulase complex in a cell-free extract is indicated by a capacity to degrade various forms of crystalline cellulose, such as filter paper, cotton, Avicell, a-cellulose fiber, or the microcrys-

Digestion; Microbes; and Symbiosis

3

Table 1.1 Enzymes of the cellulase complex of white rot fungi Enzyme3 Endo-$- 1,4-Glucanase

Cx-Cellulase Carboxymethylcellulase CMCase

Mode of action and products Random attack on (3-1,4-glucosidic bonds, generating transient cellodextrins, cellobiose, and glucose

1.4- P-D-Glucan 4-glucanohydrolase EC 3.2.1.4

E^o-P-l/t-Glucanase C1-Cellulase 1.4- (3-D-Glucan cellobiohydrolase EC 3.2.1.91

Hydrolysis of terminal or penultimate (3-1,4-gluosidic bonds of a linear (31,4-glucan chain, generating glucose or cellobiose

(3-1,4-Glucosidase

Hydrolysis of the (3-1,4-gluosidic bonds of cellobiose and cellodextrins, generating glucose

l;4-(3-D-Gluoside 4-glucohydrolase

EC 3.2.1.21

aAlternative designations appear as subentries.

talline cellulose used in preparing thin-layer plates. Neither the Cxcellulases nor the C1-cellulases are capable of degrading crystalline cellulose alone. The degradation of amorphous forms of cellulose, such as phosphoric acid—swollen cellulose, does not indicate that a com¬ plete cellulase complex is present because amorphorous cellulose can be degraded by the endoglucanases (Cx-cellulases) alone, without any contribution of activity from the exoglucanases (C1-cellulases). The Cxcellulases are conveniently assayed with a soluble derivative of cel¬ lulose, such as carboxymethylcellulose (CMC) or methylcellulose, as substrate. C1-Cellulases show only very weak activity toward these sub¬ stances. The digestion of native cellulose by white rot fungi is believed to be initiated when the Cx-cellulases attack isolated amorphous regions of the predominantly crystalline cellulose matrix, creating nick sites in the linear cellulose chains. The C1-cellulases are then presumed to attack the nonreducing ends generated at the nick sites, exposing additional sites for attack by the endoglucanases and generally disrupting the highly ordered structure of the cellulose aggregates. The continued combined action of the endo- and exoglucanases eventually brings about the com¬ plete degradation of the cellulose and the generation of glucose, cel¬ lobiose, and water-soluble cellodextrins of varying chain lengths. The f3-l,4-glucosidases complete the process by effecting the hydrolysis of cellobiose and the cellodextrins to glucose.

4

Invertebrate-Microbial Interactions

The enzymatic basis for cellulolysis in brown rot fungi and bacteria is less well understood. Multienzyme complexes are probably involved; and enzymes analogous to the endoglucanases of white rot fungi have been detected in a number of species. It has not usually been possible to detect enzymes analogous to the exoglucanases; however; and the nature of the synergism between the components of the enzyme com¬ plex remains obscure. It is also possible that nonenzymatic oxidative processes; involving H2Q2 and Fe2+, contribute to cellulose digestion by brown rot fungi. Cellulose digestion by bacteria often requires attach¬ ment of the bacterial cell to the cellulosic substrate because some of enzymes involved remain bound to the bacterial cell rather than being secreted into the medium. Cellulose Digestion in Insects

The ability of many insects to thrive on wood; foliage, and detritus has naturally stimulated investigations of the extent to which such species are able to digest the structural polysaccharides in their food. Table 1.2 lists cellulose-digesting insects. It includes only those species that have been shown to be able to digest cellulose by (1) a determina¬ tion of the assimilation efficiency of dietary cellulose, (2) a demonstra¬ tion that carbon-14 is incorporated into the body tissues or respired carbon dioxide of animals fed [U-14C]cellulose, or (3) a demonstration that the gut fluid of the insect is able to degrade some form of crystalline cellulose in vitro, that is, that the gut fluid contains a complete cellulase complex. It should be noted that the first two criteria are superior to the third as indicators of the capacity of an organism to digest cellulose in vivo. Low levels of activity toward crystalline cellulose in a gut extract, detected with an assay that has a long incubation time, do not neces¬ sarily imply that an organism can degrade native lignocellulose in its natural diet. Table 1.2 does not include species in which the gut fluid has only been shown to contain enzymes active against carboxymethylcellulose, Cellulose Azure, amorphous cellulose, or phosphoric acidswollen cellulose because activity toward these substrates does not indicate the presence of a complete cellulase complex. Wood feeders, especially termites and beetles, dominate the list of insects able to degrade cellulose. The values for assimilation efficiency that have been determined for a few species suggest that termites are the most efficient cellulose digesters. The only detritus feeders for which information is available are the immature forms of the stonefly Pteroriarcys proteus, the caddisfly Pycnopsyche luculenta, and the crane fly Tipula abdominalis. In all three of these species, the efficiency of cellulose assimilation is low (11%, 12%, and 19%, respectively). The only foliage

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Journals Ltd. Media defined in Martin and Weber (1969).

The growth of the Atta colombica. tonsipes fungus on defined media that vary in free amino acid content

Table 6.4

c c

-Q

109

110

Invertebrate-Microbial Interactions

tage to generating free amino acids rapidly if growth were limited by the rate at which carbon compounds of low molecular weight were pro¬ duced from cellulose or lignin. In fact, if amino acids were generated more rapidly than they could be assimilated, they would probably be lost to competitors. In contrast, low proteolytic capacity would be a potentially fatal limitation in an early colonizer or in a species that colonizes live plant tissue. Plant sap is a rich source of readily assimilable carbon com¬ pounds, and unless proteins were degraded rapidly, it is likely that growth would soon become limited by amino acids. The limited capac¬ ity of the ants’ fungus to degrade polypeptide nitrogen therefore sug¬ gests that it is ill equipped for life on freshly cut leaves, and we reasoned that the fungus-culturing activities of the ants must somehow compen¬ sate for the biochemical limitations of the fungus. Chemical Characteristics of the Ants’ Fecal Material The ants’ rectal fluid contains free amino acids (Martin and Martin 1970a). Although the amount present is doubtless variable and our mea¬ surements of rectal volume (Martin and Martin 1970b) not very accurate, we can estimate that the total amino acid concentration falls some¬ where in the range of 1—6 pg/fil. The rectal fluid from a single ant may contain as much as 0.5 |xg, with glutamic acid, histidine, arginine, pro¬ line, lysine, and leucine making up 82% (by weight) of the total. In

Table 6.5 The effect of added protease on the growth of the Atta colombica tonsipes fungus in Sabouraud’s dextrose broth

Culture medium3

Trypsin equivalents of proteolytic activity added to culture medium (pg/ml)

Dry weight of mycelium (mg)

Sabouraud’s dextrose broth

0.00

7.1 ± 4.9 (10)

Sabouraud’s dextrose broth plus Streptomyces griseus protease mixture

0.16

253.2 ± 43.2 (11)

Note: Each value is the mean plus or minus the standard error of the mean for the number of replicates shown in parentheses. aCulture flasks contained 50 ml of medium and were shaken for 12 days (180 rpm) at 250°C. Source: N. D. Boyd and M. M. Martin, "Faecal Proteinases of the Fungus-Growing Ant Atta teyana: Properties, Significance, and Possible Origin,” Insect Biochemistry 5 (1975) £19-635, copyright 1975, Pergamon Journals Ltd.

Attine Ants and Their Fungi

111

addition, the rectal fluid contains significant quantities of allantoic acid (1.9 jxg/ant) and allantoin (1.3 |xg/ant). Clearly when the ants defecate on substrate before inoculating it with fungus, they are supplementing the substrate with nitrogenous compounds that would be beneficial to the growth of their fungus. Any beneficial effect on growth, however, will be short-lived. As soon as the fecal amino acids are assimilated, the competitive status of the fungus will again be in jeopardy. What the fungus most needs is an additional supply of proteolytic enzymes. To test the hypothesis that a supplement of proteolytic enzymes would enhance the growth of the ants’ fungus, we compared the growth of the fungus in Sabouraud’s dextrose broth with its growth in this same culture medium amended with a commercial mixture of the proteases derived from the filamentous bacterium Streptomyces griseus (Table 6.5) (Boyd and Martin 1975a). The growth-enhancing effect of the added protease is dramatic, increasing the dry weight of mycelium produced after 12 days from 7 mg in the protease-free controls to 253 mg in the experimental cultures to which S. griseus protease had been added. Using the Azocoll assay, we readily detected proteolytic activity in the midgut and rectal fluids of A. colombica tonsipes workers (Martin and Martin 1970a, 1970b). Although the levels of activity vary consider¬ ably from one collection to another, activity is consistently higher in the rectal fluid than in the midgut fluid (Table 6.6). The ratio of rectal to midgut activity (units/microliter) can range from 1.5/1.0 to 12.0/1.0, most often falling between 2.0/1.0 and 5.0/1.0. The higher activity in the rec-

Table 6.6 Protease activity in midgut and rectal fluid of Atta colombica tonsipes workers QUt

Protease activity3

segment15

Units/ant

Units/jxl

Midgut Rectum

0.15-0.40 0.77-1.79

1.0-2.8 2.5-5.8

Note: Values have been recalculated from original data used to pre¬ pare tables in Martin and Martin (1970a, 1970b) plus data collected sub¬ sequently so that data could be presented in units more standard than those presented in the original publications. aOne unit of activity is the amount of enzyme required to bring about a change in absorbance at 580 nm of 0.001 absorbance units per minute under the conditions of the assay (37°C, pH 6.5, incubation volume 3.0 ml, 15 mg Azocoll, 20 minutes). bAssays were run on pooled samples of the midgut and rectal con¬ tents from 5-10 ants.

112

Invertebrate-Microbial Interactions

turn can be explained by the concentration of fluid that occurs in the rectum when water is resorbed by the rectal glands. To be sure that the proteolytic enzymes in the rectal fluid are still active in the ants’ fecal material when it is actually deposited on sub¬ strate, we assayed a substrate for protease activity immediately after the ants had deposited their fecal material on it. Ants from our captive colony of A colombica tonsipes readily incorporated cornflakes into their gardens, apparently treating them in exactly the same way that they treat leaves. Cornflakes recovered from the ants after being pro¬ cessed for incorporation into the gardens were soft, moist, and tacky. An extract of ant-treated cornflakes exhibited significant proteolytic ac¬ tivity, whereas an extract of untreated cornflakes had no protease ac¬ tivity. This experiment shows that when the ants defecate on substrate, they do indeed add a solution of active proteolytic enzymes to the inoculum. We have assayed proteolytic activity in the midgut and rectal fluids of an additional 16 species of attine ants from seven genera (Table 6.7) (Martin and Martin 1970b, 1971). The species selected represent the full range of substrate preferences typical of the attines. At one extreme, Acromyrmejc and Atta use only freshly cut leaf material as substrate for their gardens, whereas at the other, Cyphomyrmejc and Apterostigma use pieces of rotted wood, caterpillar feces, beetle frass, and insect carcasses. Species of Trachymyrmejc, Sericomyrmejc, and Mynrmicocrypta use vary¬ ing amounts of cut leaf material, flower parts, plant detritus, fragments of woody fruits, insect feces, and insect carcasses. In every species exam¬ ined, the rectal fluid was proteolytically active, and the level of activity was severalfold higher in the rectal fluid than in the midgut fluid. The production of proteolytically active fecal material is clearly a general characteristic of the fungus-growing ants. We have also assayed the midgut and rectal fluids of 35 nonattine species from 22 genera and 5 subfamilies (Martin and Martin 1970b, 1971). In striking contrast to our results with the attine ants, 26 of the nonattine species had no detectable protease activity in the rectal fluid, and in the 9 species in which protease activity was detectable, it repre¬ sented only a small fraction of the activity present in the midgut. Clear¬ ly, then, the production of fecal material in which proteolytic enzymes have been concentrated is not characteristic of all ants. It is a specific feature of the fungus growers. We have routinely assayed for protease activity using Azocoll, a gen¬ eral substrate against which most proteolytic enzymes exhibit activity. We have also shown, however, that the fecal fluid of A. tejcana degrades bovine serum albumin (BSA), casein, gelatin, and acid-denatured hemo-

Attine Ants and Their Fungi

113

Table 6.7 Protease activity in midgut and rectal fluids of attine ants Activity in midgut3

Activity in rectum3

Species

Units/ant

Units/|xl

Units/ant

Units/fxl

Cyphomyrmey costatus (50) C. rimosus trinitatis (40) Apterostigma dentigerum (10) A. mayri (12)

0.013 0.015 0.25 0.12 0.11 0.04 0.021 0.07 0.19 0.013 0.32 0.30 0.18 0.83 0.29 0.23

0.7 0.4 1.9 2.4 1.1 2.3 0.7 1.1 2.1 0.2 1.7 1.5 0.9 6.9 3.2 0.9

0.020 0.043 0.25 0.10 0.34 0.04 0.05 1.00 1.47 0.13 1.05 1.67 0.54 5.57 1.06 0.44

2.0 1.1 3.1 5.0 2.3 3.4 1.7 1.6 10.5 2.0 8.8 2.6 2.7 24.2 5.0 2.8

Apterostigma sp. (10) Myrmicocrypta ednaella (40) Trachymymey bugnioni (47) T. cornetzi (16) T. septentrionalis (5) Seriocomyrmey amabalis (20) S. urichi (5) Acromyrmey lobicornis (5) A. octospinosus (5) A. versicolor (5) Atta cephalotes (5) A. seydens (5)

Note: Values are for single runs on pooled samples from vaiying numbers of ants (shown in parentheses). Values have been recalculated from original data used to prepare tables in Martin and Martin (1970b, 1971) so that data could be presented in units more standard than those presented in the original publications. Genera are listed in presumed approximate phylogenetic position, Cyphomyrmey being the most primitive. “Activity is as defined in Table 6.6 (37°C, pH 6.65, incubation volume 3.0 ml, 15 mg Azocoll, 20 minutes).

globin (Boyd and Martin 1975a). It degrades native BSA 60-70% as rapid¬ ly as it degrades denatured BSA. The proteases of the rectal fluid are remarkably stable to denaturation and autolysis at pH 5.0-7.5 (Boyd and Martin 1975a). This range includes the pH encountered in the fungus gardens (5.0) as well as in the ants’ gut (crop, 7.2; midgut, 6.5; rectum, 5.8). The protease activity of the rectal fluid exhibits a broad maximum in the pH range 6.5-9.0 (Boyd and Martin 1975a). This broad activity maxi¬ mum results from the summation of the pH profiles of the component enzymes (Figure 6.7), the isolation of which will be discussed shortly. Protease API is most active under weakly alkaline conditions, whereas Proteases AP2 and AP3, which have superimposable pH profiles, are most active under weakly acidic or neutral conditions. Even at pH 5.0, the pH of the fungus garden, Proteases AP2 and AP3 exhibit 60-65% of their maximum possible activity. Stability, activity under acidic conditions, and broad-spectrum ac-

114

Invertebrate-Microbial Interactions Figure 6.7 Dependence of pro¬ teolytic activity (toward Azocoll) of proteases API, AP2, and AP3 on pH. Proteases AP2 and AP3 exhibit such similar pH profiles that a single curve has been drawn through the points for these two enzymes. [Re¬ drawn from N.D. Boyd and M.M. Martin, "Faecal Proteinases of the Fungus-Growing Ant Atta teyana: Properties, Significance, and Possi¬ ble Origin,’’ Insect Biochemistry 5 (1975) :619—635, copyright 1975, Pergamon Journals Ltd.]

tivity are clearly important properties in enzymes that must survive their period of residence in the ants’ digestive tract and then contribute to the degradation of leaf proteins in the ants’ fungus gardens after they have been deposited there by defecation. The Addition of the Ants' Fecal Material or Fecal Proteases to a Fungus Culture Once we had demonstrated that the ant fungus grows much faster in a culture medium containing free amino acids than in polypeptide, that its growth in a culture medium containing polypeptide as the major source of nitrogen is enhanced by the addition of a mixture of active Streptomyces griseus proteases, and that the ants’ fecal material contains active proteases, the functional significance of the ants’ ap¬ plication of fecal material to substrate prior to incorporation into a fungus garden seemed clear. Nonetheless, we decided to establish for certain that the fecal proteases of the ants have the same positive effect on the growth of the ants’ symbiotic fungus in Sabouraud’s dextrose broth that was observed when the proteases of S. griseus were added to the culture medium. As predicted, the fungus grew much faster in a medium to which the ants’ fecal material had been added (Table 6.8). To confirm that the growth-enhancing effect of the ants’ fecal mate¬ rial is due to the proteases present and not to some other component, we tested the effect of a mixture of the purified proteases from the rectal fluid on the growth of the ants’ fungus. When the chromatographic fractions containing Proteases API, AP2, and AP3 (see below) were re¬ combined and added to Sabouraud’s dextrose broth, growth enhance¬ ment of the fungus comparable to that brought about by the unfraction¬ ated fecal fluid was observed (Table 6.8). In these experiments, we have added much less fecal protease to the

Attine Ants and Their Fungi

115

Table 6.8 The effect of added fecal material and fecal proteases from Atta colombica tonsipes on the growth of the ants’ fungus in Sabouraud’s dextrose broth

Culture medium3 Run I Sabouraud’s dextrose broth

Typsin equivalents of proteolytic activity added to culture medium (|xg/ml)

Dry weight of mycelium (mg)

0.00

24.7 ± 8.9 (13)

Sabouraud’s dextrose broth plus ant fecal fluid

0.10

300.4 ± 19.1 (5)

Sabouraud’s dextrose broth plus chromatographic fractions containing partially purified fecal proteases

0.09

313.8 ± 20.6 (4)

0.00

7.1 ± 4.9 (10)

0.14

198.5 ± 34.1 (6)

Run 2 Sabouraud’s dextrose broth Sabouraud’s dextrose broth plus ant fecal fluid

Note: Each value is the mean plus or minus the standard error of the mean for the number of replicates shown in parentheses. “Culture flasks contained 50 ml of medium and were shaken for 12 days (180 rpm) at 25°C. Source: N. D. Boyd and M. M. Martin, ‘‘Faecal Proteinases of the Fungus-Growing Ant Atta teyana: Properties, Significance, and Possible Origin,” Insect Biochemistry 5 (1975):619-635, copyright 1975, Pergamon Journals Ltd.

liquid culture medium than the ants add to their licked and chewed substrate. The rectal fluid of a single worker often contains protease activity equivalent to 300-500 ng of trypsin. If we assume that the ants deposit one-tenth of their rectal fluid each time they defecate on a fragment of substrate being prepared for inoculation with their fungus, and if we further assume that a piece of substrate has a volume of 1 mm3, then the concentration of added protease in the treated substrate fragment would be 30-50 |Jig (trypsin equivalents)/ml. In the experi¬ ments in which we demonstrated a growth-enhancing effect of added fecal protease, we added only about 0.1 |xg (trypsin equivalents)/ml of protease activity to the culture flasks. If low concentrations of the fecal proteases enhance the growth of the ants’ fungal symbiont in a liquid culture medium in which the nitrogen is in the form of soluble polypep¬ tides, it is reasonable to conclude that high concentr ations would en-

116

Invertebrate-Microbial Interactions

hance growth on natural substrates in which the nitrogen takes the form of proteins. These experiments confirm the hypothesis that the application of the ants’ fecal material compensates for the inability of the fungus to bring about the rapid hydrolysis of protein and fosters rapid initial growth on protein-containing substrates. Other Enzymes in the Ants' Fecal Material Proteases are not the only active enzymes present in the fecal mate¬ rial of the attine ants. We have also established that the fecal fluid of A colombica tonsipes is able to depolymerize starch; pectin; sodium polypectate; xylan, carboxymethylcellulose; and chitin (Martin et al. 1973; 1975). The highest activity is toward pectin and sodium polypectate (Table 6.9). The fecal material of A colombica tonsipes also has pectin methyl esterase activity. Finally; glycosidase activity is present; as indi¬ cated by a capacity to catalyze the hydrolysis of maltose, trehalose, sucrose, turanose, cellobiose, and the p-nitrophenylglycosides of a- and (3-glucose and a- and (3-galactose (Martin et al. 1975). It is easy to envisage several important functions for these enzymes in the fungus-culturing activities of the ants. Obviously the degradation of the structural polysaccharides of the leaf tissue would provide nutrients that would be useful to the fungus immediately following inoculation. Indeed, we have observed a growth-enhancing effect of the ants’ fecal

Table 6.9 Enzymatic activity of Atta colombica tonsipes fecal fluid toward starch, caiboxymethylcellulose, xylan, pectin, and sodium polypectate Activity3 Substrate Starch CMC Xylan Pectin NaPP

Units/ant

Units/pi

27.3 22.8 27.9 900.0 1294.0

88.1 73.5 90.0 2903.0 4174.0

Note: Values are the average of duplicate determinations that differed by

less than 6.5%, run on pooled samples of 5-10 ants. CMC = carboxymethylcellulose. NaPP = sodium polypectate. aA unit of activity is the amount of enzyme required to liberate 1 pmole of maltose equivalents per hour under the conditions of the assay (0.5% substrate, 37°C, pH 5.5). Source: Recalculated from Martin et al. (1975).

Attine Ants and Their Fungi

117

material when the fungus is cultured in a synthetic medium containing starch or carboxymethylcellulose as the primary carbon source (Table 6.10) (Martin et al. 1975). Another way in which the enzymes in the ants’ fecal material, es¬ pecially the pectinases and the proteases, are likely to contribute to the initial growth of the fungus is by bringing about the rapid maceration of the plant tissue used as substrate. Enzymes that degrade pectin and protein have been implicated in the process of tissue maceration by pathogenic fungi (Bateman and Millar 1966; Cooper 1983) as well as by some insects (Talmadge and Albersheim 1969; King 1973). The macera¬ tion process is critical during the initial stages of growth because it facilitates hyphal invasion of the tissue and subsequent ramification of the fungus within the tissue. In addition, the maceration of the plant tissue improves access of other catabolic enzymes to potential sub¬ strates. Thus all of the processes involved in substrate preparation—the licking, chewing, and defecation behavior—can be interpreted as mech-

Table 6.10

The effect of added fecal fluid from Atta colombica tonsipes on the growth of the ants’ fungus in a synthetic medium

Carbon source in medium3

Fecal fluidb

Starch

Absent

Starch

Present

CMC

Absent

CMC

Present

Dry weight of mycelium (mg) 89.7 ± 7.6 (6) 135.4 ± 18.7 (4) 23.3 ± 2.7 (5) 66.3 ± 9.1 (3)

Note: Each value is the mean plus or minus the standard error of the

mean for the number of replicates shown in parentheses. CMC = carboxymethylcellulose. “Culture flasks contained 50 ml of medium (Martin and Weber 1969) that included 1.0 g of the carbon source. Flasks containing starch were shaken for 10 days, whereas flasks containing CMC were shaken for 16 days (180 rpm, 25°C). bWhen fecal fluid was included, the amount from 50 ants was present in each flask. Source: M. M. Martin, N. D. Boyd, M. J. Gieselmann, and R. G. Silver, "Activity of Faecal Fluid of a Leaf-Cutting Ant toward Plant Cell Wall Polysac¬ charides," Journal of Insect Physiology 21 (19751:1887-1892, copyright 1975, Pergamon Journals Ltd.

118

Invertebrate-Microbial Interactions

anisms for enhancing the competitive status of the fungus on fresh plant tissue^ a substrate that the fungal symbiont would not otherwise be able to exploit. The Origin of the Attine Ants' Fecal Proteases When we decided to attempt to isolate and characterize the ants’ fecal proteases, the idea had not yet occurred to us that the rectal enzymes might be ingested fungal enzymes. We had assumed all along that the rectal enzymes were the ants’ normal digestive proteases, which had somehow avoided the fate of the digestive proteases of other ant species. Our initial objective in isolating the fecal proteases of an attine ant was to understand why these enzymes were more stable than the proteases of other ants and to determine how the attines had managed to direct such a large proportion of their digestive proteases into their hindguts. The outcome of this effort, however, was to establish that the fecal proteases were not enzymes secreted by the ants’ midgut epi¬ thelium, but rather were ingested fungal enzymes (Boyd and Martin 1975a, 1975b). Purification of the Fecal Proteases of Atta colombica tonsipes and Atta tepana Before we undertook a full-fledged attempt at purifying the fecal proteases of an attine ant, it was essential for us to accumulate far more fecal material than we had ever had available to us in our earlier studies. To fulfill this mundane but critical condition, we had to find a better way to obtain the fecal material than by the tedious and time-consum¬ ing process of dissection. We found one when we discovered that im¬ mersion in ether would induce the workers to defecate. Ants, in groups of many hundreds, were quickly submerged in ether in large crystalliz¬ ing dishes. After a minute or so, ether and dead ants were poured off, leaving the bottom of the container covered with amber droplets of fecal fluid. After the last traces of ether had been removed in vacuo, the rectal fluid was washed from the bottom of the dish and from the dead ants with a small amount of water or buffer (0.01 M phosphate, pH 7.0). Dissection of ants treated in this manner revealed that the rectums were nearly empty, whereas the midguts were full and intact. Using this technique, we were able to accumulate fecal material with protease activity equivalent to 7.6 mg of trypsin from approximately 2 pounds of A. teyana workers. This quantity proved to be more than enough for all of our subsequent work. By subjecting the fecal material of A. colombica tonsipes and A. teyana to a conventional protein purification sequence (precipitation

Attine Ants and Their Fungi

119

with ammonium sulfate, gel permeation chromatography on Sephadex G-75, and anion-exchange chromatography on DEAE-cellulose), we es¬ tablished that the protease activity was due to three separate enzymes, designated API, AP2, and AP3. AP indicates that the enzymes are ant proteases, whereas the numbers indicate the order of their elution from the Sephadex G-75 column. Protease API accounted for about 20% of the total activity of the fecal material; Proteases AP2 and AP3 accounted for 30—35% and 45—50%, respectively. Properties of API, AP2, and AP3 Preincubation of the fecal material with diisopropylphosphofluoridate (DFP) resulted in a 20% loss of protease activity, whereas preincu¬ bation with ethylenediamine tetraacetate (EDTA) caused a loss of 80% of the activity (Table 6.11). DFP is a specific inhibitor of enzymes with a serine residue at the active site. EDTA, on the other hand, is a chelating agent that inhibits metalloenzymes. With both inhibitors present in the preincubation mixture, 100% loss of protease activity occurred. When we examined the separate enzymes, we found that Protease API is completely inhibited by DFP but is unaffected by EDTA. Proteases AP2 and AP3, on the other hand, are completely unaffected by DFP but are completely inhibited by EDTA. Also, Proteases AP2 and AP3, but not API, were more than 95% inhibited by 1,10-phenanthroline, another chelat¬ ing agent that inhibits metalloenzymes. The addition of zinc nitrate following treatment by phenanthroline regenerated 74% and 80% of the activity in Proteases AP2 and APS, respectively.

Table 6.11 The effect of inhibitors on the proteolytic activity of fecal fluid and the purified proteases (API, AP2, AP3) from Atta teyarta Concentration of inhibitor

Residual activity (%) Rectal fluid

API

AP2

AP3

None DFP EDTA 1,10-Phenanthroline l,10-Phenanthrolinea

0 10~3 10“2 IQ-3

100 80 20 —

100 0 100 100

—

—

100 100 0