Clinical small animal internal medicine 9781118496992, 111849699X, 9781118497036, 1118497031

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Clinical Small Animal Internal Medicine

Clinical Small Animal Internal Medicine Volume 1 Edited by David S. Bruyette, DVM, DACVIM (SAIM) Chief Medical Officer, Anivive Lifesciences, Long Beach, CA, USA

SECTION EDITORS Nick Bexfield, BVetMed, PhD, DSAM, DipECVIM‐CA, PGDipMEdSci, PGCHE, FHEA, MRCVS University of Cambridge

Johnny D. Chretin, DVM, DACVIM (O) TrueCare for Pets

Linda Kidd, DVM, PhD, DACVIM (SAIM) Western University of Health Sciences

Stephanie Kube, DVM, DACVIM (N)

Veterinary Neurology and Pain Management

Catherine Langston, DVM, DACVIM (SAIM) Ohio State University

Tina Jo Owen, DVM, DACVS Washington State University

Mark A. Oyama, DVM, MSCE, DACVIM‐Cardiology University of Pennsylvania

Nathan Peterson, DVM, DACVECC VCA West Los Angeles Animal Hospital

Lisa V. Reiter, DVM, DACVD McKeever Dermatology Clinics

Elizabeth A. Rozanski, DVM, DACVIM (SAIM), DACVECC Tufts University

Craig Ruaux, BVSc (Hons), PhD, MACVSc, DACVIM (SAIM) Massey University

Sheila M.F. Torres, DVM, MS, PhD University of Minnesota

This edition first published 2020 © 2020 by John Wiley & Sons, Inc. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions. The right of David S. Bruyette to be identified as the author of the editorial material in this work has been asserted in accordance with law. Registered Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA Editorial Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print‐on‐demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty The contents of this work are intended to further general scientific research, understanding, and discussion only and are not intended and should not be relied upon as recommending or promoting scientific method, diagnosis, or treatment by physicians for any particular patient. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of medicines, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each medicine, equipment, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Cataloging‐in‐Publication data applied for ISBN 9781118497067 Cover image: Cell images (left, middle) © Kateryna Kon/Shutterstock, Cell image (right) © David Bruyett, Dog and Cat image © David Bruyette Cover design by Wiley Set in 10/12pt Warnock by SPi Global, Pondicherry, India 10 9 8 7 6 5 4 3 2 1

­Dedication To CB Chastain, Dick Nelson and Ed Feldman for trying their best to make me a better clinician. For someone interested in internal medicine and endocrinology I could not have had better mentors. To all of my former students, interns and residents, thanks so much for always keeping me honest and forcing me to try to stay ahead of those I was supposed to be teaching. To all the wonderful folks at Wiley (Erica Judisch, Gillian Whitley, Purvi Patel, Holly Regan-Jones, and Katrina Maceda) that made this book possible and tried their best to keep me on track. You turned the idea into reality and improved the book at every step in the process. To my amazing children McKenna, Taylor, Ivy and Cole, thanks for letting me be a part of your lives. Being your Dad is the best job in the world. To my amazing wife and best friend Maya. You make my life complete and I look forward to sharing every day with you.

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Contents Preface  xix List of Contributors  xxi About the Companion Website  xxix VOLUME I Section 1 

Evaluation and Management of the Patient  1

  1 The Concept of One Medicine  3 Lonnie J. King   2 Statistical Interpretation for Practitioners  7 Philip H. Kass   3 Using Data for Clinical Decision Making  17 Philip H. Kass Section 2 

Endocrine Disease  27

  4 Principles of Endocrinology  29 Robert Kemppainen  5 Neuroendocrinology  35 Maya Lottati   6 Feline Acromegaly  43 David S. Bruyette   7 Pituitary‐Dependent Hyperadrenocorticism in Dogs and Cats  49 David S. Bruyette   8 Polyuria and Polydipsia  65 Jennifer L. Garcia   9 Canine Hypothyroidism  71 David S. Bruyette 10 Feline Hyperthyroidism  75 David S. Bruyette

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Contents

11 Hypoadrenocorticism in Dogs and Cats  81 Patty Lathan 12 Diabetes Mellitus in Dogs and Cats  93 Jacquie S. Rand 13 Hypoglycemia in Patients without Diabetes Mellitus: Evaluation and Management  103 Rhett Nichols 14 Canine Autoimmune Polyglandular Syndromes  113 Deborah Greco Section 3 

Cardiovascular Disease  117

15 Approach to the Patient with Suspected Cardiovascular Disease  119 Ingrid Ljungvall and Jens Häggström 16 Imaging in Cardiovascular Disease  127 Valérie Chetboul 17 Electrocardiography 165 Anna R.M. Gelzer and Marc S. Kraus 18 Pathophysiology of Heart Failure  175 Barret J. Bulmer 19 Management of Heart Failure  185 Steven Rosenthal and Mark A. Oyama 20 Ventricular Arrhythmias  199 Amara H. Estrada and Romain Pariaut 21 Supraventricular Arrhythmias  205 Romain Pariaut and Amara H. Estrada 22 Systemic Hypertension  219 Rebecca L. Stepien 23 Pulmonary Hypertension  225 Heidi B. Kellihan 24 Congenital Heart Disease  231 Brian A. Scansen 25 Valvular Heart Disease  245 Michele Borgarelli 26 Canine Myocardial Disease  253 M. Lynne O’Sullivan 27 Feline Myocardial Disease  267 Virginia Luis Fuentes

Contents

28 Pericardial Disease  275 Ashley B. Saunders and Sonya G. Gordon Section 4 

Respiratory Disease  287

29 A Respiratory Pattern‐Based Approach to Dyspnea  289 Christopher G. Byers 30 Feline Bronchial Asthma  297 Christine M. Serafin 31 Canine Chronic Bronchitis  305 Kevin Kumrow 32 Pulmonary Thromboembolism  313 Robert Goggs 33 Surgical Approaches to Thoracic Disease  325 Raegan Wells 34 Pleural Effusion  333 Ashley Allen and Gareth Buckley Section 5 

Critical Care Medicine  345

35 Approach to the Patient in the Critical Care Setting  347 Sarah Allen 36 Fluid Therapy  355 Teresa Rieser 37 Cardiopulmonary Resuscitation  365 Nathan Peterson 38 Respiratory Monitoring in Critical Care  373 Mathew Mellema 39 Acute Respiratory Failure  381 Matthew Mellema 40 Mechanical Ventilation  393 Kate Hopper and Mathew Mellema 41 Approach to the Patient with Shock  403 James W. Barr 42 Cardiogenic Shock  413 Nathan Peterson and James W. Barr 43 Septic Shock  421 James W. Barr

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44 Disorders of Heat and Cold  431 Sarah Allen 45 Acute Poisoning  437 Ben O’Kelley 46 Medical Management of Trauma and Burns  445 Nathan Peterson 47 Venomous Snake Bites  459 Nathan Peterson Section 6 

Gastrointestinal Disease  467

48 Gastrointestinal Imaging  469 Susanne M. Stieger‐Vanegas 49 Gastrointestinal Endoscopy  507 Craig Ruaux 50 Diseases of the Oral Cavity and Salivary Glands  533 Maria M. Soltero‐Rivera and Alexander M. Reiter 51 Gastritis and Gastric Ulceration in Dogs and Cats  547 Katie Tolbert and Emily Gould 52 Disorders of the Esophagus  557 Silvia Funes and Craig Ruaux 53 Motility Disorders of the Alimentary Tract  563 Reto Neiger and Silke Salavati 54 Exocrine Pancreatic Insufficiency in Dogs and Cats  583 Panagiotis G. Xenoulis 55 Pancreatitis in the Dog  591 Caroline Mansfield 56 Pancreatitis in the Cat  601 Craig Ruaux 57 Rectoanal Diseases – Medical and Surgical Management  609 Craig Ruaux and Milan Milovancev 58 Dysbiosis and the Use of Pre‐, Pro‐ and Synbiotics  621 Jan S. Suchodolski 59 Diagnosis and Management of Chronic Enteropathies  627 Karin Allenspach Section 7 

Diseases of the Liver, Gallbladder, and Bile Ducts  639

60 Approach to the Patient with Liver Disease  641 Emma O’Neill

Contents

61 Imaging in Hepatobiliary Disease  659 Esther Barrett 62 Metabolic, Toxic, and Neoplastic Diseases of the Liver  677 Jan Rothuizen 63 Feline Inflammatory Liver Disease  687 Nicki Reed 64 Canine Inflammatory Liver Disease  695 Nick Bexfield 65 Cirrhosis and its Consequences  705 Katherine Scott 66 Portosystemic Shunts and Microvascular Dysplasia  713 Geraldine Hunt 67 Diseases of the Gallbladder and Extrahepatic Biliary Ducts  721 Ben Harris Section 8 

Neurologic Disease  727

68 The Neurologic Examination  729 Alexander de Lahunta 69 Central Nervous System Trauma  741 Simon R. Platt 70 Seizures and Movement Disorders  759 Michael Podell 71 Disorders of the Forebrain  773 Sam N. Long 72 Vestibular Disease  789 Tammy Stevenson 73 Meningoencephalitis and Meningomyelitis  795 Christopher L. Mariani 74 Diseases of the Neuromuscular Junction  803 David Lipsitz 75 Myopathies 811 Marguerite F. Knipe 76 Myelopathy 815 Joan R. Coates 77 Neuroophthalmology 823 Bradford J. Holmberg

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VOLUME II Section 9  Part 1 

Infectious Disease  829

Diagnostic Considerations  831

78 Epidemiology of Infectious Disease  833 Peggy L. Schmidt and Helen T. Engelke 79 Laboratory Diagnosis of Infectious Diseases  839 Laia Solano‐Gallego and Gad Baneth Part 2 

Select Infectious Diseases and Disease Agents  849

80 Canine Distemper  851 David S. Bruyette 81 Canine Herpesvirus  855 Yvonne Drechsler 82 Canine Viral Enteritis  857 Margaret C. Barr 83 Viral Papillomatosis  861 Margaret C. Barr 84 Canine Influenza Virus  865 Ellen Collisson 85 Feline Parvovirus  869 Margaret C. Barr 86 Feline Coronavirus  873 Yvonne Drechsler 87 Feline Leukemia Virus  877 David S. Bruyette 88 Feline Immunodeficiency Virus  883 Tom Phillips 89 Feline Viral Upper Respiratory Tract Disease  887 Yvonne Drechsler 90 Rabies in Dogs and Cats  891 Emily Beeler and Karen Ehnert 91 West Nile Virus  899 Tracey McNamara 92 Ebola Virus  901 Linda Kidd

Contents

  93 Ehrlichiosis and Anaplasmosis  903 Pedro P. Vissotto de Paiva Diniz   94 Salmon Poisoning Disease  913 Pedro P. Vissotto de Paiva Diniz  95 Wolbachia pipientis Infection  917 Pedro P. Vissotto de Paiva Diniz  96 Bartonellosis  919 Pedro P. Vissotto de Paiva Diniz  97 Hemotropic Mycoplasma  927 Séverine Tasker  98 Nonhemotropic Mycoplasma, Ureaplasma, and L‐Form Bacteria  931 Joachim Spergser   99 Spotted Fever and Typhus Group Rickettsia  937 Linda Kidd 100 Lyme Borreliosis  941 Meryl P. Littman 101 Leptospirosis 945 Katharine F. Lunn 102 Yersiniosis 951 Maria Grazia Pennisi 103 Tularemia 955 Linda Kidd 104 Q Fever  959 Linda Kidd 105 Brucellosis 963 Erin E. Runcan and Marco A. Coutinho da Silva 106 Tetanus and Botulism  967 Andrea Fischer 107 Anthrax 971 Wayne E. Wingfield and Jerry J. Upp 108 Actinomycosis, Nocardiosis, and Mycobacterial Infections  977 Joanna Whitney and Vanessa R. Barrs 109 Fungal Infections  985 Jane E. Sykes 110 Protozoal and Protozoa‐Like Infections  1003 Gad Baneth and Laia Solano‐Gallego

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111 Coccidia 1023 Chris Adolph 112 Surgical, Traumatic, and Bite Wound Infections  1029 Laura L. Nelson 113 Canine Infectious Respiratory Disease Complex  1035 Jonathan Dear Part 3 

Therapeutic Considerations  1039

114 Antimicrobial Therapy in Dogs and Cats  1041 Katrina R. Viviano 115 Antifungal Therapy  1049 Daniel S. Foy Part 4 

Special Topics  1055

116 Nosocomial and Multidrug‐Resistant Infections  1057 Jason W. Stull and J. Scott Weese 117 Management of Infectious Disease in Kennels and Multicat Environments: Creating a Culture of Compliance  1063 Frank Bossong Section 10 

Renal and Genitourinary Disease  1067

118 Disorders of Sodium and Water Homeostasis  1069 Julien Guillaumin and Stephen DiBartola 119 Disorders of Phosphorus and Magnesium  1079 Rosanne Jepson 120 Acute Kidney Injury  1089 Adam E. Eatroff 121 Glomerular Disease  1101 Shelly L. Vaden 122 Obstructive Uropathy  1109 Edward Cooper and Brian A. Scansen 123 Urolithiasis in Small Animals  1123 Alice Defarges, Michelle Evason, Marilyn Dunn, and Allyson Berent 124 Prostatic Diseases  1157 Serge Chalhoub 125 Management of Chronic Kidney Disease  1165 Jessica Quimby

Contents

126 The Role of Dialysis  1175 Adam E. Eatroff 127 Micturition and Associated Disorders  1181 Julie K. Byron 128 Urinary Tract Infections  1189 Nicole Smee Section 11 

Oncologic Disease  1197

129 Approach to the Cancer Patient  1199 Lisa DiBernardi 130 Biology of Cancer and Cancer Genetics  1205 Mary‐Keara Boss 131 Endocrine Manifestations of Cancer: Ectopic Hormone Production  1213 Cory Brown 132 Paraneoplastic Syndromes  1217 Cory Brown 133 Lymphoid Leukemias, Myeloid Neoplasia, and Myelodysplastic Syndrome  1223 Angela R. Kozicki 134 Lymphomas 1231 Kristine Elaine Burgess 135 Plasma Cell Disorders  1241 Orna Kristal 136 Central Nervous System Tumors in Dogs and Cats  1247 David Ruslander 137 Cancer of the Nose and Mouth  1253 Lauren Askin Quarterman 138 Tumors of the Eye and Ocular Adnexa  1261 Erin M. Scott and Paul E. Miller 139 Cancer of the Heart  1271 Nick A. Schroeder and Lisa DiBernardi 140 Cancer of the Airway and Lung  1277 Joanne L. Intile 141 Cancer of the Esophagus and Stomach  1283 Avenelle I. Turner 142 Cancer of the Small and Large Intestine  1287 Edwin Brodsky

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143 Exocrine Pancreatic Cancer  1293 Avenelle I. Turner 144 Pancreatic Endocrine Tumors  1297 Karen Eiler 145 Liver and Biliary Tract Tumors  1303 Amanda K. Elpiner 146 Tumors of the Urinary Tract  1307 Pedro A. Boria 147 Tumors of the Male Reproductive System  1311 Trina Hazzah 148 Gynecologic Cancers  1317 Trina Hazzah 149 Mammary Cancer  1321 Julie Bulman‐Fleming 150 Tumors of Bone and Joint  1327 Stephanie L. Shaver, William T.N. Culp, and Robert B. Rebhun 151 Soft Tissue Sarcomas  1333 Lauren Askin Quarterman 152 Hemangiosarcoma 1339 Christine B. Oakley and John D. Chretin 153 Melanoma 1347 Philip J. Bergman 154 Nonmelanoma Skin Cancers  1353 Barbara E. Kitchell 155 Mast Cell Neoplasia  1359 Zachary M. Wright 156 Apheresis in Companion Animals  1369 Steven Suter Section 12 

Skin and Ear Diseases  1373

157 Approach to the Patient with Dermatologic Disease  1375 Lisa V. Reiter 158 Principles of Therapy of Dermatologic Diseases  1397 Sandra N. Koch 159 Atopic Dermatitis  1403 Alison Diesel

Contents

160 Allergic Skin Diseases  1413 Patrick Hensel 161 Cutaneous Adverse Food Reactions  1419 Ralf S. Mueller 162 Autoimmune and Immune‐Mediated Skin Diseases  1423 Nicole A. Heinrich 163 Approach to Alopecia  1433 Linda A. Frank 164 Canine Sterile Papular and Nodular Skin Diseases  1441 Sandra Diaz 165 Parasitic Skin Diseases  1449 Elizabeth E. Toops 166 Bacterial Pyodermas  1461 Jennifer R. Schissler 167 Otitis 1471 Sue Paterson 168 Cutaneous Manifestations of Systemic Disease  1481 Kinga Gortel 169 Superficial Necrolytic Dermatitis  1491 Mitchell D. Song 170 Miscellaneous Skin Diseases  1495 Lisa V. Reiter and Sheila M.F. Torres Section 13 

Diseases of Bone and Joint  1507

171 Skeletal Development and Homeostasis  1509 Matthew J. Allen and Gert J. Breur 172 Metabolic Bone Diseases  1521 Keren E. Dittmer 173 Osteoarthritis in Small Animals  1529 Steven A. Martinez 174 Developmental Orthopedic Diseases  1537 Gert J. Breur, Nicolaas E. Lambrechts, and Heather A. Towle Millard Section 14 

Social and Ethical Issues in Veterinary Medicine  1553

175 Canine and Feline End of Life Care  1555 Robin Downing

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Section 15 

Preventive Care  1565

176 Role of Immunization  1567 Melissa Kennedy 177 Behavior Triage for Internists and the General Practitioner  1571 Karen Lynn C. Sueda Section 16  Index  1583

Laboratory Support  1581

xix

Preface Internal medicine is hard. Progress in science and technology makes it almost impossible to keep abreast of recent advances and on top of that, all of us want to balance our life both at work and home. Is there enough time in the day for everything? There are a variety of ways in which we learn and the path is not the same for everyone. For those of us with an interest in internal medicine, and I’m assuming that includes the reader of this book, we rely on a number of resources. Our colleagues, continuing education seminars, the literature, professional educational networks, and reference textbooks. A concern of mine over the past few years has been the increasing reliance on technology to arrive at a clinical diagnosis rather than emphasizing the need to understand physiology and the value of a complete history and a thorough physical examination. We feel increasing pressure to arrive at a specific and definitive diagnosis and, more importantly, to arrive at that diagnosis almost immediately. Your goal in internal medicine is not to arrive at a diagnosis. I would much prefer to see a pet respond to my treatment and recover without a definitive diagnosis than to arrive at a definitive diagnosis at necropsy. If you are successful in achieving a diagnosis, that can be very rewarding. However, your goal should be to accurately identify problems and address those problems in a logical, timely, and cost‐effective manner, always weighing the risk/benefit of running myriad diagnostics versus improving the quality of life of your patient and the pet owner. While not every pet owner will be able to afford or desire to follow each and every one of our recommendations, it is our job to make sure that whatever decision the owner makes is based on being fully informed. Illness is really physiology gone awry. If you have an understanding of what is normal, it makes your job of identifying the abnormal much easier. The body has a limited repertoire of responses to an insult so often many diseases will have very similar clinical presentations.

While we all like to make lists of the 20 differential diagnoses for a given patient’s abnormalities, it’s important to recognize that in clinical practice, common things occur commonly and those differentials should be at the top of your rule‐out list. There are numerous excellent textbooks on the market and I use many of them on a daily/weekly basis. Some serve as definitive reference works for the topic under discussion. Some are brief, bullet point, clinically oriented texts that one can use to find something quickly. What I thought was currently lacking was a text that would be continually updated, provide enough background physiology to help the reader understand normal versus abnormal, and provide useful and, more importantly, clinically relevant material to help you with your patients. The goal of this text was to first identify section editors who were recognized experts in their field both academically and clinically. The section editors then identified topics they felt were of greatest clinical importance and selected authors who could translate this information into a text that would be used every day. I hope we have achieved that goal. We will be updating the text online with new information and updated references on a quarterly basis, adding additional sections and chapters with future editions, and uploading podcasts consisting of interviews with the authors to highlight and emphasize the material in the text and any recent advances in the field. We hope that you will find the text helpful and all credit for that success goes to the authors. Any omissions or errors lie with me so please let me know both the good and the bad so we can improve things going forward. As my favorite philosopher once said “It’s a magical world, Hobbes, ol’ buddy, let’s go exploring.” All the best and enjoy exploring the mysteries of medicine. Dave

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List of Contributors Chris Adolph, DVM, MS, PhD, DACVM (Parasitology)

Zoetis Inc. Tulsa, OK, USA Ashley Allen, DVM, DACVECC

College of Veterinary Medicine University of Florida Gainesville, FL, USA Matthew J. Allen, VetMB, PhD

Department of Veterinary Medicine University of Cambridge Cambridge, UK Sarah Allen, DVM, DACVECC

Massachusetts Veterinary Referral Hospital Woburn, MA, USA Karin Allenspach, Dr.Med.Vet., PhD, DECVIM‐CA

College of Veterinary Medicine Iowa State University Ames, IA, USA Gad Baneth, DVM, PhD, DECVCP

School of Veterinary Medicine Hebrew University Rehovot, Israel James W. Barr, DVM, DACVECC

BluePearl Veterinary Partners Tampa, FL, USA Margaret C. Barr, DVM, PhD

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Vanessa R. Barrs, BVSc (Hons), MVetClinStud, MACVSc (Small Animal), FACVSc (Feline), GradCertEd (Higher Ed)

University of Sydney Sydney, Australia Emily Beeler, DVM, MPH

Veterinary Public Health Program Los Angeles County Department of Public Health Los Angeles, CA, USA Allyson Berent, DVM, DACVIM (SAIM)

The Animal Medical Center New York, USA Philip J. Bergman, DVM, PhD, DACVIM (Oncology)

Katonah‐Bedford Veterinary Center Clinical Studies, VCA Antech Bedford Hills, NY, USA Nick Bexfield, BVetMed, PhD, DSAM, DECVIM‐CA, PGDipMEdSci, FHEA, MRCVS

The Queen’s Veterinary School Hospital University of Cambridge Cambridge, UK Michele Borgarelli, DVM, PhD, DECVIM (Cardiology)

Virginia‐Maryland Regional College of Veterinary Medicine Blacksburg, VA, USA Pedro A. Boria, DVM, MS, DACVIM (Oncology)

Blue Pearl Veterinary Partners Northfield, IL, USA Mary‐Keara Boss, DVM, PhD, DACVR (Radiation Oncology)

Colorado State University Fort Collins, CO, USA Frank Bossong, DVM

Esther Barrett, MA, VetMB, DVDI, DECVDI, MRCVSt

Wales and West Imaging Chepstow, UK

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

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­List of Contributors

Gert J. Breur, DVM, MS, PhD, DACVS

Johnny D. Chretin, DVM, DACVIM (Oncology)

College of Veterinary Medicine Purdue University West Lafayette, IN, USA

TrueCare for Pets Studio City, CA, USA

Edwin Brodsky, DVM, DACVIM (Oncology)

College of Veterinary Medicine University of Missouri Columbia, MO, USA

Veterinary Medical Center of Long Island West Islip, NY, USA Cory Brown, DVM, DACVIM (SAIM)

Joan R. Coates, DVM, MS, DACVIM (Neurology)

Ellen Collisson, MS, PhD

VetScan Mobile Diagnostics Powell, OH, USA

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

David S. Bruyette, DVM, DACVIM (SAIM)

Edward Cooper, VMD, MS, DACVECC

Anivive Lifesciences Long Beach, CA, USA Gareth Buckley, MA, VetMB, MRCVS, DACVECC

College of Veterinary Medicine University of Florida Gainesville, FL, USA Julie Bulman‐Fleming, DVM, DACVIM (Oncology)

Veterinary Cancer Group Tustin, CA, USA Barret J. Bulmer, DVM, MS, DACVIM (Cardiology)

Tufts Veterinary Emergency Treatment & Specialties Walpole, MA, USA Kristine Elaine Burgess, DVM, DACVIM (Oncology)

Cummings School of Veterinary Medicine Tufts University North Grafton, MA, USA Christopher G. Byers, DVM, DACVECC, DACVIM (SAIM), CVJ

CriticalCareDVM.com Omaha, NE, USA Julie K. Byron, DVM, MS, DACVIM (SAIM)

College of Veterinary Medicine Ohio State University Columbus, OH, USA

College of Veterinary Medicine Ohio State University Columbus, OH, USA William T.N. Culp, VMD, DACVS, ACVS Founding Fellow of Surgical Oncology

School of Veterinary Medicine University of California, Davis Davis, CA, USA Jonathan Dear, DVM, DACVIM (SAIM)

School of Veterinary Medicine University of California, Davis Davis, CA, USA Alice Defarges, DVM, DACVIM (SAIM)

Ontario Veterinary College University of Guelph Guelph, ON, Canada Alexander de Lahunta, DVM, PhD, DACVIM (Neurology), DACVP

College of Veterinary Medicine Cornell University Ithaca, NY, USA Sandra Diaz, DVM, MS, DACVD Department of Veterinary Clinical Sciences

Ohio State University Columbus, OH, USA

Serge Chalhoub, DVM, DACVIM (SAIM)

Stephen DiBartola, DVM, DACVIM (SAIM)

Faculty of Veterinary Medicine University of Calgary Calgary, AB, Canada

Ohio State University Columbus, OH, USA

Valérie Chetboul, DVM, PhD, DECVIM‐CA (Cardiology)

Lisa DiBernardi, DVM, DACVIM (Oncology), DACVR (Radiation Oncology)

National Veterinary School at Alfort Maisons‐Alfort, France

Gulf Coast Veterinary Specialists Houston, TX, USA

­List of Contributors

Alison Diesel, DVM, DACVD

Michelle Evason, DVM, DACVIM (SAIM)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Atlantic Veterinary College University of Prince Edward Island Charlottetown, PE, Canada

Pedro P. Vissotto de Paiva Diniz, DVM, PhD

Andrea Fischer, DVM, DECVN, DACVIM (Neurology)

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Keren E. Dittmer, PhD, BVSc, DACVP

School of Veterinary Science Massey University Palmerston North, New Zealand Robin Downing, DVM, MS, DAAPM, DACVSMR

The Downing Center for Animal Pain Management, LLC Windsor, CO, USA Yvonne Drechsler, PhD

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Marilyn Dunn, DVM, DACVIM (SAIM)

Centre Hospitalier Universitaire Vétérinaire University of Montreal Saint‐Hyacinthe, QC, Canada Adam E. Eatroff, DVM, DACVIM (SAIM)

ACCESS Specialty Animal Hospitals Culver City, CA, USA Karen Ehnert, DVM, MPVM, DACVPM

Veterinary Public Health Program Los Angeles County Department of Public Health Los Angeles, CA, USA Karen Eiler, DVM, MS, DACVIM (SAIM)

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Helen T. Engelke, BVSc, MPVM, DACVPM, MRCVS

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Ludwig‐Maximilians University Munich, Germany Daniel S. Foy, MS, DVM, DACVIM (SAIM), DACVECC

College of Veterinary Medicine Midwestern University Glendale, AZ, USA Linda A. Frank, DVM, MS, DACVD

College of Veterinary Medicine University of Tennessee Institute of Agriculture Knoxville, TN, USA Virginia Luis Fuentes, MA, VetMB, PhD, CertVR, DVC, MRCVS, DACVIM (Cardiology), DECVIM (Cardiology)

Royal Veterinary College University of London Hatfield, Herts, UK Silvia Funes, DVM, MS, DACVIM (SAIM)

VCA Bay Area Veterinary Specialists and Emergency Hospital San Leandro, CA, USA Jennifer L. Garcia, DVM, DACVIM (SAIM)

Sugar Land Veterinary Specialists and VetCompanion Houston, TX, USA Anna R. M. Gelzer, DVM, PhD, DACVIM (Cardiology), DECVIM‐CA (Cardiology)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA Robert Goggs, BVSc, PhD, DACVECC, DECVECC

Cornell University College of Veterinary Medicine Companion Animal Hospital Ithaca, NY, USA Sonya G. Gordon, DVM, DVSc, DACVIM (Cardiology)

VCA Great Lakes Veterinary Specialists Cleveland, OH, USA

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Amara H. Estrada, DVM, DACVIM (Cardiology)

Kinga Gortel, DVM, MS, DACVD

University of Florida Gainesville, FL, USA

Tri Lake Animal Hospital and Referral Centre Lake Country, BC, Canada

Amanda K. Elpiner, DVM, DACVIM (Oncology)

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Emily Gould, DVM, MS

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Deborah Greco, DVM, PhD, DACVIM (SAIM)

Desert Veterinary Specialists Palm Desert, CA, USA Julien Guillaumin, DVM, DACVECC

Colorado State University Fort Collins, CO, USA Jens Häggström, DVM, PhD, DECVIM‐CA (Cardiology)

Swedish University of Agricultural Sciences Uppsala, Sweden Ben Harris, MA, VetMB, CertSAM, MRCVS

Northwest Veterinary Specialists Sutton Weaver, Cheshire, UK

Rosanne Jepson, BVSc, MVetMed, PhD, DACVIM (SAIM), DECVIM-CA, PGCertVetEd, FHEA, MRCVS

Royal Veterinary College University of London London, UK Philip H. Kass, BS, DVM, MPVM, MS, PhD

Population Health and Reproduction Davis, CA, USA Heidi B. Kellihan, DVM, DACVIM (Cardiology)

University of Wisconsin Madison, WI, USA Robert Kemppainen, DVM, PhD

Auburn University Auburn, AL, USA Melissa Kennedy, DVM, PhD, DACVM

College of Veterinary Medicine University of Tennessee Knoxville, TN, USA Linda Kidd, DVM, PhD, DACVIM

Trina Hazzah, DVM, DACVIM (Oncology)

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Nicole A. Heinrich, DVM, DACVD

McKeever Dermatology Clinics Eden Prairie, MN, USA Patrick Hensel, Dr. Med.Vet., DACVD, DECVD

Tierdermatologie Basel, Switzerland Bradford J. Holmberg, DVM, MS, PhD, DACVO

Animal Eye Center Little Falls, NJ, USA Kate Hopper, BVSc, DACVECC, PhD

School of Veterinary Medicine University of California, Davis Davis, CA, USA Geraldine Hunt, BVSc, MVetClinStud, PhD, FACVSc

School of Veterinary Medicine University of California, Davis Davis, CA, USA Joanne L. Intile, DVM, DACVIM (Oncology) College of Veterinary Medicine

North Carolina State University Raleigh, NC, USA

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Lonnie J. King, DVM, MS, MPA, DACVPM

The Ohio State University Columbus, OH, USA Barbara E. Kitchell, DVM, PhD, DACVIM (Internal Medicine, Oncology)

VCA Veterinary Care Animal Hospital and Referral Center Albuquerque, NM, USA Marguerite F. Knipe, DVM, DACVIM (Neurology)

School of Veterinary Medicine University of California, Davis Davis, CA, USA Sandra N. Koch, DVM, MS, DACVD

College of Veterinary Medicine University of Minnesota St Paul, MN, USA Angela R. Kozicki, DVM, DACVIM (Oncology)

Bluepearl Veterinary Partners Southfield, MI, USA Marc S. Kraus, DVM, DACVIM (SAIM, Cardiology), DECVIM‐CA (Cardiology)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA

­List of Contributors

Orna Kristal, DVM, DACVIM (Oncology)

Veterinary Specialty and Emergency Hospital Wan Chai, Hong Kong Stephanie Kube, DVM, DACVIM (Neurology)

Veterinary Neurology and Pain Management Center of New England Walpole, MA, USA Kevin Kumrow, DVM, DACVIM (SAIM)

Orchard Park Veterinary Medical Center Orchard Park, NY, USA Nicolaas E. Lambrechts, DVM, DECVS, DACVSMR

College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO, USA Catherine Langston, VM, DACVIM (SAIM)

Department of Veterinary Clinical Sciences Ohio State University Columbus, OH, USA Patty Lathan, VMD, DACVIM (SAIM)

College of Veterinary Medicine Mississippi State University Mississippi State, MS, USA David Lipsitz, DVM, DACVIM (Neurology)

Veterinary Specialty Hospital San Diego, CA, USA Meryl P. Littman, VMD, DACVIM (SAIM)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA Ingrid Ljungvall, DVM, PhD, DECVIM‐CA (Cardiology)

Swedish University of Agricultural Sciences Uppsala, Sweden Sam N. Long, BVSc, PhD, DipECVN

Centre for Animal Referral and Emergency Melbourne, Australia Maya Lottati, DVM, PhD, DACVIM (SAIM)

Caroline Mansfield, BSc, BVMS, MVM, PhD, MANZCVS, DECVIM-CA

Melbourne Veterinary School University of Melbourne Melbourne, Australia Christopher L. Mariani, DVM, PhD, DACVIM (Neurology)

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA Steven A. Martinez, DVM, MS, DACVS, DACVSMR

College of Veterinary Medicine Washington State University Pullman, WA, USA Tracey McNamara, DVM, DACVP

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Mathew Mellema, DVM, PhD, DACVECC

School of Veterinary Medicine University of California, Davis Davis, CA, USA Heather A. Towle Millard, DVM, MS, DACVS‐SA

Blue Pearl Pet Hospital Overland Park, KS, USA Paul E. Miller, DVM, DACVO

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA Milan Milovancev, DVM, DACVS‐SA

School of Veterinary Medicine Oregon State University Corvallis, OR, USA Ralf S. Mueller, Dr. Med.Vet., DACVD, FANZCVSc

Centre for Clinical Veterinary Medicine Ludwig‐Maximilians‐University of Munich Munich, Germany Reto Neiger, Dr.Med.Vet., PhD, DACVIM (SAIM), DECVIM-CA

TrueCare for Pets Studio City, CA, USA

Small Animal Clinic Hofheim Hofheim, Germany

Katharine F. Lunn, BVMS, MS, PhD, MRCVS, DACVIM (SAIM)

Laura L. Nelson, DVM, MS, DACVS

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA

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­List of Contributors

Rhett Nichols, DVM, DACVIM (SAIM)

Antech Diagnostics Farmingdale, NY, USA Christine B. Oakley, DVM

VCA Veterinary Specialists of the Valley Woodland Hills, CA, USA Ben O’Kelley, DVM, DACVECC

BluePearl Veterinary Partners Tampa, FL, USA Emma O’Neill, BSc, BVSc, PhD, DSAM, DECVIM‐CA, MRCVS

School of Veterinary Medicine University College Dublin Dublin, Eire M. Lynne O’Sullivan, DVM, DVSc, DACVIM (Cardiology)

Atlantic Veterinary College University of Prince Edward Island Charlottetown, PEI, Canada Tina Jo Owen, DVM, DACVS

Washington State University Pullman, WA, USA Mark A. Oyama, DVM, MSCE, DACVIM (Cardiology) Department of Clinical Sciences and Advanced Medicine

University of Pennsylvania Philadelphia, PA, USA

Romain Pariaut, DVM, DACVIM (Cardiology), DECVIM-CA (Cardiology)

College of Veterinary Medicine Cornell University Ithaca, NY, USA Sue Paterson, MA, VetMB, DVD, DECVD, FRCVS

Veterinary Dermatologicals Altrincham, Cheshire, UK Maria Grazia Pennisi, DVM, PhD

Specialist Applied Microbiology University of Messina Messina, Italy Nathan Peterson, DVM, DACVECC

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Tom Phillips, DVM, MS, PhD

Deceased Formerly College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Simon R. Platt, BVM&S, FRCVS, DACVIM (Neurology), DECVN

College of Veterinary Medicine University of Georgia Athens, GA, USA Michael Podell, MSc, DVM, DACVIM (Neurology)

Chicago Veterinary Emergency and Specialty Center Chicago, IL, USA Lauren Askin Quarterman, DVM, DACVR (Radiation Oncology)

PetCure Oncology San Jose, CA, USA Jessica Quimby, DVM, PhD, DACVIM (SAIM)

Ohio State University Veterinary Medical Center Columbus, OH, USA Jacquie S. Rand, BVSc, DVSc, MANZVS, DACVIM (SAIM)

School of Veterinary Science University of Queensland Gatton, Queensland, Australia and Australian Pet Welfare Foundation Kenmore, Queensland, Australia Robert B. Rebhun, DVM, PhD, DACVIM (Oncology)

Department of Surgical and Radiological Sciences University of California, Davis Davis, CA, USA Nicki Reed, BVM&S, CertVR, DSAM (Feline), DECVIM‐CA, MRCVS

Veterinary Specialists West Lothian, Scotland, UK Alexander M. Reiter, Dipl.Tzt., Dr.Med.Vet., DAVDC, DEVDC

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA Lisa V. Reiter, DVM, DACVD

McKeever Dermatology Clinics Eden Prairie and Inver Grove Heights, MN, USA Teresa Rieser, DVM, DACVECC

Veterinary Specialty Care Mt. Pleasant, SC, USA Steven Rosenthal, DVM, DACVIM (Cardiology)

CVCA Cardiac Care for Pets Towson, MD, USA

­List of Contributors

Jan Rothuizen, DVM, PhD, DECVIM‐CA

Erin M. Scott, DVM, DACVO

Department of Clinical Science of Companion Animals Utrecht University Utrecht, The Netherlands

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Elizabeth A. Rozanski, DVM, DACVIM (SAIM), DACVECC

VCA Alameda East Veterinary Hospital Denver, CO, USA

Tufts University Medford, MA, USA Craig Ruaux, BVSc (Hons), PhD, MACVSc, DACVIM (SAIM)

School of Veterinary Science Massey University Palmerston North, New Zealand Erin E. Runcan, DVM, DACVT

Department of Veterinary Clinical Sciences Ohio State University Columbus, OH, USA David Ruslander, DVM, DACVIM (Oncology), DACVR (Radiation Oncology)

Veterinary Specialty Hospital of the Carolinas Cary, NC, USA Silke Salavati, Dr.Med.Vet., PhD, DipECVIM‐CA, MRCVS

Royal School of Veterinary Studies University of Edinburgh Edinburgh, Scotland Ashley B. Saunders, DVM, DACVIM (Cardiology)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Brian A. Scansen, DVM, MS, DACVIM (Cardiology)

Katherine Scott, DVM, DACVIM (SAIM)

Christine M. Serafin, DVM, DACVIM (SAIM)

The Animal Surgical Center of Michigan Flint, MI, USA Stephanie L. Shaver, DVM, DACVS

School of Veterinary Medicine University of California, Davis Davis, CA, USA Marco A. Coutinho da Silva, DVM, MS, PhD, DACT

Ohio State University Columbus, OH, USA Nicole Smee, DVM, MS, DACVIM

Las Vegas Veterinary Specialty Center Las Vegas, NV, USA Laia Solano‐Gallego, DVM, PhD, DECVCP

Facultat de Veterinária Universitat Autònoma de Barcelona Barcelona, Spain Maria M. Soltero‐Rivera, DVM, DAVDC

VCA San Francisco Veterinary Specialists San Francisco, CA, USA Mitchell D. Song, DVM, DACVD

Colorado State University Fort Collins, CO, USA

VETMED Specialty Referral and 24‐Hour Emergency Care Veterinary Hospital Phoenix, AZ, USA

Jennifer R. Schissler, DVM, MS, DACVD

Joachim Spergser, Dipl.Tzt., Dr. Med.Vet., DECVM

College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO, USA Peggy L. Schmidt, DVM, MS, DACVPM

College of Veterinary Medicine Kansas State University Manhattan, KS, USA

Institute of Microbiology University of Veterinary Medicine Vienna, Austria Rebecca L. Stepien, DVM, MS, DACVIM (Cardiology)

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA

Nick A. Schroeder, DVM, DACVIM (Cardiology)

Tammy Stevenson, DVM, DACVIM

LeadER Animal Specialty Hospital Cooper City, FL, USA

Veterinary Specialty Hospital San Diego, CA, USA

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­List of Contributors

Susanne M. Stieger‐Vanegas, DVM, PhD, DECVDI

Avenelle I. Turner, DVM, DACVIM (Oncology)

Carlson College of Veterinary Medicine Oregon State University Corvallis, OR, USA

Veterinary Cancer Group Culver City, CA, USA

Jason W. Stull, VMD, MPVM, PhD, DACVPM

Midtown Animal Hospital Gering, NE, USA

Department of Veterinary Preventive Medicine Ohio State University Columbus, OH, USA Jan S. Suchodolski, MedVet, Dr. Med.Vet., PhD, AGAF, DACVM

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Karen Lynn C. Sueda, DVM, DACVB

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Steven Suter, VMD, MS, PhD, DACVIM

Canine Bone Marrow Transplant Unit North Carolina State University College of Veterinary Medicine Raleigh, NC, USA Jane E. Sykes, BVSc, PhD, DACVIM (SAIM)

Jerry J. Upp, DVM

Shelly L. Vaden, DMV, PhD, DACVIM (SAIM)

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA Katrina R. Viviano, DVM, PhD, DACVIM (SAIM), DACVCP

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA J. Scott Weese, DVM, DVSc, DACVIM (SAIM)

Ontario Veterinary College University of Guelph Guelph, ON, Canada Raegan Wells, DVM, DACVECC

Phoenix Veterinary Referral and Emergency Center Phoenix, AZ, USA Joanna Whitney, BSc, BVSc, PhD, MVetStud, MACVS (SMAnimMed, ECC)

University of California, Davis Davis, CA, USA

Small Animal Specialist Hospital North Ryde, NSW, Australia

Séverine Tasker, BSc, BVSc (Hons), PhD, DSAM, DACVIM‐CA, PGCertHE, MRCVS

Wayne E. Wingfield, MS, DVM, DACVS, DACVECC

Bristol Veterinary School University of Bristol Bristol, UK Katie Tolbert, DVM, PhD, ADCVIM (SAIM)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Elizabeth E. Toops, DVM, MS, DACVD (Dermatology)

Virginia Veterinary Specialists Charlottesville, VA, USA Sheila M.F. Torres, DVM, MS, PhD, DACVD

College of Veterinary Medicine University of Minnesota St Paul, MN, USA

Department of Clinical Sciences Colorado State University Fort Collins, CO, USA Zachary M. Wright, DVM, DACVIM (Oncology)

VCA Animal Diagnostic Clinic Dallas, TX, USA Panagiotis G. Xenoulis, DVM, Dr. Med.Vet., PhD

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX

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About the Companion Website This book is accompanied by a companion website: www.wiley.com/go/bruyette/clinical The website includes: Podcasts

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Section 1 Evaluation and Management of the Patient

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1 The Concept of One Medicine Lonnie J. King, DVM, MS, MPA, DACVPM Department of Veterinary Preventive Medicine, The Ohio State University, Columbus, OH, USA

Today’s world is rapidly changing, complex, and progressively more interconnected. The convergence of people, animals, and our environment has created a new dynamic characterized by a profound and unprecedented interdependence in which the health of all three domains is now inextricably linked and elaborately connected. Over the last three decades, approximately 75% of new emerging human diseases have been zoonotic. The human–animal interface is expanding, accelerating, and becoming increasingly more consequential. At the same time, we have permanently altered a significant portion of our environment, ecosystems, and habitats and have created a new ecological milieu that is changing both the conditions of our human–animal interface and the conditions for microbial adaptation and the emergence and reemergence of infectious diseases worldwide. Our new interdependence includes social, economic, political, and biological conditions that are creating new threats to the health of people, animals, and our environment. We can no longer focus on these threats separately from each other. Our contemporary challenge is to create and implement a new mindset and strategies to address our threats to health based on a holistic and integrated approach with a special emphasis on prevention and attacking problems closer to their origin that is often within the animal and environmental domains. This approach is the essence of the concept of One Health. One Health can be defined as the collaborative effort of multiple disciplines – working locally, nationally, and globally to attain optimal health for people, animals, and our environment. The scope of One Health is ­impressive, broad, and growing. Much of the recent focus on One Health has been limited to emerging infectious diseases yet the concept clearly embraces environmental and ecosystem health, social sciences, biodiversity, ecology, noninfectious diseases and chronic diseases and much more.

While the concept of One Health is not new, it has enjoyed a new recognition based on today’s complex challenges to our health. The veterinary profession is especially well equipped through our broad training in herd health, comparative medicine, epidemiology, problem solving, and disease ecology to play an important role in implementing new One Health strategies.

­Factors Driving One Health In a publication entitled Microbial Threats to Health: Emergence, Detection, and Response, authors from the Institute of Medicine suggested that a group of factors have simultaneously converged to create a “perfect microbial storm” [1]. The most important of these ­factors include: ●● ●● ●● ●● ●● ●● ●● ●●

adaptation of microbes global travel and transportation host susceptibility intent to do harm climate change economic development and land use human demographics and behavior a breakdown of both public and animal health infra­structures.

­The Human Domain The world population has a growth rate of 1.2% per year and the next century will represent a period of exponential growth. We add approximately 10 000 people per hour to our global population every day. Approximately 90% of the world’s population growth is occurring in the developing countries where we are most concerned about a lack of adequate public and animal health infrastructure. In addition, almost 1 billion people live in

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 1  Evaluation and Management of the Patient

periurban or slum settings in the developing world’s largest cities and these sites are where the most rapid growth in our human populations will continue. Concurrently, there is unprecedented immigration, translocation, and movement of people worldwide. Unique diasporas have emerged and there are large numbers of immunocompromised individuals dispersed throughout the US and global populations who are especially susceptible to infections involving food‐ and water‐borne illnesses.

­The Animal Domain While there is also legitimate concern about the approximately 800 million people who are undernourished, we are concurrently observing a relative increase in wealth in the developing world and as per capita incomes rise, people eat more calories and consume different products, including a demand for meat and protein from animal sources. Thus, a new agricultural phenomenon is emerging  –  the Livestock Revolution; we will need to increase protein from animal sources by 50% over the next several decades. We will also need to increase food‐ animal production with minimal impacts on the environment and our ecosystems. Veterinarians will be engaged with ecologists to help ensure the sustainability of our lands. At the same time, companion animals and pets are also rapidly increasing globally and there are untold numbers of wildlife that are increasingly involved at the human– animal interface and responsible for carrying and transmitting more and more zoonotic diseases. West Nile virus, Lyme disease, Nipah, Hendra, and hantaviruses are some important examples. With a new appreciation of the need for detecting and identifying human infectious disease threats at animal and ecosystem level, our scope of disease surveillance must also be expanded to include these domains. Currently, 80% of select agents are zoonotic which further emphasizes the need for integrating animal disease surveillance into human disease surveillance programs. In addition to the benefits of earlier detection and warning, surveillance within these domains will give us new insights and analyses on disease prediction and new possibilities to prevent future disease occurrences and exposures. Thus, animal disease surveillance has taken on a new importance and relevance. Until we address the underlying factors that lead to disease emergence and reemergence, we will continue to address infectious disease problems one at a time and based on reactive responses. A new commitment is necessary to refocus our efforts to prevent and control diseases in ways that also address these underlying conditions and the animals, animal products, and their vectors.

For example, human exposure to Borrelia burgdorferi, the etiologic agent of Lyme disease, is maintained and spread based on the complex relationships among wildlife populations, tick life cycles, host preferences, habitat, ecosystem, and landscape design and other dynamics of this disease ecology. Effective prevention and control may need to focus on ecologic interventions rather than treating the human illness after exposure and infection. Thus, rather than a narrow focus on humans and their interactions with pathogens, we need to focus on the dynamic interplay among humans, animals, and animal products, and our changing environment  –  in other words, a One Health approach.

­Role for Companion Animal Experts The purpose of including this introduction on One Health for a text on internal medicine is to help create a new awareness of the concept across the veterinary profession and to suggest that companion animal practitioners also have new opportunities, as well as obligations, to support public health in the future. Practitioners can be extremely helpful and contribute in the following ways: ●●

●●

●●

●●

●●

enhance surveillance for emerging and zoonotic diseases identify cases of zoonoses that will then prevent human illnesses educate clients regarding One Health and zoonotic diseases be involved in a potential surge capacity in case a significant epidemic or natural disaster calls for your support promote the human–animal bond but with knowledge of both the benefits and potential negative impacts of the human–animal interface, including wildlife.

­Conclusion Our global interdependence and growing convergence of animals, animal products, and people ensure that diseases will continue to be a significant threat to health. There is nothing on the horizon suggesting that the era of the perfect microbial storm will lessen or be abated. The factors creating this reality are well entrenched and emerging and reemerging zoonoses are growing and expanding in scope, scale, and impact. These diseases now have a much greater impact and influence beyond health. Global outbreaks of SARS and influenza have demonstrated how such diseases create immense economic losses in goods, services,

1  The Concept of One Medicine

travel, and trade in addition to their profound health costs and consequences. Contemporary societal issues such as global trade and commerce, movements and migrations of populations, changes and insults to our ecosystems, population growth in developing countries, the Livestock Revolution, and disruptions of the environment are helping to create a complex, nonlinear and disruptive world that is riskier and impacts health in the domains of people, animals, and our environment. The ability to improve and optimize health in the future will depend on working in these three domains concurrently, holistically, and synergistically. A One Health mindset and accompanying actions are now a necessity and no longer just a concept. In order to reconcile our significant contemporary challenges and threats with our habitual, narrow and traditional thinking in medicine, new levels of thinking across disciplines and professions will become the new normal. Veterinarians are becoming essential leaders in this

movement and a new concept of community health, including veterinary medicine, is emerging. Public health and animal health are no longer the domain of any single discipline, profession or boundary; rather, they require the work of epidemiologists, clinicians, ecologists, veterinarians, entomologists, engineers, and many more experts coming together to provide new insights and perspectives using new scientific and technological tools. Addressing, predicting, and preventing diseases certainly demand the richness of new scientific and medical teams. For veterinary medicine, our horizon has never been richer nor our possibilities more promising. Our profession’s challenge is to be relevant and meet the changing needs of society regarding public health, global food systems, ecosystem management, and biomedical research. One Health is the framework, mindset, and new collective purpose to help define our expanding responsibilities, including, and beyond, traditional clinical medicine.

­Reference 1 Institute of Medicine Committee on Emerging Microbial

Threats to Health in the 21st Century, Smolinski M, Hamburg MA, Lederburg J. Microbial Threats to Health:

Emergence, Detection, and Response. Washington, DC: National Academies Press, 2003.

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2 Statistical Interpretation for Practitioners Philip H. Kass, BS, DVM, MPVM, MS, PhD Department of Population Health and Reproduction, School of Veterinary Medicine, University of California, Davis, CA, USA

The need for an understanding of how to conduct statistical analyses and, more importantly, how to interpret them derives from a natural tension between aspiration and reality: the desire to make encompassing statements concerning the characteristics and causal properties about populations of animals, juxtaposed with the inability to study more than a small sample of them. Statistical inference therefore provides the necessary linkage between using samples to make inferences about populations.

­External Validity It is often taken as a matter of faith that studies conducted in relatively circumscribed subpopulations (such as a cohort of patients seen at an individual hospital in a defined period of time) can have relatively generalizable findings. For example, a hospital‐based study examining the relative clinical efficacy of two or more chemotherapeutic regimens to treat newly diagnosed canine lymphoma by inducing remission may motivate the authors to make recommendations for adoption well beyond the hospital’s patient catchment area. When are such generalizations justified? Sampling of populations is required to scientifically justify extrapolating results from sample‐based studies to target populations. Thus, to make scientific inferences about a population, it is necessary to study a representative sample. In many cases, the sample size need not be particularly large, and can be obtained through random (or more complex) sampling schemes. The process of random sampling ensures representative selection, and that in turn provides the key link between a study sample and a target population. Because few studies are actually conducted using true sampling of a target population, the ability to generalize study findings (that is, having “external validity”) typically depends on prior medical belief and knowledge, as well as

key assumptions. For example, the initial epidemiological research into the association between tobacco smoking and lung cancer was performed in the 1950s in a cohort of male British doctors [1]. Technically, the findings of this research strictly applied to the entire population of male British doctors who were contemporaries of the study subjects. The choice to generalize these findings – namely, that the incidence of lung cancer was many‐fold higher among smokers than nonsmokers  –  to other populations rested on key assumptions motivated by scientific reasons independent of the actual research. These assumptions included that the effect of tobacco smoking on lung cancer incidence should not meaningfully vary by gender, occupation, country of origin, ethnic identity, and birth cohort. Nothing in the original research could have provided evidence to support these assumptions; nevertheless, they helped create the basis for the landmark 1964 report in the US officially affirming a causal link between tobacco smoking and lung cancer [2]. A more recent example of the dangers of extrapolating study results to nonstudy populations can be found in an article on the association between mitotic index (MI) and survival in dogs diagnosed with mast cell tumors [3]. The authors found a substantially lower survival in dogs seen at a California veterinary medical teaching hospital whose MI was 5 or fewer versus those whose MI was 6 or greater. In contrast, Elston et al. performed similar analyses on dogs from Brazil and recommended somewhat different MI cut‐off values [4]. The original authors responded by underscoring the difficulty of externally validating studies: This underscores the fallibility of classification schemes in clinical veterinary medicine: they may work well in a study population and its corresponding reference population but may not perform nearly as well in a target population of inherently different animals or where measurement standards may not necessarily be completely

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 1  Evaluation and Management of the Patient

concordant. The latter can be influenced by how different pathologists measure, count, and ultimately determine MI, and whether such determinations are so precise as to be perfectly replicable by others. Such studies are then relegated to determining average effects across potentially heterogeneous patient populations even within a community, much less between countries. Thus, patient characteristics such as age, breed, sex, and owner propensity for diagnosis (including biopsy) would be expected to vary, perhaps substantially, between institutions such as ours and Elston et al.’s. [5] These illustrations should presumably compel veterinarians to be circumspect about relying too heavily on any single study, particularly one conducted in a geographically restricted region, and for authors to exercise restraint in advocating wider than justified implications for a single study. Although the heterogeneity existing across populations may have a negligible impact on the universality of study findings, a cautious approach to generalizability is often warranted.

­Internal Validity Unlike external validity, which relies on information not usually obtained in a study to make inferences beyond the bounds of a study population, internal validity depends on the relative presence or absence of two sources of error pervasive to all clinical studies: imprecision (random error) and invalidity (systematic error). Imprecision has multiple formal statistical definitions and interpretations, but can best be understood as how variable the results are expected to be from multiple studies measuring the same quantity. As an example, if a veterinarian were to repeatedly weigh a patient using a scale, precision would imply obtaining similar weights each time; conversely, imprecision implies considerable disagreement in weights with each attempt. Standard deviations, standard errors, variances, and ranges are all statistical measures for quantifying imprecision. We can make two broad qualitative statements about variability without resorting to mathematical formulas: (1) the further apart observations are from each other, the more imprecise any summary statistic based on them will be; and (2) the larger a study is, the more precise any statistic derived from it will be. This has an intuitive appeal: the findings from a large study are more likely to be accepted as more precise than those from a small study. Provided there are no biases, the source of statistical variability in a study is typically ascribed to random error.

It would be a grievous error, however, to accept a study’s outcomes solely on the basis of high precision. Invalidity, another source of error in studies and equivalently called bias, is a distinctly different statistical measure, and represents the disparity between what one empirically measures in a study and what one strives to measure (namely, factual truth). Returning to the previous example, if a hospital scale is miscalibrated, then regardless of how precise or imprecise the replicate measurements are, the average weight will invariably be incorrect (biased), with the degree of the bias proportional to the degree of miscalibration. All medical research is potentially susceptible to imprecision and bias, and it is only through circumspect study design that these errors can be prevented or controlled. Such errors exist on a quantitative continuum (parenthetically often reduced to crude descriptors, such as “very” imprecise or “small” bias), so it becomes important to recognize that all studies must be judged by both error criteria and not just one (or neither). Such lack of circumspection is pervasive, even in academic settings, and helps explain the often contradictory results one so often reads about in the popular media [6]. It also underscores the often unappreciated theme that even very large‐sample studies are susceptible to bias, and despite their exceedingly high precision can conceivably be as misleading as a much smaller study.

­Hypothesis Testing Perhaps the most obvious manifestation of the use of statistical analysis in clinical research is through much misunderstood hypothesis testing. Although investigators and clinicians typically want to know if the differences between study groups are real, or if the association between risk factors and health outcomes is real, hypothesis tests are unable to answer these questions. Moreover, the P‐values so ubiquitously reported in clinical articles fail to inform readers about the probabilities that measured differences or associations are real. In reality, their interpretation is surprisingly counterintuitive. Consider the example of comparing the average (mean) alanine aminotransferase (ALT) levels between young dogs and geriatric dogs. The average in young dogs is 40 μ/L, and in geriatric dogs is 80 μ/L. The null hypothesis is that the averages in the two groups are equal; the two‐ sided (meaning that either group could have a higher/ lower average than the other) alternative hypothesis is that they are unequal. A two‐group Student’s t‐test is performed, and a P‐value of 0.10 is calculated. If the level of significance (α) is 0.05, then the difference is not statistically significant. What can one conclude from this?

2  Statistical Interpretation for Practitioners

First, it would be an error to state that there was “no difference” between the groups. Clearly, there was a difference (of 40 μ/L). It would, however, be correct to state that because the null hypothesis of no group difference was not rejected, there was no significant difference between the groups, assuming the assumptions of the statistical model are correct (because (P = 0.10) > (α = 0.05)). Second, the P‐value does not provide a quantitative assay of the probabilities that the null or alternative hypotheses are correct. Conventional (superiority) hypothesis testing is predicated on the veracity of the null hypothesis, and so does not provide any assessment of its truth. Instead, the P‐value addresses an entirely different question: how likely (probabilistically speaking) is it that one would observe differences at least as large as the one found in the study (40 μ/L) when the null hypothesis is true? In other words, instead of the P‐value equaling the probability that the null hypothesis is true given the data observed in the study, it provides the probability of observing the difference in the data observed (or more extreme) given the null hypothesis is true. It follows that “large” P‐values indicate substantial concordance with the null hypothesis, while “small” P‐values indicate poor concordance with the null hypothesis (presumably motivating its rejection in favor of its alternative). Third, a P‐value is only numerically correct under a particular statistical model. With the Student’s t‐test example, the underlying model assumes that the ALT values in each group are approximately normally distributed. If the assumption of normality is violated, however, the P‐value will no longer be correct; the greater the departure of the data distribution from normality, the more incorrect the P‐value will be. Fourth, another assumption is that the study data are independent, meaning that knowing one individual’s ALT value does not allow the ability to predict another individual’s ALT value. In this example, such an assumption is reasonable when each individual contributes only one ALT value. However, when replicates from a single individual are included, it is plausible to assume that knowing an individual’s value at one time can better predict the same individual’s value at another time than the value from a different individual. Such a violation of the data independence assumption leads to the estimation of an incorrect P‐value; typically, the use of correlated (nonindependent or dependent) data underestimates P‐ values, and hence more type I statistical errors (improperly rejecting the null hypothesis) arise. Fifth, it is important to recognize that statistical analysis is more than the analysis of single numbers (i.e., point estimates, as in averages) in groups; instead, it is more correctly described as the analysis of variability. For this reason, the presence or absence of statistical significance

is more than just reflective of the magnitude of differences or associations and the chosen level of significance; it is also a function of the variance of the point estimates. Because such variances are inversely proportional to sample sizes, two studies with identical differences or associations can have completely different P‐values: the smaller study’s differences could be nonsignificant, while the larger study’s differences may be significant. Sixth, it directly follows that any group differences or associations can eventually be made statistically significant if the study size becomes sufficiently large. To illustrate this, two random samples of 25 individuals each were created, one assuming blood hemoglobin was normally distributed with a mean of 15 g/dL and standard deviation of 2 g/dL, and the other with a mean and standard deviation of 15.1 and 2 g/dL, respectively. No one would seriously argue that a hemoglobin difference of 0.1 g/dL is clinically important, and indeed it is not significant at α = 0.05 (P = 0.79). However, if the two groups were constructed to have 2500 individuals each, with the same means and standard deviations, this same clinically unimportant difference (0.1 g/dL) becomes statistically significant (P = 0.023). This underscores an important distinguishing principle of statistical analysis: statistical significance is distinctly different from and does not imply medical importance. In a large enough study, even trivial and unimportant differences can become statistically significant; in a small study, differences that appear to be worthy of medical pursuit may be statistically insignificant. In recognition of this principle, alternative methods of hypothesis testing have been developed that instead of examining superiority of one group over another, evaluate whether groups are noninferior or equivalent based on a determination of what constitutes an acceptably important difference or association [7]. Finally, each decision to reject or not reject a null hypothesis following hypothesis testing is prone to error. What is often unappreciated is that the more tests that are performed, the greater the probability that at least one decision will be incorrect. In a clinical setting, this is perhaps most manifest when performing clinical laboratory testing panels to screen for hematologic or chemical abnormalities in blood. Reference ranges for blood parameters are typically constructed to encompass approximately 95% of normal animals, implying that 5% of normal animals will have values falling outside the ranges. When a reference range is appropriate for a patient’s age, sex, and any other factors that can influence a particular blood parameter, it is reassuring that a normal animal’s value will fall within the reference range 95% of the time. However, a universally accepted practice is to run laboratory panels simultaneously evaluating many parameters. Suppose, for example, that a blood chemistry

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panel contains 30 different parameters to evaluate. Although each separate parameter has a 5% probability of being incorrect in a normal animal, when considering all parameters the probability of at least one error rises to 78.5%. In other words, a veterinarian should be surprised if at least one parameter does not fall outside the reference range even in a healthy patient. Another related problem arises when using screening tests with imperfect properties (false‐positive and ‐negative results) in populations with low prevalences of illnesses. Even when the test properties are excellent, with error rates less than 5%, if a disease is rare enough in a population of patients being screened, there can paradoxically be more false‐positive results than true‐positive results. This should be discomforting to veterinarians whose clinical decisionmaking depends on the veracity of diagnostic testing. One approach to resolving this issue (apart from running the test again to see if the result was aberrant) is to use screening tests judiciously, reserving them for patients with a heightened suspicion of actual illness. In other words, clinical judgment and experience are as integral to the use and interpretation of diagnostic tests as contrasting the actual test results to reference ranges. Such circumspection serves to elevate the likelihood of illness in the patient being tested, or analogously the prevalence of the illness in the population being screened. This will contribute to making fewer unjustified conclusions about abnormal results in otherwise healthy patients.

­Parametric versus Nonparametric Tests All statistical tests have assumptions underlying their use. Those that assume that data follow a particular probability distribution function (also known as the sampling model) are called parametric tests (by virtue of the fact that the functions have one or more parameters in them). Probably the most common distribution function used in parametric statistical analyses is the normal distribution (with two parameters: the mean and standard deviation). As noted earlier, Student’s two‐group t‐test assumes that the data in each group are approximately normally distributed. Other well‐known parametric tests also rely on normality: the paired t‐test assumes the differences are approximately normally distributed; analysis of variance and least‐squares linear regression assume the model residuals (the difference between observed and predicted values) are approximately normally distributed. How can a clinician know if data are normally distributed? Various approaches exist, including distributional tests (for example, Shapiro–Wilk, Shapiro–Francia,

Kolmogorov–Smirnov tests) and plots (for example, normal probability plots). Such tests assume data are normally distributed; rejection of the null hypothesis indicates incompatibility with the null hypothesis of normality. The problem with such tests is that they are notoriously inadequate to detect departures from normality when sample sizes are small. For example, consider the following six data points: 100, 100, 80, 70, 50, 50. This is a U‐shaped distribution and so is clearly not normally distributed. Nevertheless, a Shapiro–Wilk test fails to reject the null hypothesis that the data are normally distributed when α = 0.05 (P = 0.22), so if we relied on this test result we would erroneously conclude that the sample data derived from a population with a normal distribution. Interestingly, if this dataset is doubled in size (n = 12), then the Shapiro–Wilk test would become significant (P = 0.019) and the null hypothesis of normality would be rejected. When sample sizes are small or when normality is an untenable assumption, then alternative statistical tests are sometimes available. These tests are called nonparametric tests, because they do not assume that data follow a particular distribution, and hence are also known as distribution‐free tests. Table  2.1 contains a list of the most commonly used nonparametric tests, as well as their equivalent parametric test (if one exists). These tests have more statistical power (for example, the ability to reject the null hypothesis when it is false) than parametric tests when data do not conform to a parametric distribution (such as normality), and are almost as powerful as parametric tests when the data do conform to a parametric distribution. When the assumption of normality is questionable based on statistical tests or the sample size is too small for reliable testing, nonparametric tests are preferable whenever possible.

­Describing Data Distributions For data that are approximately normally distributed, it is customary to describe the center of the distribution with the sample average (i.e., mean), and to describe the dispersion around the mean with either the standard deviation or the standard error of the mean (i.e., the standard deviation divided by the square root of the sample size). When data are not normally distributed, however, it is generally advisable to describe the data with statistics that do not assume a symmetric distribution around the mean. Preferable measures to use include medians, ranges, and percentiles of the data (for ­example, the 2.5th and 97.5th percentiles provide a 95% ­percentile‐based interval).

2  Statistical Interpretation for Practitioners

Table 2.1  Commonly used parametric and equivalent nonparametric statistical tests in biomedical research. Types of data

Continuous

Categorical

Type of statistical problem

Parametric test

Nonparametric test

Two independent samples

Student’s t

Wilcoxon‐Mann‐Whitney

Two dependent samples

Paired t

Wilcoxon signed rank

Three or more independent samples

Analysis of variance (one‐way)

Kruskal‐Wallis

Three or more ordered independent samples

Jonckheere‐Terpstra

Unordered 2 × 2 table

Fisher’s exact

Unordered R × C tablea Single ordered R × C table

Ordinal

Pearson’s Chi‐square a

Kruskal‐Wallis

Doubly ordered R × C tablea   Two dependent samples           K independent ordered proportionsb K dependent samples

                  Repeated measures analysis of variance (one‐way)

Jonckheere‐Terpstra   McNemar’s (two levels) Marginal homogeneity (three or more levels)       Cochran‐Armitage trend   Friedman

Association of two samples

Pearson correlation

Spearman correlation

a

 R = number of rows, C = number of columns.  K = number of proportions or samples, K > 2.

b

Frequency

Figure 2.1  Histogram showing a hypothetical asymmetric data distribution. The green line illustrates the poor fit of a normal (Gaussian) distribution to the data.

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To illustrate this point, Figure 2.1 shows a histogram of a data distribution that is clearly not normally distributed. Superimposed over the histogram is a curve of a normal distribution whose shape is completely determined by the mean and standard deviation of the actual data. Because the curve fails to closely approximate the histogram, the mean and standard deviation used to

c­ reate the curve similarly fail to describe the center and dispersion of the data. For example, the mean = 7.5 while the median (i.e., the 50th percentile) is only 6.0. Similarly, the range of the data extends from 1 to 24, but a 95% confidence interval of the data distribution based on the mean and standard deviation would have a lower bound less than 0, which is less than the smallest possible

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observed value. This underscores the point that when data do not conform to a particular parametric distribution, it is preferable to use a method of describing the data that does not depend on specifying a particular distribution. Graphically representing such data is likewise inappropriate when using bar charts based on means and error bars using standard deviations or standard errors. One indispensable way of graphically portraying such data is through the use of a box and whiskers plot, as shown for the data above in Figure 2.2. The advantage of such graphs is that they are constructed based only the data, and not on any distributional assumptions (such as normality). The lower and upper ends of the box correspond to the 25th and 75th percentiles respectively; the horizontal line within the box corresponds to the 50th percentile (i.e., the median), the bars (i.e., the “whiskers”) above and below the box contain all data above and below 1.5 times the vertical width of the box (i.e., the interquartile range), and the individual points beyond the whiskers represent extreme (outlying) values. Figure  2.2 reinforces the observation from the histogram that the data are skewed towards higher values, which causes the mean to be larger than the median. This type of plot is also valuable in assessing symmetry of a data distribution (asymmetric distributions should not be described with

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Figure 2.2  Box and whiskers plot for data shown in Figure 2.1. Lower and upper ends of the box represent the 25th and 75th percentiles, respectively; the orange line within the box equals the 50th percentile (median), the bars (“whiskers”) above and below the box contain all data above and below 1.5 times the interquartile range, and individual points beyond the whiskers represent outliers.

means and standard deviations) and for identifying outliers that may represent data errors. Another common error arises when replicate measurements are taken from the same patient over time. This arises, for example, when two or more treatments are successively tested in patients with before/after comparisons, or when monitoring blood values over time. As mentioned earlier, n replicate measurements taken from a single individual cannot be treated as equivalent to single measurements coming from n different individuals. Figure 2.3 illustrates this principle by examining before/ after measurements taken on five patients at different times. Both panel 1 (left) and panel 2 (right) contain the identical 10 points but panel 1 ignores the pairing within individuals, and a regression line fitted to the 10 points suggests a negative correlation between the measurements over time. Panel 2, in contrast, links the before/after pairs within individuals and an entirely different picture emerges: every individual’s values actually increased over time. Ignoring such pairing not only obscures actual effects but also in cases such as this suggests a paradoxical reversal of effects. Therefore, in studies where two or more measurements are taken from the same patients over time, it is important to not combine the patients’ values at individual time points through the use of histograms, boxplots, or other graphical representations. Panel B is often called a “spaghetti plot” because it can, with larger numbers of individuals, take on an appearance of many intertwined lines across two or more times.

­Measures of Association and Effect As noted earlier, the construct and interpretation of the P‐value is unsatisfying, if not bewildering to consumers of statistical analyses who are not well versed in their subtleties. The preoccupation with statistical testing in the medical literature should presumably be subordinate to and supplanted by a more favorable inclination towards estimation: the mathematical calculation of statistics that measure the magnitude of differences and associations in data. The imperative for this is no more evident than in the often seen but erroneous exposition that “no differences were found from a set of analyses because none were statistically significant.” Among the most common intuitively appealing measures of association are those conjoined from descriptive measures of prevalence and incidence. Prevalence Prevalence represents the proportion of individuals in a population who, at a single point or restricted period of time, possess a health characteristic of interest, as in

2  Statistical Interpretation for Practitioners 1

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10 Before/after measurements

Figure 2.3  Scatterplots showing before and after measurements taken on five individuals. Panel 1 (left) contains a regression line with a negative slope estimated ignoring the pairing within individuals. Panel 2 (right) shows the paired observations, demonstrating that all measurements within individuals increased over time, in contrast to the incorrect impression conveyed in panel 1.

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­ aving a disease (or conversely being disease free). The fach tors that constitute why an individual possesses such a health outcome are intertwined, reflecting contributions from the risk of developing the outcome, the probabilities of recovering, succumbing, or remaining with it over a short or long time (i.e., its duration), and the probability of remaining in the population. Because the contribution of the risk of developing the outcome, also known as incidence, cannot be isolated from the effects of the other respective factors influencing prevalence, prevalence measures cannot serve as surrogates for measures of risk, incidence, or effects. To illustrate this, suppose 100 dogs are born in January 2015 and all become chronically infected with a virus in their first three months of life so that their three‐month risk of infection is 100%. Also suppose that infection only begins to be cleared after 18 months of age and is eliminated at 24 months of age. Then consider another 100 dogs born in January 2016, which all become infected in their first three months as well. Therefore, the three‐month risk of infection in both groups is 100%. Unlike the other dogs, these dogs begin to clear their infection almost immediately (perhaps due to vaccination or postinfection treatment), and by six months are disease free. However, suppose this incidence information was unknown, and a cross‐sectional study of both sets of dogs (n = 200) is performed on June 30, 2016 (i.e., there is no attrition from the combined population). On this date, the prevalence in all 200 dogs will precisely be 50% because the prevalence will be 0% in the 6‐month‐old dogs (n = 100) and 100% in the 18‐month‐old dogs (n = 100). It would be an error, however, to use these prevalence figures to claim that older age is a risk factor for infection because, unknown to the study investigator,

100% of the dogs under three months of age developed the infection; if we assume recovery from infection confers immunity, it is impossible to even address the issue of age above 3 months as a risk factor. This underscores the fallacy of using prevalence to infer risk when factors that affect risk also affect recovery or mortality or leaving a population. Prevalence nevertheless has value in quantifying the relative burden or presence of a health outcome in a population at a particular point in time. Clinicians may, for example, use disease prevalence to guide their pharmaceutical inventories or preventive medicine programs. Measures of relative prevalence can also be informative in hypothesis generation. To understand this, suppose the prevalence of antibodies to a particular organism is found more frequently in dogs with immune‐mediated polyarthritis than in dogs without arthritis. This is no guarantee that the antibody response associated with infection of the organism causally leads to cross‐reactivity with synovial membranes or joint surfaces, because there is no way by measuring only prevalence to distinguish infection occurring prior to onset of polyarthritis from infection occurring afterwards. Such a finding could, however, lead to experimental confirmation (or refutation) of the hypothesis that infection leads to polyarthritis. Serologic surveys are frequently used for this hypothesis‐generating purpose. Cumulative Incidence In contrast, incidence is more appropriate for inferring risk (which can be defined as the average probability of developing a health outcome of interest in a specified time interval). Incidence, by definition, is predicated on the initial absence of the outcome among the members

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of a population at risk of experiencing the outcome. The population is then followed forward in time, and within a specific time period cumulative incidence is defined as the proportion of the individuals initially at risk that newly develop the outcome (assuming no individuals have been lost to follow‐up), and can only strictly fall between 0 and 1. If 300 Yorkshire terriers were followed from birth for five years without any dental interventions, and at the end of this time the presence of tooth resorption was found in 100 dogs, then the five‐year cumulative incidence of tooth resorption would be 0.33 (assuming that there were no dogs that spontaneously recovered from it). Cumulative incidence alone is not a measure of association. However, comparing two or more cumulative incidences under different conditions allows such associations to be quantified. Suppose that in addition to the finding in Yorkshire terriers above, the five‐year cumulative incidence of tooth resorption in border collies was also measured as 0.10. Two important statements can now be made about these two figures. First, that over the five‐year follow‐up period the difference in cumulative incidences between Yorkshire terriers and border collies is 0.20 (0.30 – 0.10). Second, that over the five‐year follow‐up period there is a three‐fold increase (0.30/0.10) in the cumulative incidence in Yorkshire terriers compared to border collies. These two measures of association are known as the cumulative incidence difference and the cumulative incidence ratio, respectively. Other terms that remain in use include risk difference and attributable risk for the former, and risk ratio and relative risk for the latter (these terms should be used with caution, however, because risk implies causation, and statistical measures are distinct from causal measures without corresponding strong causal assumptions that must be invoked). Nevertheless, the cumulative incidence ratio is particularly well suited to effectively communicating relative impacts of factors on health in clinical practice because of its intuitive understandability. Statistical novices can appreciate, for example, a statement such as: “The incidence of parvovirus in unvaccinated puppies under 6 months of age is 10‐fold greater than in vaccinated puppies.” Our understanding of how vaccines, the immune system, and viral infections work could also reasonably allow us to modify this statement by substituting the word “risk” for “incidence.” Such strong biologic knowledge is not always apparent, however, which should motivate caution when using words with causal interpretations like “risk” or “likelihood.” Physicians would have no hesitancy in making the correspondence between the association of tobacco smoke with lung cancer and heart disease with the statement that the risk of lung cancer

and heart disease is greater among smokers than nonsmokers. However, no one would seriously entertain making such a correspondence between the association of carrying matches with lung cancer and heart disease and any causal effect. Therefore, statements about “risk” differences and ratios are best reserved for situations where the justification for alleging a causal relation is supported by knowledge other than purely statistical. Incidence Rates When patients are not all followed for the same amount of time, the cumulative incidence should not be utilized because the reasons for loss to follow‐up (known as right censoring) could be related to the risk of the outcome developing. Instead, another, less intuitive measure of incidence that allows for different follow‐up in each individual is the incidence rate (also sometimes called the hazard rate). In contrast to cumulative incidence, a proportion that has an interpretation always linked to a defined period of time, the incidence rate in a population is a joint function of the number of new (incident) outcome occurrences divided by the sum of the individual times at risk of an outcome among the members of the population, and is expressed as the number of outcomes per unit of time (for example, days, months, years, etc.). Incidence rates are analogous to speeds; across populations, they estimate the average number of outcomes expected to occur per quantity of time. Incidence rates have particular utility in studies measuring the time from onset of follow‐up to when an event occurs. Such events can be recovery, death, remission, or other outcomes. One of the most common applications for incidence rates is in survival studies of comparative treatment efficacy, where particular interest is in the rate of death (or, conversely, survival). Suppose that in Yorkshire terriers the incidence rate of tooth resorption = 0.3 case/dog‐year (or, equivalently, 30 cases/100 dog‐years). This can be interpreted to mean that if 100 dogs were each followed for one year, that 30 cases would be expected; it can also be interpreted to mean that if 50 dogs were each followed for two years, 30 cases would be expected, and so on. The independence of the incidence rate from the number of individuals is perhaps best understood through the analogy of measuring the average speed of automobiles: whether 60 cars each travel one mile in one minute or one car travels 60 miles in 60 minutes, under both scenarios the total distance traveled is 60 miles, the total amount of time traveled is 60 minutes, and the speed is 60 miles per hour. As with cumulative incidence, it is frequent to see incidence rates compared on a ratio scale to measure relative associations; these are called incidence rate ratios or hazard ratios. An incidence rate ratio greater than 1 implies

2  Statistical Interpretation for Practitioners

that the rate of the outcome occurring under the numerator condition (such as treatment A) is faster than the rate under the denominator condition (such as treatment B); conversely, an incidence rate ratio between 0 and 1 indicates that the rate of the outcome occurring under the numerator condition is slower than the rate under the denominator condition. Returning to the above example, if the rate of resorption in Yorkshire terriers is 30 cases/100 dog‐years, and the rate of resorption in border collies is 10 cases/100 dog‐years, then the incidence rate ratio is 3.0. Not only does this indicate that the rate of developing tooth resorption is three times faster in Yorkshire terriers compared to border collies, but it also means that the time to developing tooth resorption is less in Yorkshire terriers, and that the proportion of dogs remaining free of tooth resorption (i.e., “surviving”) is greater in border collies at all times.

I­ ncidence Odds and Incidence Odds Ratios Perhaps the least understood yet ubiquitously found measure of association in the veterinary medical literature is the odds ratio. Although odds ratios arise under different study designs (both experimental and nonexperimental), and their interpretations can vary, these differences are less critical than the properties they share as measures of association and potential impact of factors on the incidence of health outcomes. In order to understand odds ratios, it is necessary to first understand the statistical meaning of an odds. Probabilities are measures of the likelihood of an event occurring, and are only strictly between (and including) 0 and 1; a probability of 0 implies impossibility and a probability of 1 implies inevitability. The odds is another measure of likelihood, and is calculated as the probability of an event occurring divided by the probability of an event not occurring (note that the sum of these two probabilities must equal 1). When the probability of an event occurring equals 0, the odds equals 0. However, when the probability of an event occurring equals 1, then the probability of an event not occurring equals 0, and the odds (which would equal 1 divided by 0) is undefined but infinitely large. The restricted range for probabilities versus the infinitely large range for odds necessarily makes the former more understandable and desirable when communicating biostatistical findings. For example, if the probability of a 14‐year‐old cat developing hyperthyroidism in the ensuing years of its life is 0.60 (which can equivalently be expressed as 60%), then the odds of developing

hyperthyroidism is 1.5 (but there is no percentage equivalent). This leads to two related but important findings. First, as the probability approaches 1, the odds becomes unintelligibly large (for example, a probability of 0.95 corresponds to an odds of 19, while a probability of 0.999 corresponds to an odds of 999). Second, when the probability is small (between 0 and 0.05), such as with the incidence of a rare disease, the odds is similarly small (between 0 and 0.053, respectively). This implies that for rare events, probabilities and odds are nearly equal, and for interpretive purposes are essentially interchangeable. While odds, just like probabilities, can be expressed as unconditional (not depending on the values of other factors) estimates, it is more common to see them expressed as conditional odds; for example, the odds of developing tooth resorption among Yorkshire terriers is a conditional statement because it only applies to Yorkshire terriers and not other dog breeds. Just as ratios of two different cumulative incidences and two different incidence rates can be calculated to measure proportionate changes in incidence between two levels of a variable, so can ratios of odds. Returning to the earlier example of the cumulative incidence of tooth resorption, if the five‐ year cumulative incidence in Yorkshire terriers is 0.3 and the five‐year cumulative incidence in border collies is 0.1, then the five‐year incidence odds in the two breeds are 0.43 and 0.11, respectively. Therefore, the odds ratio relating five‐year incidence of tooth resorption in Yorkshire terriers compared to border collies is 0.43/0.11 = 3.86. This can be contrasted with the cumulative incidence ratio of 3.00, and the disparity between the two measures primarily arises because the odds in (0.43) does not closely approximate the cumulative incidence (0.3) in Yorkshire terriers because the latter is not rare. Just as cumulative incidence, as a probability, is much more comprehensible than an incidence odds, so the cumulative incidence ratio is more comprehensible than an incidence odds ratio. Knowing in the above example that the cumulative incidence ratio = 3.0 makes the interpretation straightforward: the five‐year cumulative incidence in Yorkshire Terriers is three times the incidence in border collies. In the absence of any biases, we could attach a causal interpretation to this and claim that the average five‐year risk of dental resorption in Yorkshire terriers is three times the corresponding risk in border collies. In contrast, no such interpretation can be attributed to the odds ratio of 3.86 above, because it is not in general interpretable either as a relative measure of either risks or incidence rates. While it is legitimate to interpret it literally as a relative odds, other interpretations that include “risk,” “likelihood,” “probability,” and “cumulative incidence” are incorrect and should not be used. This begs

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the question: of what value is the incidence odds ratio if it is interpretatively inferior to other ratio measures? The answer is that in case–control studies, where study groups are selected based on whether or not they possess (cases) or do not yet possess (controls) the outcome of interest, it is not possible to directly calculate measures of incidence because the ratio of cases to controls is determined by the study investigator. Without the ability to calculate individual conditional incidence measures in subgroups, investigators are therefore similarly unable to calculate cumulative incidence or incidence rate ratio measures of association that depend on knowing individual ones. Fortunately, case–control studies are often reserved for studying rare diseases, even conditional on the levels of other variables such as potential risk factors, so using the aforementioned rarity condition that interpretatively equilibrates the cumulative incidence and the incidence odds, the incidence odds ratio can essentially be understood as the cumulative incidence ratio. Subtle modifications in case–control study designs can also sometimes provide more

interpretative latitude in extending such equivalence to incidence rate ratios [8].

­Conclusion This chapter addresses many, but certainly not all, of the most common statistical interpretive issues that face readers of the veterinary medical literature. Appreciating these issues is an important avenue to understanding the basis for evaluating clinical research articles. But statistical analyses only represent one component of medical research; not addressed here are the myriad issues that arise in the course of designing and implementing experimental and nonexperimental studies. Because study design is also inextricably linked to precision and validity issues, statistical inference complements and cannot be divorced from causal inference in evidence‐based veterinary medicine. Topics involving the design of studies to generate data for clinical decision making are the subject of the following chapter.

­References 1 Doll R, Bradford Hill A. The mortality of doctors in

relation to their smoking habits. 1954; 328(7455): 1451–5. 2 Terry L. Smoking and Health: Report of the Advisory Committee to the Surgeon General of the United States. Public Health Service Publication No. 1103. Washington, DC: Department of Health, Education, and Welfare, 1964. 3 Romansik ER, Reilly CM, Kass PH, Moore PF, London CA. Mitotic index is predictive for survival. Vet Pathol 2007; 44(3): 335–41. 4 Elston LB, Sueiro FAR, Cavalcanti JN, Konradin M. Letter to the editor: the importance of the mitotic index as a prognostic factor for survival of canine cutaneous

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mast cell tumors: a validation study. Vet Pathol 2009; 46(2): 362–4. Kass PH, Romansik EM, Reilly CM, Moore PF, London CA. Letter to the Editor: Author Response. Vet Pathol 2009; 46(2): 364–5. Kass PH. Evidence‐based veterinary medicine: evaluation of nonexperimental studies. Pulse 2014; 58(5): 16–17. Buhles W, Kass PH. Understanding and evaluating veterinary clinical research. J Am Anim Hosp Assoc 2012; 48(5): 285–98. Weng HY, Messam LL. Making inferences from a case‐control study: implications and sampling. Vet J 2012; 194(3): 282–7.

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3 Using Data for Clinical Decision Making Philip H. Kass, DVM, MPVM, MS, PhD Department of Population Health and Reproduction, School of Veterinary Medicine, University of California, Davis, CA, USA

The advent of the electronic medical record was accompanied by a new‐found feasibility for veterinary medical practitioners, either at individual hospitals or through multiinstitutional collaborations, to gather, analyze, interpret, and publish their clinical data. While it is still common to read individual case or case series reports in medical journals, electronic data and searchable databases extend the ability of veterinarians to design more sophisticated studies that go beyond the bounds of being purely descriptive. Such design enhancements allow the measurement of differences and associations between two or more factors of interest, and in some cases lead to estimation of measures of effect that under assumptions of unbiasedness can lead to causal inferences. Using clinically derived data to make informed medical decisions is a central tenet underlying the practice of evidence‐based medicine, an evolving form of scientific scholarship that seeks to bring objectivity and elucidation to the implementation of medical investigation, and that exists at the confluence of medicine, basic sciences, applied mathematics/biostatistics, epidemiology, and causal inference. Using data analytically is only one component of evidence‐based medicine, and cannot be dissociated from equally important others. While collaboration is all but essential in ensuring that these respective component specialties are well represented, of most proximate importance to veterinarians seeking to advance beyond the hypothesis phase of their clinical research is a fundamental understanding of hospital‐ based epidemiologic study design, for this is the study type that determines the data collected, and in turn how it should be analyzed and interpreted [1]. The most common study designs employed in clinical veterinary research will be considered in subsequent sections of this chapter. These will be broadly partitioned into two temporal types. The first involves data measurements taken on an assemblage of patients all present at a point in time, or on a succession of patients enrolled over

a typically brief period of time to obtain an adequate total sample size, which are called “cross‐sectional studies.” The second involves those with data on groups of patients with some defining inclusion criteria collected over a specified period of time, allowing measurement of a temporal change in patient characteristics and health status, which are called “longitudinal” studies. The latter can be further subdivided into those that are interventional (the clinical analog of experimental), where the investigators have control over the treatment(s) of primary interest the patients receive, and observational (nonexperimental), where the investigators can passively or actively record, but not manipulate, the treatment(s) of primary interest. The focus here on treatments arises from the nature of clinical research: any treatment in a clinical setting should be pursuant to the best interests of the patient and owner, and should not be any lesser quality of treatment than the accepted standard of care in veterinary medical practice. As noted, analyses of treatments are not restricted to interventional longitudinal studies, although it will be shown that observational longitudinal studies of treatment effects are prone to potentially severe biases. Longitudinal studies also need not be restricted to studying treatment effects: intrinsic characteristics of patients, and how those may be associated with clinical sequelae, may instead be a primary objective of a researcher.

­Hospital‐Based Cross‐sectional Studies Cross‐sectional studies are conducted when a clinician wants to establish an inventory of disease burden, patient characteristics, and exposure to factors in a population. A shelter veterinarian, for example, may be interested in knowing the crude proportion of a shelter population at any defined time point that has clinical signs of upper respiratory disease. This proportion is

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Section 1  Evaluation and Management of the Patient

called the “prevalence” (see Chapter 2 for examples), and is contextually tied to a specific point in time. Knowing such information may be valuable in planning preventive initiatives and having an appropriate inventory of pharmaceuticals available. Hospitals, however, may not have an adequate number of patients at a single time to estimate prevalence, so instead patients fitting inclusion criteria may be captured over a period of time, and the frequency of the factor of interest is then obtained as patients are enrolled. This special type of prevalence is called “period prevalence” and is contextually defined by the beginning and end dates of patient accrual (e.g., the proportion of patients entering a hospital over one month present with clinical signs of upper respiratory disease). It is important to note that no assumptions need be made about when individuals with the outcome actually developed it; it is sufficient to note that at the time of measurement, they either had it or did not (the latter from either never having developed it or from recovering from it). Prevalences measured from cross‐sectional studies are not restricted to being crude (unconditional) measures. Factor‐specific (conditional) prevalences are also estimable, as in sex‐specific or age‐specific prevalences. Moreover, such specific measures can be jointly used for comparative purposes, as in relative prevalences (prevalence ratios). For example, it may be of interest to estimate the proportionate change in the prevalence of feline leukemia virus (FeLV) infections in intact outdoor males compared to sterilized outdoor males. Sampling in cross‐sectional studies can be accomplished in several ways. The most straightforward is to either study an entire population of individuals or take a random sample from one. With either approach, individuals can be cross‐classified by their health outcome status as well as other factors to assess joint associations. Using the previous example, the prevalence of FeLV can be measured in intact and sterilized outdoor male cats, or conversely the prevalence of male sexual status can be measured in cats with and without FeLV. Prevalence ratios provide an objective measure of how relatively common or rare an outcome of interest is between groups, but should not be used for causal inference (such as making statements about likelihood, risk, or probability of a health outcome) because when measured simultaneously, it is often not possible to know if prevalence factors, as putative risk factors, occurred prior to or after the health outcome occurrence. In addition, such factors may not potentially alter only disease incidence but also disease duration and recovery, all of which affect prevalence. Prevalence ratios, as measures of association, therefore have utility for hypothesis generation but lack a foundation for making inferences about causal associations.

An alternative approach to obtaining prevalence ratios when there is a health outcome is to sample subsets of a population defined by certain characteristics. For example, an investigator may choose to deliberately sample a fixed number of sexually intact and sexually altered male outdoor cats, and subsequently ascertain the prevalence of FeLV in these two groups. A prevalence ratio relating the prevalence of FeLV in one sexual status group versus the other is estimable, but not  the converse due to the sampling being based on sexual status. Another sampling variant of cross‐sectional studies is the two‐stage prevalence case–control study. This approach is indicated when a relatively rare disease in a clinical population is studied, and the goal is to estimate not the prevalence of the disease but instead factors that may be associated with it but cannot be readily measured without additional data gathering and consideration of cost/time expenditures. Initially, a cross‐sectional study is undertaken in a clinical population to ascertain disease status. When the cost and effort of obtaining prevalence factor information prohibit procurement of it for every individual in the population, those with the rare disease (prevalent cases) can have it measured, along with only a sample of the remaining nondiseased individuals (nonprevalent controls). Thus, prevalence factor information is efficiently obtained only on a subset of the entire population – either all or some of the diseased individuals and a sample of the nondiseased individuals. The ratio of individuals in the two groups is at the discretion of the investigator, although a 1:1 ratio is statistically the most efficient for a fixed study size. The prevalence case–control study is the cross‐sectional counterpart of the (longitudinal) incidence case–control study discussed later.

­Longitudinal Observational Studies The distinguishing feature of longitudinal medical studies, compared to cross‐sectional studies, is that individuals are monitored over time, typically being followed from a state absent a particular health outcome to a later state that may or may not have experienced a change in outcome status. The two principal types of such studies are cohort studies and case–control studies, which differ in how individuals in them are sampled for study inclusion. Longitudinal studies may also be characterized by their temporality; relative to the study onset, they can be retrospective, with historical data being used during the follow‐up period, or prospective, with future data being collected following study onset. Less commonly, studies can be ambispective, with both retrospective and prospective components.

3  Using Data for Clinical Decision Making

Cohort Studies Assemblages of patients that share one or more defining conditions, typically diseases or syndromes, are often referred to as a “case series” but can be more expansively thought of as a patient cohort. In a hospital setting, such patients present over a period of time; if the condition is common then the period of time for sufficient patient accrual may be brief, but if the condition is rare, it may take many years for an adequate number of patients to be enrolled unless a decision is made to extend patient enrollment to multiple hospitals. While population‐ based (epidemiologic) cohort studies typically study risk factors for disease incidence, the goal of hospital‐based cohort studies is different: to study the outcomes of patients already diagnosed with diseases. Such outcomes are contextual to the specific diseases, but often include the occurrence of or time to remission, relapse, development of another disease condition, recovery, or death. By subdividing cohort members into groups defined by factors thought to influence the respective outcomes, it becomes possible to quantify and compare their effects. There are two main types of outcomes of interest in the study of factors affecting cohorts of patients that share a disease in common. In this example, death will be the illustrative outcome, and sex will be the hypothesized determinant of death. The duality of outcomes is: ●●

●●

whether or not death from the disease occurs following its occurrence in patients how long it takes death from the disease to occur (if it does) following its occurrence in patients.

The first outcome is not as straightforward to evaluate as might initially appear, because it depends on two related factors: how long a patient is diseased, and how long a patient is followed up. To illustrate these points, consider a patient diagnosed with immune‐mediated hemolytic anemia (IMHA) on January 1, 2013, and that succumbed to the disease on January 1, 2019. From the perspective of the investigator, did death occur? The answer is no if the follow‐up time is less than six years (although death occurred after six years, the investigator would not have been aware of it); the answer is yes if the time period evaluated is longer than six years. Therefore, the qualified answer to the question depends on specifying the time of follow‐up after the diagnosis is made. Clearly, the longer a patient is followed up, the greater the probability they will die (although not necessarily from IMHA). By convention, patients who do not experience the health outcome at the point that they are no longer observed at risk of it, because they were lost to follow‐up, withdrew from the study, recovered, or died from a competing cause, are called “censored.”

Suppose that the patient was lost to follow‐up after four years, but was known to be alive up to that time. Did death from IMHA occur? The answer at the four‐year time point is no, because the patient was not observed long enough for death to occur and be recorded. However, such an individual who did not die from IMHA after four years should not statistically be treated as equivalent to a patient still alive after an even longer period of observation. Because most patients are followed for different periods of time after disease diagnosis, an investigator would provide a distorted impression of the natural course of the disease by responding to the simplistic question, “What proportion of patients died?” A legitimate analysis would instead have to be limited to specifying an identical follow‐up period for each patient, such as “What proportion of patients succumbed to IMHA within one year of diagnosis?” Of course, patients followed for less than one year would necessarily be excluded even from this analysis. Because follow‐up times vary so much in patients recruited over long periods of time, a preferable analytic approach takes into account not only if a patient eventually died from the disease, but also how long they were alive prior to death. The apposite question instead becomes: “What is the probability of patients dying from IMHA after accounting for different periods of time under observation following diagnosis?” The answer, a probability that depends on the time interval after diagnosis and characteristics of patients that influence survival, will fall between 0 and 1. This kind of analysis is generically known as a “survival analysis.” One approach commonly used to estimate these probabilities is called the “product‐limit method of survival/failure function estimation” or, equivalently and eponymously, a “Kaplan– Meier analysis.” To illustrate the distinction between counting the proportion of patients who die and the probability of a patient dying as a function of time, consider a hypothetical cohort of 12 patients. Four patients each have either five, six, or seven years of follow‐up, and within each group of four patients, two die and two either do not die or are lost to follow‐up at the end of their follow‐up period. The crude proportion of these 12 patients dying is 50%, but that figure is entirely misleading because of unequal follow‐up times. In contrast, a statistical analysis projects that the probability of dying after five years is 17%, after six years is 38%, and after seven years is 69%. The latter analysis is valid because it accounts for not only whether or not death occurred, but also how long a patient was at risk of dying (by including censored patients). It is also possible to estimate the time it takes to reach a certain probability of death. For example, the time it

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Section 1  Evaluation and Management of the Patient

takes for an estimated 50% probability of patients to die is called the median survival time. In the above example, the median time to death is seven years. With larger datasets, it would also be achievable to estimate the time it would take for a 10%, 25%, 75%, 95%, etc. probability of death. In the less common case that patients are followed for identical periods of time, or inferences are restricted to briefer periods that most or all patients were followed through, it becomes possible to estimate the risk of outcomes for such time periods, either unconditionally or conditional on possible determinants of the outcomes. For example, the one‐year risk of death from patients diagnosed with IMHA conditional on sex can be calculated, and expressed (typically) as a risk ratio (sometimes referred to as a relative risk). It is conventional to report ratio statistics accompanied by: (1) their respective 95% confidence intervals to demonstrate the degree of precision associated with the statistic, and (2) P‐values corresponding to a test of the null hypothesis that the risk ratio = 1 (i.e., there is no difference in the risks of the outcome between the groups compared in an analysis), which are interpretable as the probabilities of finding risk ratios >1 at least as large (or risk ratios 1 at least as large (or rate ratios α) for a particular significance test. The above examples present clinical trial outcomes as means and proportions. However, it is not uncommon in clinical trials to instead use interval‐scaled ordinal variables, such as pain or severity indices, when quantitative measurements are unavailable. These variables generally require nonparametric statistical analyses for group comparisons, as well as for sample size and power calculations, rather than ones that assume that the group outcomes follow a normal (Gaussian) distribution. Nonparametric methods are invaluable for clinical research, particularly when sample sizes are small and underlying distributional assumptions are tenuous (as with interval‐scaled data), and are discussed in more detail in Chapter 2. Cross‐over Trials

In studies of medical interventions with transient effects, there can be a distinct advantage of using each patient as her or his own control. Earlier, when groups composed of randomized individuals were compared, there was always an assumption that they were similar with respect to the distribution of unknown or unmeasurable variables (confounders) that could influence the outcome. This assumption becomes tenuous when group‐specific sample sizes are small or there is so much variability in these factors that the groups would still not be comparable. In this case, sequentially administering different treatments to the same individual allows them to “cross over” between exposure states, permitting within‐individual comparisons and controlling for confounding by intrinsic patient characteristics. This approach requires two important assumptions. One is “temporal stability”: that time itself is not a determinant of the study outcome. It implies that over the study period, the incidence of the outcome remains constant in the absence of any treatments. The second is “causal transience”: that order of treatment administered (e.g., an experimental drug versus placebo) is irrelevant because there are no carry‐over effects of either treatment (i.e., the effects of both are transient), which implies an eventual return to an original state. To make this assumption more tenable, a “wash‐out” period is typically included between treatment options, being sufficiently long to ensure that any effects have become dissipated. Measurements are characteristically taken at multiple times under the different treatments, allowing the assessment of time effects, treatment effects, and the interaction between these two main effects. It is also possible to extend such studies to include evaluating the effects of subgroups of different individuals, such as those defined by age and sex categories.

3  Using Data for Clinical Decision Making

Studies where individuals serve as their own controls lead to collection of dependent data, in which measurements from the same individual are correlated. This arises, for example, when readings are taken from both eyes, when blood parameters are sequentially monitored, or when drug concentrations are evaluated for half‐lives. Conventional statistical procedures assume that data are independent, but when data are paired or repeatedly collected from the same individual, the independence assumption is violated, which leads to invalid hypothesis test results, typically erroneously low P‐values and higher than planned type I error proportions. Analysis of dependent data typically requires specialized statistical methods, from Student’s t‐tests or the nonparametric Wilcoxon signed‐rank tests for paired data, to one‐way repeated measures analysis of variance or the nonparametric Friedman test for sequential observations, to multivariate models, such as mixed effects analysis of variance or linear regression models for more complex designs.

­Conclusion Using data to make causal inferences is a hallmark of evidence‐based medicine, but no inferences will be correct if the evidence used to establish them is flawed. Research can generate new knowledge, but it can also embed flawed conclusions in the literature if sedulous attention is not afforded to proper study design and conduct. Even the most sophisticated statistical methods cannot supplant biases that insidiously enter studies through flaws in study design. Despite the zeal that investigators have for finding statistically significant study results, reported point and variance estimates, confidence intervals, and P‐values will be incorrect unless great care is exercised in designing and establishing the central features of a clinical study: thoughtful selection of comparison groups, unbiased and masked collection of information, diligent follow‐up of patients, proper specification of statistical models, and verification of the model assumptions.

­References 1 Buhles W, Kass PH. Understanding and evaluating

veterinary clinical research. J Am Anim Hosp Assoc 2012; 48(5): 285–98. 2 Lecouteur RA. It’s time. Vet Surg 2007; 36(5): 390–5.

3 Hulley SB, Cummings SR, Browner WS, Grady

DG, Newman T. Designing Clinical Research, 4th edn. Philadelphia, PA: Lippincott Williams & Wilkins, 2013.

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Section 2 Endocrine Disease

29

4 Principles of Endocrinology Robert Kemppainen, DVM, PhD Department of Anatomy, Physiology and Pharmacology, Endocrine Diagnostic Service, Auburn University, Auburn, AL, USA

­Overview of Endocrinology The classic definition of an endocrine tissue is a ductless gland specialized to release a hormone into the blood­ stream to act on a distant target. Glands such as the pituitary, thyroid, islet cells in the pancreas, and adrenal fit this characterization. It is increasingly clear, however, that many (if not all) tissues in the body release sub­ stances into the blood (or at least locally) in a regulated fashion and these substances influence the activity of other cells. Examples include leptin and adiponectin from adipose tissue, natriuretic factors from the heart, and erythropoietin from the kidney. Hormones regulate a vast array of biologic processes, including, but not limited to, fuel mobilization, storage, and distribution; maintenance of electrolyte concentrations in extracellular fluid; and control of metabolic rate. Each ­system employs checks and balances; most utilize some form of negative feedback control to maintain homeostasis. While some systems have as their principal goal mainte­ nance of a steady level of a metabolite or electrolyte, others employ moving set points or thresholds that adapt to chang­ ing conditions. As an example of the latter, stress causes activation of the pituitary‐adrenal system and increased adrenocorticotropic hormone (ACTH) release stimulates release of cortisol from the adrenal cortex. Normally, ­elevated cortisol concentrations suppress ACTH secretion via negative feedback, but in some long‐term stressful ­situations, this negative feedback is overridden so that high ACTH concentrations in circulation are maintained in the presence of the steroid negative feedback signal.

­Chemical Classes of Hormones Classic hormones comprise three main chemical types. Each class has a different means of synthesis, storage, and mechanism of action on targets.

Protein/Polypeptide Hormones Hormones in this class are composed at minimum of three amino acids. Some consist of subunits (e.g., the pituitary hormones luteinizing hormone [LH], follicle‐ stimulating hormone [FSH], and thyroid‐stimulating hormone [TSH]) and may have additional modifications including additions of sugar residues (glycosylation), phosphates, and disulfide bonds. Protein/polypeptides hormones are manufactured by  classic transcription/translation of genes that are expressed in high amounts in the endocrine tissue of ­origin. Encoded in the genes are sequences that direct these molecules to the regulated secretory pathway, resulting in the storage of the hormone in vesicles, ready for release by exocytosis in response to the appropriate signal. The nature of this signal differs depending on the system involved. Most endocrine cells that produce and release hormones of this class store significant amounts of the hormone and have the ability to release a large amount of product if needed. Continual stimulation of release often causes induction of the gene necessary for its synthesis. Protein/polypeptide hormones are released into cir­ culation through the specialized capillaries in their ­tissue of origin and most circulate free or unbound in blood. Exceptions exist; for example, insulin‐like growth factor‐1 (IGF‐1) and growth hormone are bound to varying extent to plasma proteins. In the case of IGF‐1, the binding is significant, accounting for the prolonged half‐life of this hormone. By contrast, most other protein/polypeptide hormones have relatively short half‐lives, usually in the range of several minutes (e.g., the half‐life of ACTH is less than 15 minutes). Protein/polypeptide hormones act on target cells by binding receptors located on the cell surface (see later), rapidly activating specific signal transduction systems in target cells.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

4  Principles of Endocrinology

surface. Hormone binding results in a subtle change in the three‐dimensional structure of the receptor and this signal is carried into the cell where it is transduced into a message(s) that affects cellular biochemistry. Cell surface receptors can be classified by their struc­ ture and the nature of the signaling pathway that they activate. Many protein/polypeptide hormones bind to G‐ protein (guanosine‐protein) coupled receptors. This large family of receptors possess 7‐transmembrane spanning domains and transfer signals to another membrane pro­ tein complex (the G‐protein) consisting of three subunits (alpha, beta, and gamma). The alpha subunit at rest is bound to a GDP, hormone‐receptor binding causes the G‐alpha to replace the guanosine 5’‐diphosphate (GDP) with guanosine 5’‐triphosphate (GTP), resulting in activa­ tion of the G‐protein and dissociation of its subunits. The free subunits then carry the signal to other downstream proteins, including enzymes (e.g., adenylyl cyclase) or ion channels. Hormone activation of a G‐protein does not always result in activation of a cellular process, but may instead result in inhibition. For example, some G‐alpha subunits (Gi), when activated, suppress the activity of adenylyl cyclase, reducing cyclic AMP production in cells. Other types of cell surface receptors are enzymes themselves. For example, the insulin receptor has tyros­ ine kinase activity, meaning that after insulin binds, the intracellular portion of the receptor acts to place phos­ phates on tyrosine residues on the receptor itself and on substrate molecules that contact the activated receptor. Cell surface receptor activation can activate many pathways in target cells, employing signal amplification and cross‐talk. Signaling may also reach the nucleus, therefore affecting gene transcription. Cell surface receptor number can change with condi­ tions. Exposure to high levels of the activating hormone can result in receptor downregulation and reduce the target tissue response. In other situations, target cells express more receptors which can result in upregulation of the response. In many cases, “spare receptors” are pre­ sent on targets, meaning that only a small percentage of receptors need to be occupied in order to obtain a maxi­ mal response in the cell. Steroid and thyroid hormones principally affect their targets by binding to intracellular receptors followed by movement and binding of the hormone receptor com­ plex to specific regions on DNA. This binding then affects (positively or negatively) the rate of gene tran­ scription and ultimately the production of certain cell‐ specific proteins. The receptor molecules act in pairs. Thyroid hormone entry into target cells has been shown to be mediated by specific cell membrane transporters such as monocarboxylate transporter 8 and organic anion transporting polypeptide 1C1. Similar transport­ ers have not been identified for steroid hormones.

Evidence exists for steroid and thyroid hormone effects that do not involve the classic genomic signaling path­ way, and in fact, membrane receptors for both steroids and thyroid hormones have been identified. In addition, these hormones may act inside cells by binding their receptors and then the hormone–receptor complex interacts with intracellular proteins, such as transcrip­ tion factors, without the need to bind DNA. It is believed that much of the antiinflammatory activity of glucocorti­ coids comes about through this process, where the glu­ cocorticoid receptor complex binds to nuclear factor kappa B (NFkappaB), preventing this molecule from stimulating the production of proinflammatory genes. In general, activation of cell surface receptors results in rapid responses in target cells while the effects of steroid and thyroid hormones take longer to become apparent. This difference can be illustrated by compar­ ing the response to ACTH (protein hormone) to that of thyroid hormone. When performing an ACTH response test, a post‐ACTH blood sample is usually collected at one hour for cortisol measurement, the time it takes for the ACTH‐induced cortisol secretion to reach maxi­ mum levels in circulation. On the other hand, noticea­ ble clinical responses in a dog with hypothyroidism treated with T4 replacement therapy require several days (increase in activity) to weeks (stimulation of hair regrowth) to become apparent.

­Regulation of Hormone Secretion Although control of each endocrine system is unique, negative feedback is a dominant feature in most, if not all, systems. The nature of the feedback signal depends on the system; in some cases, it is an ion (calcium in the case of the parathyroid) or fuel molecule (glucose in the case of insulin) while in others it is a hormone itself (cor­ tisol in the case of the ACTH‐adrenal axis). Regulatory control in some systems is multifaceted and more com­ plex; for example, regulation of aldosterone involves the renin‐angiotensin system with sensing mechanisms tied to extracellular fluid volume and sodium levels. In oth­ ers, such as calcium and parathyroid hormone, the rela­ tionship is straightforward with the parathyroid gland responding rapidly to a decline in calcium levels in the blood with an increase in secretion of parathyroid hor­ mone that, in turn, works to restore calcium levels to a set point. Feedback relationships are central to diagnostic testing and to localizing disease. Dexamethasone suppression testing relies on feedback, as this potent glucocorticoid activates negative feedback in the hypothalamus and pituitary, suppressing ACTH secretion and thus secretion (and therefore blood levels) of cortisol. ACTH‐secreting

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4  Principles of Endocrinology

While some diagnostic laboratories still use RIA, most hormone determinations today are made using reporters other than radioactivity. The basic methodology is simi­ lar to RIA (competitive binding to antibodies) but the tag employed is nonradioactive, using methods such as chemiluminescense or an enzyme label (as in enzyme‐ linked immunosorbent assay [ELISA]). Another method that is particularly well suited to the measurement of protein hormones employs two antibodies, directed against different regions of the molecule. The presence of the hormone allows for formation of a “sandwich” complex, which is reported by a tag present on one of the antibodies. Measurement of hormones in companion and domes­ tic animals presents some special challenges. Many assays are commercially available to measure hormones in samples from humans. Some, but not all, of these can

be used to measure the hormone in samples from other animal species. Considerations involved in determining the usefulness of such assay methods include the fact that circulating levels of a particular hormone can vary significantly across species (e.g., T4 levels are approxi­ mately four times higher in humans compared with dogs and cats), protein binding differs across species and other materials in serum or plasma (e.g., lipid content) can impact results obtained in a particular assay. Protein hormones can vary structurally across species and this may have a profound effect on the usefulness of an assay to validate across species. For example, TSH levels ­cannot be determined in samples from dogs or cats using human TSH assay methods. Especially in the case of ­protein hormones, users should contact the reference laboratory to determine if the assay has been validated for use in that species.

­Further Reading Chakravarti B, Chattopadhyay N, Brown EM. Signaling through the extracellular calcium‐sensing receptor (CaSR). Adv Exp Med Biol 2012; 740: 103–42. Dooley R, Harvey BJ, Thomas W. Non‐genomic actions of aldosterone: from receptors and signals to membrane targets. Mol Cell Endocrinol 2012; 350(2): 223–34. Finch NC, Syme HM, Elliott J. Parathyroid hormone concen­ tration in geriatric cats with various degrees of renal function. J Am Vet Med Assoc 2012; 241(10): 1326–35.

Peterson ME. More than just T₄: diagnostic testing for hyperthyroidism in cats. J Feline Med Surg 2013; 15(9): 765–77. Rutter GA, Pullen TJ, Hodson DJ, Martinez‐Sanchez A. Pancreatic β‐cell identity, glucose sensing and the control of insulin secretion. Biochem J 2015; 466(2): 203–18.

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5 Neuroendocrinology Maya Lottati, DVM, PhD, DACVIM (SAIM) TrueCare for Pets, Studio City, CA, USA

Neuroendocrinology refers to the orchestration between the nervous system and the endocrine system to maintain homeostasis by regulating various bodily functions such as stress responsiveness, growth and development, and metabolism. To achieve homeostasis, the body appropriately senses and responds to alterations in the external and internal environment through the modulation of chemical messengers, or hormones. The key players of the neuroendocrine system are the hypothalamus and pituitary gland which are intricately linked via secretory peptides and neuronal connections, as well as target end‐organs (adrenal, thyroid, gonads, liver, and breast tissue) whose role is to exert a physiologic response. These players are integrated with one another through a series of feedback loops that allow controlled secretions of endocrine hormones throughout the body. In this way, the hypothalamus, pituitary gland, and target tissues regulate physiologic function. Alterations to these pathways and regulatory mechanisms may result in hormone disturbances that lead to disease processes.

­ natomic Considerations of the A Hypothalamus and Pituitary System The Hypothalamus The hypothalamus is a highly specialized region of the posterior forebrain that is critical to regulation of vital bodily functions, including temperature regulation, immune function, growth, stress responsiveness, water balance, sexual behavior and reproduction, and metabolism. It is situated below the thalamus just above the midbrain, and arises from the diencephalon during embryologic development. The hypothalamus is composed of neurosecretory neurons that produce various peptide hormones and biogenic amines (dopamine), which enter the circulatory

system and act as endocrine hormones on distant target tissues. The unique capability of the hypothalamus to convert neural signals into hormonal output is called neuroendocrine transduction. Hypothalamic hormones enter the general circulation in one of two ways (Figure 5.1). Some hypothalamic hormones are transported along axons to the median eminence where they enter the portal circulation to reach the anterior pituitary. Here, they stimulate or inhibit anterior pituitary hormone release into the general ­circulation. Other hypothalamic nuclei have axons that terminate at the posterior pituitary. These particular hypothalamic hormones (e.g., vasopressin and oxytocin) are transported to the posterior pituitary where they can be released into the general circulation. The intermediate lobe of the pituitary gland is under direct neural control by the hypothalamus. Hypophysiotropic hormones can be releasing or inhibitory in nature. The hypothalamic and pituitary hormones and their primary effects are listed in Table 5.1. The Pituitary Gland The pituitary gland is referred to as the hypophysis, and it is situated beneath the hypothalamus at the base of the brain. It is securely nestled within a depression of the sphenoid bone called the sella turcica whose shape is described as “saddle‐like.” The pituitary is composed of an anterior lobe (AL; adenohypophysis) and a posterior lobe (PL; neurohypophysis). Their diverse function is a reflection of their distinct embryologic origin. Adenohypophysis and Intermediate Lobe

The adenohypophysis is the glandular portion of the pituitary gland. During embryologic development, it arises from an ectodermal pouch (Rathke’s pouch) growing upwards from the oral cavity. In many mammalian species, there is a distinct intermediate lobe (IL) that

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5 Neuroendocrinology

Table 5.1  Hypothalamic‐pituitary hormones and their primary functions Endocrine gland

Hormone

Primary function

Target

Hypothalamus

Thyrotropin‐releasing hormone (TRH)

Stimulates the release of thyroid‐stimulating hormone (TSH) and prolactin (PRL)

Anterior pituitary

Gonadotropin‐releasing hormone (GnRH)

Stimulates the release of luteinizing hormone (LH) and follicle‐stimulating hormone (FSH)

Corticotropin‐releasing hormone (CRH)

Stimulates the release adrenocorticotropic hormone (ACTH) and proopiomelanocortin (POMC)‐derived peptides

Growth hormone‐ releasing hormone (GHRH)

Stimulates the release of growth hormone (GH)

Prolactin‐releasing hormone (PRLH)

Stimulates the release of prolactin (PRL)

Somatostatin (SS)

Inhibits the release of GH and thyroid‐stimulating hormone (TSH)

Dopamine (AKA prolactin‐inhibiting hormone)

Inhibits the release of PRL

Vasopressin

Regulates blood volume

Distal convoluted tubules and collecting ducts of kidney

Stimulates smooth muscle contraction

Blood vessels

Posterior pituitarya

Oxytocin

Anterior pituitary

Stimulates uterine contraction during parturition

Uterine smooth muscle

Stimulates milk ejection during lactation

Myoepithelial cells of mammary gland ducts

ACTH

Stimulates the release of glucocorticoids and sex steroids

Adrenal cortex

GH

Promotes growth and development

Various tissuesb

Prolactin

Stimulates milk production

Mammary tissue

TSH

Stimulates the release of thyroid hormone

Thyroid gland

FSH

Stimulates estrogen production and promotes ovarian follicle development in the female. Stimulates spermatogenesis in the male

Gonads

LH

Stimulates progesterone production and promotes ovarian follicle maturation in the female. Stimulates testosterone production in the male.

a

 These hormones are synthesized in the hypothalamus and released from the posterior pituitary gland.  See GH section.

b

­ aintenance of the corpus luteum. Together, the gonm adotropins stimulate production of the  sex hormones estradiol and progesterone. In male gonads, LH stimulates testosterone production by Leydig cells, and FSH stimulates Sertoli cell proliferation and maintains sperm quality. Gonadotropin secretion is pulsatile, and release is stimulated by gonadotropin‐releasing hormone (GnRH) binding to the anterior pituitary. LH and FSH circulate to target tissues and stimulate sex hormone production. As gonadotropin levels rise, LH and FSH exert negative feedback on the hypothalamus and pituitary to decrease their own production. Sex hormone products also feed

back on the hypothalamus and pituitary gland to variably regulate gonadotropin production. Testosterone and estrogen inhibit LH and FSH release by the anterior pituitary, while estradiol may have variable effects on gonadotropin production depending on its plasma concentration, a feature that may be important during various stages of reproduction. High levels of estradiol exert positive feedback on GnRH to promote sustained ­gonadotropin release, while low levels negatively feedback to inhibit hypothalamic GnRH and pituitary ­gonadotropin release. Inhibin, a protein produced by ovarian granulosa cells in females and by Sertoli cells in males, inhibits FSH release both by direct feedback on

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Section 2  Endocrine Disease

its ­production by the pituitary and indirectly by altering GnRH receptor sensitivity. Activin, a protein produced by the ovaries, and dopamine are known stimulators of FSH release. Endogenous opioids have an inhibitory influence on gonadotropin production through alteration of GnRH secretion. Somatomammotropic Hormones

Growth hormone and prolactin (PRL) are grouped together as the somatomammotropic hormones. Their amino acid sequences are similar, which may account for some common functions. Growth Hormone  As the name suggests, GH regulates

growth and development. In young animals, it stimulates growth of connective tissue, long bones, muscles, and endocrine glands. GH is a metabolic counterregulatory hormone, which opposes the actions of insulin and raises glucose by stimulating hepatic gluconeogenesis and impairing glucose uptake in muscle. GH stimulates lipolysis in adipose tissue, which results in increased circulating free fatty acids, and has an anabolic effect on protein synthesis. GH stimulates production of insulin‐like growth factor 1 (IGF‐1) by hepatocytes. IGF‐1 is a primary mediator of the effects of GH. Growth hormone is a single‐chain polypeptide containing two disulfide bridges. The sequence is highly conserved among species; the sequences of canine and feline GH differ by only one amino acid, while those of canine and porcine GH are identical. Receptors for GH have been identified throughout the body, but are found in the highest concentrations in skeletal muscle, liver, adipose tissue, heart, kidneys, lungs, and cartilage. This diffuse distribution in part explains GH’s diverse effects. GH receptors are known to have altered sensitivity to various secretagogues throughout life, which aids in preferential growth and development in young, and specialized metabolic function in adults. Growth hormone secretion is pulsatile, and results from the complex interactions between growth hormone‐releasing hormone (GHRH), SS, and peripheral factors (both neural and hormonal). GH release is primarily a reflection of the delivery and binding of hypothalamic GHRH to pituitary somatotropic cells. SS is a known inhibitor of GH secretion, thus high GHRH and low SS levels result in GH release. SS is a more potent inhibitor of GH release than of TSH release. Insulin‐like growth factor 1 is secreted in response to GH, but is also responsive to systemic alterations in energy balance and nutrition. Low levels of IGF‐1 stimulate GH release. As GH and IGF‐1 levels rise, they feed back on the hypothalamus to decrease GHRH and increase SS secretion, which results in reduced production of GH. Ghrelin, an endogenous ligand produced by

enteral cells of the stomach, is a potent GH secretagogue. Other factors known to directly or indirectly influence GH secretion include nutritional states, stress, sex steroids, and exercise. For example, GH secretion is  stimulated by alpha‐adrenergic agonists and inhibited  by beta‐adrenergic agonists and hyperglycemia. Estradiol modulates GH levels indirectly via inhibitory effects on SS. Prolactin  Prolactin is involved in the regulation of lacta-

tion and reproduction. PRL works together with sex steroids to stimulate mammary gland development and regulate milk production during lactation. Although PRL is secreted in the male, its physiologic role is not yet fully elucidated. Prolactin is a single‐chain polypeptide containing three disulfide bridges. There is significant sequence variation between species. PRL secretion is pulsatile, and stimulated by the hypothalamic hormones prolactin‐ releasing hormone (PRLH) and TRH. PRL is also released in response to the neural stimulation of suckling during lactation. The definitive identity of PRLH is not yet known, but vasoactive intestinal peptide (VIP) has been suggested. PRL release is inhibited by dopamine, which is also referred to as prolactin‐inhibiting hormone. In the regulation of PRL release, the inhibitory component is known to predominate over the stimulatory component. Upon binding to its receptors in lactotropic cells, dopamine inhibits PRL release. PRL, in turn, acts in a short feedback loop by circulating and binding to PRL receptors on dopaminergic neurons in the hypothalamus. PRL release is also stimulated by oxytocin and angiotensin II, and inhibited by gamma‐aminobutyric acid (GABA). Unlike other hypophyseal hormones, PRL is not subject to typical feedback inhibition by target tissue hormone products. Instead, its release is solely regulated through hypothalamic input and the suckling reflex.

Proopiomelanocortin‐Derived Peptides

Proopiomelanocortin is a large prohormone which encodes numerous secretory products (Figure 5.2). It is synthesized by corticotropic cells of the anterior pituitary and melanotropic cells of the IL. Proteolytic enzymes hydrolyze POMC into ACTH and beta‐lipotropin (beta‐ LPH). These products are then further cleaved into alpha‐MSH, which stimulates melanin production by epithelial cells and leads to darkening of skin, beta‐ endorphin (an endogenous opioid), and other related peptides. The function of many of these peptides is not yet fully understood and remains under investigation. Proopiomelanocortin synthesis is under direct neural control by the hypothalamus primarily through inhibitory dopaminergic innervation. Thus, dopamine exerts tonic inhibition of POMC release under normal

5 Neuroendocrinology

Signal peptide

γ1,2MSH γ3MSH

JP

MSH

β-MSH

CLIP

γ-END

ACTH

pro-γMSH N-POC

Enk

β-END

γ-LPH β-LPH

Figure 5.2  Schematic representation of proopiomelanocortin (POMC) and production of ACTH and related peptides. POMC contains four major domains: the signal peptide domain; the N‐terminal peptide domain (N‐POC) which contains the (pro) gamma‐MSH peptide and joining peptide (JP); the ACTH domain which contains melanocyte‐stimulating hormone (MSH) and corticotropin‐like intermediate lobe peptide (CLIP); and the beta‐lipotropin (beta‐LPH) domain which generates metenkephalin (Enk) and the endorphin (END) family of peptides. Vertical lines represent potential sites of proteolytic cleavage. Source: Mol and Meij 2008. Reproduced with permission of Elsevier.

circumstances. Corticotropin‐releasing hormone (CRH) will stimulate POMC production, which results in ACTH release. Adrenocorticotropic Hormone  Adrenocorticotropic hormone is a glycoprotein hormone that regulates the synthesis and release of cortisol. Its binding to adrenal cortical cells stimulates production of glucocorticoids, sex steroids, and, to a lesser extent, mineralocorticoids. ACTH also regulates growth of the adrenal cortex as evidenced by high ACTH causing hypertrophy of the adrenal cortex, and absent ACTH causing atrophy. Adrenocorticotropic hormone secretion is pulsatile, and stimulated by the hypothalamic hormones CRH and vasopressin. These hormones act synergistically to stimulate ACTH release, but stress will variably modulate their production by the hypothalamus. Adrenocorticotropic hormone stimulates cortisol release, and as cortisol levels rise, they feed back on the hypothalamus and pituitary to inhibit production of CRH and ACTH, respectively. Hypothalamic dopamine exerts tonic inhibition of ACTH release through dopaminergic neural connections terminating on the IL. Some factors that directly or indirectly influence ACTH release include leptin, vasoactive intestinal peptide (VIP), neuropeptide Y, cholecystokinin (CCK), cytokines such as interleukin‐1 and tumor necrosis factor‐alpha, serotonin agonists, and beta‐adrenergic agonists. Additionally, ACTH release is triggered by exercise and hypoglycemia. Neurohypophysis

The neural portion of the pituitary is called the neurohypophysis, and it is composed of axons that extend downward as a large bundle from the hypothalamus. During embryologic development, the neurohypophysis and hypothalamus both arise from the diencephalon and remain connected through life.

The hormones oxytocin and vasopressin are synthesized by the paraventricular and supraoptic nuclei of the hypothalamus. They are transported to the neurohypophysis through direct neuronal projections, and are stored in secretory vesicles in axon terminals. Upon appropriate neurogenic stimulation, they are released into the general circulation where they travel to distant target tissues and exert a physiologic effect. Vasopressin; Antidiuretic Hormone, Arginine Vasopressin

Vasopressin (VP) regulates blood volume through alteration of water excretion in the kidneys. It also promotes vascular smooth muscle contraction, and stimulates ACTH release. These diverse effects are mediated by vasopressin receptors: V1 in vascular smooth muscle, V2 in the kidney epithelium, and V3 in the anterior pituitary. Vasopressin secretion is pulsatile, and hypothalamic production is primarily stimulated when high plasma osmolality (hemoconcentration) or a significant decrease in blood volume (hypovolemia) is detected. In the absence of circulating VP, the epithelial cells lining the distal convoluted tubules and collecting ducts in the kidney are largely impermeable to water. As such, large volumes of dilute filtrate are excreted into the urine. Upon release, circulating VP binds to VP2 receptors and enhances the permeability of water‐selective channel proteins (aquaporins) to water in renal epithelial cells. This results in the passive diffusion of water into the interstitial space due to the medullary osmotic gradient, thereby restoring blood volume and suppressing VP production by the hypothalamus. The binding of VP to V1 receptors in blood vessels results in vasoconstriction through activation of calcium channels (vasopressor effects). VP binding to V3 receptors in the anterior pituitary stimulates ACTH release. The production of VP is stimulated by CRH and angiotensin II, and inhibited by glucocorticoids.

39

5 Neuroendocrinology

­Vascular Supply The vascular organization of the neuroendocrine system is responsible for the effectiveness of feedback regulation and the hormone cascade. The hypothalamus receives arterial blood from the circle of Willis, while the pituitary gland is supplied by the superior and inferior hypophyseal arteries (branches of the internal carotid artery). The hypophyseal portal circulation connects the hypothalamus with the anterior pituitary, thereby allowing direct transport of hypophysiotropic hormones to their corresponding hypophyseal receptors. Venous blood from the pituitary drains into the cavernous sinus where it enters the systemic circulation to reach target glands and tissues. Short portal veins connect the ­posterior and anterior lobes of the pituitary. A small portion of pituitary outflow is thought to ascend in retrograde to  the hypothalamus where it can directly ­participate in regulation of hypophyseal hormone production. It is important to note that the blood–brain barrier is incomplete surrounding the portal system, which ensures exposure to circulating hormones emanating from peripheral tissues.

­Regulation of Hormone Secretion Hypothalamus The hypothalamus is responsible for integrating sensory stimuli from the external environment with input from the internal environment to maintain homeostasis. Therefore, neuroendocrine transduction is regulated not only by hormonal feedback but also by neural input from higher brain centers that convey information about the

external environment such as temperature, light, pain, and other sensory stimuli. In the hypothalamus, cell bodies responsible for hormone synthesis are anatomically intermingled, and there is often redundancy in production of a single hormone by separate hypothalamic nuclei. A single hypothalamic hormone may exert an effect on multiple pituitary hormones, and multiple hypothalamic hormones may influence regulation of a single hypophyseal hormone. This overlap and redundancy throughout the neuroendocrine system allows for a high degree of regulation. Pituitary Gland Pituitary hormone regulation is not only under hormonal feedback but is also influenced by central and peripheral neural input, an example of which is illustrated by lactation and nursing. Suckling activates sensory pathways leading to the hypothalamus that trigger release of PRLH and TRH, and inhibit release of dopamine (prolactin‐ inhibiting hormone), and result in PL secretion by the anterior pituitary. Concurrently, suckling stimulates release of OT by the posterior pituitary, and together PRL and OT stimulate milk production and initiate milk ejection in mammary glands, respectively. Central factors also influence pituitary hormone release, as is seen when on occasion lactating mothers expel a small amount of milk upon the thought of their baby. The images stimulate the cerebral cortex, which results in release of OT and contraction of myoepithelium in mammary gland ducts. The majority of pituitary hormones are released in a pulsatile manner, therefore regulation of target organs and tissues relies not only on absolute hormone concentrations but also on the frequency and amplitude of hormone pulses.

­Further Reading Goldman L, Schafer AI. Goldman‐Cecil Medicine, 25th edn. St Louis, MO: Saunders Elsevier, 2012. Kaneko JJ, Harvey JW, Bruss ML. Clinical Biochemistry of Domestic Animals, 6th edn. Cambridge, MA: Academic Press, 2008.

Malven PV. Mammalian Neuroendocrinology. Boca Raton, FL: CRC Press, 1993. Rijnberk A, Kooistra HS. Clinical Endocrinology of Dogs and Cats: An Illustrated Text, 2nd edn. Hanover, Germany: Schlütersche, 2010.

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6 Feline Acromegaly David S. Bruyette, DVM, DACVIM (SAIM) Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology and Epidemiology Feline acromegaly is a disease characterized by excessive growth hormone secretion leading to a wide array of clinical signs caused by the hormones’ effects on multi­ ple organ systems. These effects can be divided into two major classes. The first are the catabolic actions of growth hormone that include insulin antagonism, lipoly­ sis, and gluconeogenesis with the net effect of promoting hyperglycemia. The second are the slow anabolic (or hypertrophic) effects of growth hormone, which are mediated by insulin‐like growth factors. Growth hor­ mone stimulates production of insulin‐like growth fac­ tors in several different tissues. Insulin‐like growth factor‐1 (IGF‐1), which is produced in the liver, is thought to be the key factor that facilitates the anabolic effects of growth hormone that are responsible for the characte­ ristic appearance of acromegalic people, dogs, and cats. Growth hormone is produced in the pars distalis (ante­ rior pituitary), specifically by acidophilic cells, called somatotrophs. The release of growth hormone is regu­ lated by many factors, the most important of which is growth hormone‐releasing hormone (GHRH) produced by the hypothalamus. Recently, another hormone, ghrelin, has also been identified as a potent stimulator of growth hormone release. Ghrelin is produced by the stomach and released following ingestion of a meal. Release of growth hormone is inhibited by the hypo­ thalamic hormone somatostatin as well as by growth hormone and IGF‐1 via negative feedback. Feline acro­ megaly is typically the result of a functional adenoma of the pituitary that releases growth hormone despite nega­ tive feedback resulting in excessive growth hormone production and release. Feline acromegaly, thought to be an uncommon ­disease until recently, is likely underdiagnosed. Three studies have

examined the incidence of acromegaly in the ­diabetic cat population. A recent study in the United Kingdom meas­ ured IGF‐1 levels in variably controlled diabetic cats. Of the 184 cases, 59 (32%) had markedly increased IGF‐1 concentrations. Eighteen of these 59 cats underwent pitu­ itary imaging, confirming a diagnosis of acromegaly in 17/18 (94%). A second study examined 225 cats with vari­ ably controlled diabetes and 40 (17.8 %) had markedly high IGF‐1 concentrations. In the largest study to date, of 1222 cats with diabetes, 323 (26.4 %) had IGF‐1 suggesting acromegaly and 90% had a pituitary mass upon imaging (CT or MRI). This suggests that 18–32% of diabetic cats may have concurrent acromegaly and acromegaly may be one of the most important predisposing factors to feline diabetes through the induction of insulin resistance. Etiology Acromegaly in humans is usually sporadic, but up to 20% of familial isolated pituitary adenomas are caused by g­ ermline sequence variants of the aryl‐hydrocarbon‐receptor inter­ acting protein (AIP) gene. The AIP gene is associated with xenobiotic metabolizing enzymes and endocrine disrupt­ ing chemicals in the environment, such as bisphenol A and PBDEs, have been linked to both feline acromegaly and hyperthyroidism. Feline acromegaly has similarities to human acromegalic families with AIP mutations. A recent study sequenced the feline AIP gene, to identify sequence variants and compare the AIP gene  sequence between feline acromegalic and control cats, and in acromegalic sib­ lings. A single nonsynonymous single‐nucleotide polymor­ phism (SNP) was identified in exon 1 (AIP:c.9T > G) of two acromegalic cats and none of the control cats, as well as both members of one sibling pair. The region of this SNP is considered essential for the interaction of the AIP protein with its receptor. This sequence variant has not previously been reported in humans. Two additional synonymous

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 2  Endocrine Disease

sequence variants were identified (AIP:c.481C > T and AIP:c.826C > T). This was the first molecular study to investigate a potential genetic cause of feline acromegaly and identified a nonsynonymous AIP SNP in 20% of the acromegalic cat population evaluated, as well as in one of the sibling pairs evaluated.

­History and Clinical Signs Feline acromegaly most commonly affects middle‐aged to older, male castrated cats. In one study, 13 of 14 cats with acromegaly were males with an average age of 10.2 years. This association may be biased, however, as most cats that are diagnosed with acromegaly present for insulin‐ resistant diabetes mellitus, which is also more common in older, male castrated cats. Based on available data, there is no known breed association for acromegaly. Most patients with acromegaly present for insulin‐ resistant diabetes mellitus (insulin doses greater than 1.5–2.2 units/kg BID) with concurrent weight gain rather than weight loss. Growth hormone has effects on all the tissues in the body and therefore the disease has a range of clinical signs. Physical characteristics of acromegaly include increased body weight, a broad­ ened face, enlarged feet, protrusion of the mandible (prognathia inferior), increased interdental spacing, stertorous breathing, organomegaly, and a poor hair­ coat. Cardiovascular signs include the presence of a heart murmur, hypertension, arrhythmia, associated with hypertrophic cardiomyopathy. Neurologic dis­ ease associated with feline acromegaly is uncommon but can occur with a pituitary macroadenoma. Neurologic signs that have been observed with acro­ megaly include dullness, lethargy, abnormal behavior, circling, and blindness. Glomerulopathy and second­ ary renal failure have also been associated with feline acromegaly. Histopathologic evaluation of the kidneys from acromegalic cats has revealed thickening of the glomerular basement membrane and Bowman’s cap­ sule, periglomerular fibrosis, and degeneration of the renal tubules. Arthropathy and peripheral (diabetic) neuropathy have been shown to cause lameness in acromegalic cats.

­Diagnosis Diagnosis of feline acromegaly starts with clinical suspi­ cion, using a thorough history, signalment, and clinical signs. Many of the abnormalities in the minimum data­ base of affected cats reflect concurrent diabetes mellitus and include erythrocytosis, hyperglycemia, increased

liver enzymes (ALT, ALP), hypercholesterolemia, hyper­ phosphatemia, hyperglobulinemia, azotemia, glucosuria, ketonuria, proteinuria, and isosthenuria. Growth hormone concentration is a common diagnos­ tic for acromegaly in humans, but assays specifically for feline growth hormone are not widely available. An assay using ovine GH as the antigen has been validated for use in cats, but is only available in Europe. However, even if an assay were available, growth hormone concentrations alone may not be a reliable diagnostic for acromegaly. Growth hormone production is cyclic and levels may vary throughout the day. A single high value may not necessarily be diagnostic for acromegaly. Additionally, it has been shown that growth hormone may be elevated in nonacromegalic diabetic cats. This may be due to the fact that portal insulin is required for the liver to produce IGF‐1. In diabetics being treated with insulin subcutane­ ously, portal insulin concentrations will remain low, resulting in decreased IGF‐1 production and theoreti­ cally decreased inhibition of GH release. In addition, GH levels may also not be elevated early in the course of the disease, but later typically increase significantly. Insulin‐like growth factor‐1 is the endocrine assay most commonly used to diagnose feline acromegaly and is widely available through the Michigan State University Diagnostic Center for Population and Animal Health (www.animalhealth.msu.edu/Forms/F.ADM.7.pdf ). Unlike GH, IGF‐1 concentrations are less likely to fluctu­ ate over the course of the day as the majority of IGF‐1 is protein bound, giving it a longer half‐life in the body. In addition, IGF‐1 increases in response to chronically ele­ vated GH concentrations and is thought to be a reflection of GH levels over the previous 24 hours. However, just as with GH, elevations in IGF‐1 concentration alone may not be diagnostic for acromegaly. One study found that IGF‐1 levels in nonacromegalic cats on long‐term insulin treat­ ment (>14 months) had higher levels of IGF‐1 than non­ diabetics. It was proposed that insulin treatment allowed for beta cell regeneration and increased portal insulin, leading to elevations in IGF‐1. A subsequent study evalu­ ating IGF‐1 levels in confirmed acromegalics, diabetics, diabetics, and healthy cats found that acromegalic diabet­ ics had significantly higher levels of IGF‐1 than diabetics and nondiabetics. This study concluded that IGF‐1 was 84% sensitive and 92% specific for diagnosing feline acro­ megaly. No correlation between long‐term insulin use and elevations in IGF‐1 concentrations was found in this study. Diagnostic Imaging Radiographic findings associated with feline acromegaly are related to the hypertrophic effects of excessive GH. Hyperostosis of the calvarium, spondylosis of the spine, and protrusion of the mandible are common findings.

6  Feline Acromegaly

Periosteal reaction, osteophyte production, soft tissue swelling, and collapse of joint spaces are signs associated with the degenerative arthropathy linked to feline acro­ megaly. Thoracic radiographs may reveal cardiomegaly (hypertrophic cardiomyopathy) and/or congestive heart failure. Nonspecific signs such as abdominal organo­ megaly (hepatic, renal, and adrenal) may be revealed by abdominal ultrasound. Advanced imaging is needed to document the presence of a pituitary macroadenoma. Computed tomography (CT) and magnetic resonance imaging (MRI) are both useful for identifying pituitary masses. However, one study found MRI to be the more sensitive imaging modality. The presence of a pituitary tumor alone is not diagnostic for feline acromegaly as other functional tumors of the pitui­ tary may also result in insulin‐resistant diabetes, such as adrenocorticotropic hormone (ACTH)‐producing tumors in patients with Cushing disease. Conversely, the absence of a pituitary mass does not rule out acromegaly as there have been reported cases where a patient had a negative MRI but a pituitary mass was identified at necropsy and histopathology confirmed a GH‐secreting adenoma. Histopathology Histopathology is needed for definitive diagnosis which makes antemortem diagnosis challenging. However, with advancements in surgical procedures such as transsphe­ noidal hypophysectomy, surgical excisional biopsy is pos­ sible. The main histopathologic change associated with acromegaly is acidophil proliferation in pituitary tumors. Adrenocortical Function Testing There is no single test for the diagnosis of feline acro­ megaly. Clinical suspicion based on a thorough history and physical exam is essential. As stated earlier, the most common presenting complaint for patients with acro­ megaly is insulin resistance with weight gain. The two most common causes of insulin resistance in cats are hyperadrenocorticism and acromegaly. Both of these diseases can be associated with a pituitary mass and bilateral adrenomegaly. As such, all suspected acromeg­ alics should undergo adrenocortical testing via the ACTH stimulation test and/or low‐dose dexamethasone suppression test. Normal results on these tests would then be an indication to screen for acromegaly.

­Treatment Medical Treatment Somatostatin is a hypothalamic hormone that acts on the pituitary to inhibit GH release. Somatostatin analogs

are commonly used in human medicine for the treat­ ment of acromegaly and have efficacy rates of 50–60%. The somatostatin analog octreotide has been evaluated in a small number of feline acromegalics with limited success. One study in four cats found no change in GH following treatment. Another study measured the short‐ term effects of octreotide in five feline acromegalics and found a decrease in growth hormone concentrations for up to 90 minutes. A small study evaluating an octreotide analog (Sandostatin® LAR) showed no benefit in cats treated for 3–6 months. However, a recent study examined the long‐term ­medical management of acromegaly in cats using pasire­ otide, a novel somatostatin analog, which had been shown to decrease serum IGF‐1 and improve insulin sensitivity in cats with acromegaly when administered as a short‐acting preparation. The study used once‐monthly administration of long‐acting pasireotide (pasireotide LAR) for treatment of cats with acromegaly. Fourteen cats with acromegaly, diagnosed based on the presence of diabetes mellitus, pituitary enlargement, and serum IGF‐1 >1000 ng/mL, were enrolled. Cats received pasire­ otide LAR (6–8 mg/kg SC) once monthly for six months. Fructosamine and IGF‐1 concentrations, and 12‐hour blood glucose curves (BGCs) were assessed at baseline and then monthly. Product of fructosamine concentra­ tion and insulin dose was calculated as an indicator of insulin resistance (Insulin Resistance Index). Eight cats completed the trial. Three cats entered diabetic remission. Median IGF‐1 (baseline 1962  ng/mL [range 1051– 2000 ng/mL]; month 6 1253 ng/mL [524–1987 ng/mL]; P < 0.001) and median Insulin Resistance Index (baseline 812 μmolU/L kg [173–3565  μmolU/L kg]; month 6 135 μmolU/L kg [0–443  μmolU/L kg]; P = 0.001) decreased significantly. No significant change was found in mean fructosamine (baseline 494 ± 127 μmol/L; month 6 319 ± 113.3 μmol/L; P = 0.07) or mean blood glucose (baseline 347.7 ± 111.0 mg/dL; month 6 319.5 ± 113.3 mg/ dL; P = 0.11), despite a significant decrease in median insulin dose (baseline 1.5 [0.4–5.2] U/kg; month 6 0.3 [0.0–1.4] U/kg; P < 0.001). Adverse events included diar­ rhea (n = 11), hypoglycemia (n = 5), and worsening poly­ phagia (n = 2). This initial study indicates that pasireotide LAR has potential as a long‐term management option for cats with acromegaly. Conflicting results with various somatostatin analogs may be related to difference in somatostatin receptor subtypes. Future studies are required to identify the somatostatin receptor subtypes in GH‐secreting feline pituitary tumors to determine if they are similar to the ones found in humans. Dopamine agonists and more recently GH receptor antagonists are also used in human medicine for the treat­ ment of acromegaly. The use of GH receptor antagonists

45

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Section 2  Endocrine Disease

has not been reported in cats but in humans, response rates have been reported to be as high as 90%. However, it has been noted that the medication has no effect on tumor size and thus would be of no benefit in patients with neurologic signs. A single case study using a dopa­ mine agonist (L‐deprenyl) for the treatment of feline acro­ megaly showed no effect on reducing insulin requirements or clinical signs of disease. In humans, dopamine agonists are typically only 10–20% effective, but are often used in combination with other medications. Increasing the dosage of insulin to improve glycemic control and clinical signs of diabetes is the most con­ servative choice for treating insulin‐resistant diabetic acromegalics. However, there have been reports that some patients suddenly and inexplicably become sensi­ tized to insulin, resulting in hypoglycemic crises. In one study, several acromegalic cats were euthanized after experiencing hypoglycemic coma. Surgical Treatment Surgical removal of the pituitary tumor (adenectomy) is the treatment of choice for acromegaly in human medicine. The procedure can be performed in cats and dogs, usually employing complete removal of the entire pituitary (hypophysectomy). In veterinary medicine, a transsphenoidal approach is used involving only a small incision through the soft p ­ alate and then approaching the pituitary gland through the basisphenoid bone. Complications associated with the surgery include hemorrhage and incision dehiscence. Post surgery, patients are treated with cortisone, L‐thyroxine, and desmopressin. The same surgical procedure is also used to treat pituitary‐dependent hyperadrenocorticism in both dogs and cats. A study in which seven cats with pituitary‐dependent hyperadrenocorticism were treated with transsphenoidal hypophysectomy resulted in five cats showing complete resolution of the disease. Four of these cats had concurrent diabetes mellitus, two of which showed increased insulin responsiveness after surgery. A few case reports exist for the treatment of feline acro­ megaly with transsphenoidal hypophysectomy. Prior to surgery, one patient was an insulin‐resistant diabetic that was still exhibiting clinical signs despite receiving 25 U of insulin (Levemir® Novo Nordisk) four times per day. Three weeks after surgery, the patient no longer required insulin therapy and up to one year later the patient’s IGF‐1 and GH concentrations were within normal limits. In another single case report, a 13‐year‐old male cas­ trated domestic shorthair was treated for acromegaly with transsphenoidal hypophysectomy. The patient had a his­ tory of insulin‐resistant diabetes mellitus (15 units of glar­ gine BID) and was diagnosed with acromegaly via elevated IGF‐1 (447 nmol/L) and visualization of a pituitary mass

on MRI. The diabetes resolved two weeks postoperatively and remained in remission for eight months at which time the patient was euthanized as a result of feline infectious peritonitis. In more recent studies, hypophysectomy has been offered to owners who presented diabetic cats with con­ firmed acromegaly (IGF‐1 >1000 ng/mL, pituitary mass) to the Royal Veterinary College since 2012. All cats were operated on by one neurosurgeon. Hypophysectomy was performed by manual extirpation using fine surgical tools via a transoral transsphenoidal approach. Cats received intensive peri‐ and postoperative monitoring of electro­ lytes, glucose and blood pressure, and were initially administered conjunctival desmopressin (DDAVP), intra­ venous infusions of insulin and hydrocortisone, before being transitioned to conjunctival DDAVP, oral hydrocor­ tisone and levothyroxine and subcutaneous glargine insu­ lin. In total, 21 diabetic cats underwent hypophysectomy from April 2012 to October 2014 (median, range; age: 10.3 years, 5.4–14.8; pituitary height: 6.0 mm, 4.0–10.6; IGF‐1: 1833 ng/mL, 1138 to >2000; fructosamine: 574 μmol/L, 339–1076). Other than mild pelvic limb weakness, no cat displayed overt neurologic deficits prior to surgery. Three (14%) cats died postoperatively. Two cats did not recover from anesthesia and were euthanized within 24 hours; one cat developed septic meningitis and was euthanized 17 days postoperatively. All surviving cats (n = 18) had a reduction of serum IGF‐1 and 16 cats (89%) showed IGF‐1 normalization (median postoperative serum IGF‐1: 38 ng/ mL (15–1955), P < 0.001). Fourteen of the 18 s­urviving cats (78%) achieved diabetic remission; the  remaining four achieved superior glycemic control with lower insu­ lin dosages (median fructosamine pre‐ and postopera­ tively: 692 and 547μmol/L respectively; median insulin dose pre‐ and postoperatively: 20.5 and 3.5 units/kg/day respectively). Congestive heart failure was encountered as a transient problem in 4/19 cats that recovered from the surgery; all four cases occurred prior to implementing a reduction in volume of intravenous fluid delivered as part of the postoperative protocol. Two cats developed paresis of the left orbicularis oculi muscle, which resolved in the surviving cat. Cardiac arrest occurred in one cat postop­ eratively at time of ­jugular catheter placement, which was successfully revived and made an uneventful recovery. One cat developed a left pelvic limb monoparesis, which improved but did not resolve. Palatal wound breakdown was not encountered. This large case series suggests that hypophysectomy as  a  treatment for feline acromegaly results in a high i­ncidence of diabetic remission and reso­ lution of acromegaly. An alternative procedure, cryohypophysectomy, has been reported in a small number of cats but the proce­ dure has been less effective and results in a higher ­complication rate.

6  Feline Acromegaly

Radiation Radiation therapy is another option for the treatment of feline acromegaly, especially if the tumor is inoperable, the patient is not a suitable candidate for anesthesia, or surgical treatment is not available in the area. In human medicine, radiation therapy is regarded as a second‐line treatment as beneficial effects may take years to develop and the patient typically experiences undesired late‐term CNS radiation effects. The majority of studies that have been performed in vet­ erinary medicine focus on radiation treatment of pituitary masses regardless of functional status. There is no standard treatment protocol for pituitary masses in veterinary medi­ cine and varying methods have been used, including both single and multiple dose fractions administering total dos­ ages ranging from 1500 to 4000 cGY. The majority of the cats included in these studies had insulin‐resistant diabetes (suspected acromegaly or Cushing disease) and/or neuro­ logic signs. Radiation therapy was shown to be successful in  improving insulin resistance and neurologic signs. Neurologic improvement was generally seen within weeks to months and an improved insulin response was seen within the first month, but most patients still required insu­ lin therapy. In cases where repeat imaging was available, a decrease in tumor size was also noted. Disadvantages of radiation therapy are the early and delayed effects of radiation, repeated anesthesia, and expense. Early effects from radiation therapy include hair loss, skin pigmentation, and otitis externa. Reported late‐term side‐effects include brain necrosis, tumor regrowth, and visual and hearing impairment. In one study, 12 cats with pituitary tumors were treated with a coarse fractionated radiation protocol delivering a total dose of 37 Gy in five once‐weekly doses. Eight of these cats had insulin‐resistant diabetes mellitus secondary to acromegaly. Of these eight cats, five no longer required insulin therapy, two became stable diabetics, and one required less insulin. In addition, 3/4 cats had improved neurologic signs. The mean survival time of cats in this study was approximately 18 months. In another study, 14 cats with confirmed acromegaly and insulin‐resistant dia­ betes were treated with a total dose of 3700 cGy divided into 10 fractions (three per week). Thirteen of the 14 cats

had improved insulin responses. The average insulin dos­ age reduction was approximately 75%. Six of the cats went into complete diabetic remission and at the time the article was written, 3/6 remained in remission. The median sur­ vival time of cats in this study was 28 months. The most promising results with radiation may involve the use of stereotactic radiation therapy (SRT). Fifty‐ three client‐owned cats were referred to Colorado State University for SRT to treat pituitary tumors causing poorly controlled diabetes mellitus (DM) secondary to acromegaly. Diagnosis of acromegaly was based on history, physical examination, laboratory results, and cross‐sectional imaging of the pituitary. Signalment, radiation protocol, insulin requirements over time, adverse effects, and sur­ vival were recorded. Median survival time was 1072 days. Of the 41 cats for which insulin dosage information was available, 95% (39/41) experienced a decrease in required insulin dose, with 32% (13/41) achieving diabetic remis­ sion. Remission was permanent in 62% (8/13) and tem­ porary in 38% (5/13) cats. Median duration to lowest insulin dose was 9.5 months. Of the treated cats, 14% developed hypothyroidism and required supplementa­ tion after SRT. Cats treated with SRT have improved sur­ vival time and control of their DM when compared to previously reported patients treated with non‐SRT.

­Conclusion Feline acromegaly is likely an underdiagnosed disease in older male cats, especially in patients with insulin‐resistant diabetes. There is no single diagnostic test for acromegaly. The diagnostician should use history, clinical signs, labora­ tory tests (GH and IGF‐1), and advanced imaging to arrive at a diagnosis. There are several treatments options, but clinical studies on long‐term safety and efficacy are limited and often lack controls. Until more work is done evaluat­ ing medical treatments such as somatostatin analogs and growth hormone antagonists, most patients are best treated with either surgery or radiation therapy to control GH levels, improve glycemic control, and improve or pre­ vent the development of neurologic signs.

­Further Reading Abraham LA, Helmond SE, Mitten RW, et al. Treatment of an acromegalic cat with the dopamine agonist L‐deprenyl. Aust Vet J 2002; 80: 479–83. Abrams‐Ogg ACG, Holmberg DL, Stewart WA, et al. Acromegaly in a cat: diagnosis by magnetic resonance imaging and treatment by cryohypophysectomy. Can Vet J 1993; 34: 682–5.

Berg IM, Nelson RW, Feldman EC, et al.Serum insulin‐like growth factor‐I concentration in cats with diabetes mellitus and acromegaly. J Vet Intern Med 2007; 21: 892–8. Blois SL, Holmberg DL. Cryohypophysectomy used in the treatment of a case of feline acromegaly. J Small Anim Pract 2008; 49: 596–600.

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Brearly MJ, Polton GA, Littler RM, et al. Coarse fractionated radiation therapy for pituitary tumors in cats: a restrospective study of 12 cases. Vet Comp Oncol 2006; 4: 209–17. Dunning MD, Lowrie CS, Bexfield NH, et al. Exogenous insulin treatment after hypofractionated radiotherapy in cats with diabetes mellitus and acromegaly. J Vet Intern Med 2009; 23: 243–9. Elliott DA, Feldman EC, Koblik PD, et al. Prevalence of pituitary tumors among diabetic cats with insulin resistance. J Am Vet Med Assoc 2000; 216: 1765–8. Feldman EC, Nelson RW. Disorders of growth hormone. In: Canine and Feline Endocrinology and Reproduction, 3rd edn. Philadelphia, PA: WB Saunders, 2004, pp. 69–84. Feldman EC, Nelson RW. Acromegaly and hyperadrenocorticism in cats: a clinical perspective. J Feline Med Surg 2000; 2: 153–8. Goldman L, Schafer AI. Growth hormone excess: acromegaly and gigantism. In: Goldman‐Cecil Medicine, 24th edn. Philadelphia, PA: Saunders Elsevier, 2012, pp. 1437–8. Gostelow R, Scudder C, Keyte S, et al. Pasireotide long‐ acting release treatment for diabetic cats with underlying hypersomatotropism. J Vet Intern Med 2017; 31(2): 355–64. Kenny P, Scudder C, Keyte S, et al. Treatment of feline hypersomatotropism. Efficacy, morbidity and mortality of hypophysectomy. J Vet Intern Med 2015; 29: 1271. Kittleson MD, Pion PD, DeLellis DA, et al. Increased serum growth hormone concentration in feline hypertrophic cardiomyopathy. J Vet Intern Med 1992; 6: 320–4.

Meij BP, Auriemma E, Grinwis G, et al. Successful treatment of acromegaly in a diabetic cat with transsphenoidal hypophysectomy. J Feline Med Surg 2010; 12: 406–10. Meij BP, Voorhout G, van den Ingh TS, et al. Transsphenoidal hypophysectomy for treatment of pituitary‐dependant hyperadrenocorticism in seven cats. Vet Surg 2001; 30: 72–86. Niessen SJM, Khalid M, Petrie G, Church DB. Validation and application of an ovine radioimmunoassay for the diagnosis of feline acromegaly. Vet Rec 2007; 160: 902–7. Peterson ME, Taylor RS, Greco DS, et al. Acromegaly in 14 cats. J Vet Intern Med 1990; 4: 192–201. Posch B, Dobson J, Herrtage M. Magnetic resonance imaging finding in 15 acromegalic cats. Vet Radiol Ultrasound 2011; 4: 422–7. Scudder CJ, Niessen SJ, Catchpole B, et al. Feline hypersomatotropism and acromegaly tumorigenesis: a potential role for the AIP gene. Domest Anim Endocrinol 2017; 59: 134–9. Singerland LI, Voorhout G, Rijnberk A, et al. Growth hormone excess and the effect of octreotide in cats with diabetes mellitus. Domest Anim Endocrinol 2008; 35: 352–61. Starkey SR, Tan K, Church DB. Investigation of serum IGF‐I levels amongst diabetic and non‐diabetic cats. J Feline Med Surg 2004; 6: 149–55. Timian J, Lunn KF. Evaluation of a long‐acting somatostatin receptor ligand for the treatment of feline acromegaly. J Vet Intern Med 2012; 26: 757. Wormhoudt TL, Boss MK, Lunn K, et al. Stereotactic radiation therapy for the treatment of functional pituitary adenomas associated with feline acromegaly. J Vet Intern Med 2018; 32: 1383–91.

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7 Pituitary‐Dependent Hyperadrenocorticism in Dogs and Cats David S. Bruyette, DVM, DACVIM (SAIM) Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology and Epidemiology Pituitary‐dependent hyperadrenocorticism (PDH), also known as Cushing disease, is a common endocrine dis­ order in older dogs though relatively uncommon in cats. This disorder is caused by a pituitary adenoma (PA) that secretes inappropriate amounts of adrenocorticotropic hormone (ACTH), which results in bilateral adrenal hyperplasia and disorderly and excessive production of cortisol by the adrenal gland. Classification Basics of Classification

As with all central nervous system tumors, the Tumor– Node–Metastasis (TNM) system used by the World Health Organization does not apply. Current classifica­ tion systems for PAs in veterinary patients are based pri­ marily on secretory characteristics of the tumor. However, in humans, PAs are currently classified based upon: ●● ●●

tumor size and degree of invasiveness (Box 7.1) tumor endocrine activity (hormone secretion)

or functional classification based on immunohisto­ logic findings such as ACTH, thyroid‐stimulating hor­ mone (TSH), follicle‐stimulating hormone (FSH), etc. immunostaining. In both humans and animals, pituitary corticotroph adenomas that are responsible for Cushing disease (i.e., PDH) are classified as functional ACTH‐secreting PAs (ACTH‐PAs). Further Classification

The World Health Organization classification system for PAs in humans has been refined to include designations for benign adenoma, atypical adenoma, and pituitary

carcinoma on the basis of proliferation indices (p53 immunoreactivity, MIB‐I Index, mitotic activity) and the absence/presence of metastases. More comprehensive molecular classification systems based on relevant gene expression have not been system­ atically used to further characterize pituitary tumors. Similar work to classify canine pituitary tumors both morphologically and functionally is currently under way. Prevalence Humans

In humans, PAs are common tumors, with an overall prev­ alence in the general US population estimated at 16.7%. ●●

●●

●●

Corticotroph adenomas, comprising functional (ACTH‐PAs) and silent corticotroph adenomas, rep­ resent approximately 10–15% of all PAs. Functional ACTH‐PAs are the most common cause of Cushing syndrome (hypercortisolemia from any source), accounting for an estimated 70% of all cases. Prevalence of Cushing disease is estimated to be 1.2– 2.4 per 1 million people, and affects approximately 12 000 people in the US. This number, however, may be much higher, given that Cushing disease is fre­ quently misdiagnosed and diagnosis is often delayed.

Dogs

Functional ACTH‐PAs have a reported incidence of 0.2% in all dogs (1–2 cases/1000 dogs/year), with approxi­ mately 100 000 dogs affected yearly in the US. PDH accounts for approximately 85–90% of cases of hyper­ adrenocorticism, with the remainder of cases resulting from functional adrenal tumors, meal/food‐induced cases, occult or atypical disease. Meal‐ or food‐induced Cushing syndrome is thought to occur as the result of a congenital defect resulting in the aberrant expression of the gastric inhibitory polypeptide

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Box 7.1  Classification of pituitary tumors in humans by size and anatomic location Classification by size based on radiologic findings Microadenomas Macroadenomas

Less than 10 mm diameter Equal to or greater than 10 mm diameter

Classification based on radioanatomic findings Stage I Stage II Stage III Stage IV

Microadenomas (0.31 indicates an enlarged pituitary P/B ratio ≤0.31 indicates a nonenlarged pituitary.

A recent study investigated the expression of prolifera­ tion markers Ki‐67 and minichromosome maintenance‐7 (MCM7) in canine corticotroph adenomas in enlarged and nonenlarged pituitaries, and evaluated their relation to the size of canine pituitary corticotroph adenomas. ●●

●●

In the pituitary glands of humans and mice, cortico­ trophs and melanotrophs have a specific marker in com­ mon, the T‐box transcription factor (Tpit or Tbx19), which regulates the late differentiation of corticotrophs and melanotrophs and therefore may contribute to the pathogenesis of corticotroph adenomas. A recent study in 14 dogs with PDH examined the expression and mutation analysis of Tpit in normal canine pituitary and corticotroph adenomas. ●●

Nelson syndrome in which the pituitary tumor grows rapidly following bilateral adrenalectomy or suppressive medical therapy.

●●

Corticotroph and Melanotroph Cell Marker: Tpit

Canine corticotroph adenomas in enlarged pituitaries showed greater proliferation potential compared with adenomas in nonenlarged pituitaries. MCM7 expression was significantly greater than Ki‐67 expression in canine pituitary corticotroph adenomas.

Thus, MCM7 may be superior to Ki‐67 as a proliferation marker in canine pituitary tumors.

●●

Tpit was expressed in corticotroph and melanotroph cells of normal and adenomatous canine pituitary, and remained present in nonadenomatous corticotrophs of pituitaries from PDH dogs. No tumor‐specific mutation in Tpit cDNA from corti­ cotroph adenomas was found; however, a missense polymorphism (see Polymorphism versus Mutation) in the highly conserved DNA‐binding domain, the T‐ box, was discovered in one dog.

The study concluded that Tpit can be used as a reliable marker for corticotroph and melanotroph cells in canine pituitary tissue, but that mutations in the Tpit gene are unlikely to play a major role in pathogenesis of canine corticotroph adenomas. Corticotroph Differentiation Markers: LIF and LIFR

Leukemia inhibitory factor (LIF) is a cytokine of the interleukin (IL)‐6 family that activates the hypotha­ lamic–pituitary–adrenal axis and promotes corticotroph differentiation during development. LIF and leukemia inhibitory factor receptor (LIFR) expression were stud­ ied in pituitary glands of control dogs and specimens of corticotroph adenoma tissue were collected from dogs with PDH. The results demonstrated that: ●●

●●

LIFR is highly co‐expressed with ACTH and alpha‐ melanocyte‐stimulating hormone in the control canine pituitary gland and corticotroph adenomas there was a strong co‐expression of LIFR and ACTH1‐24 in the cytoplasm of cells in the pars distalis and pars intermedia of control pituitary tissue. In pituitary glands harboring an adenoma, the cytoplasmic expression of LIFR followed that of ACTH1‐24. Nontumorous cells of the pars distalis showed no cytoplasmic staining but did demonstrate nuclear to perinuclear immunoreactivity for LIFR in 10 of 12 tissue specimens from PDH dogs. This nuclear immunoreactivity was not observed in the control pituitary tissues or in the pituitary gland with corticotroph hyperplasia.

Role of ACTH Production and Glucocorticoids

As mentioned earlier, a characteristic biochemical feature of corticotroph adenomas is their relative resistance to negative feedback by glucocorticoids. In a recent study,

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gene expression related to ACTH production and secre­ tion and the negative feedback by glucocorticoids in canine corticotroph adenoma was evaluated in pituitary tumors in 10 dogs with Cushing disease. The results dem­ onstrated increased ACTH production and resistance to negative feedback by glucocorticoids in canine cortico­ troph adenomas. Therapeutic Role of EGFR

Since tumors in dogs and humans express epidermal growth factor receptor (EGFR), in another study we examined whether EGFR might provide a therapeutic target for Cushing disease. ●●

●●

In cell cultures from surgically resected human and canine corticotroph tumors, blocking EGFR also sup­ pressed expression of proopiomelanocortin (POMC), the ACTH precursor. In mouse corticotroph EGFR transfectants, ACTH secretion was enhanced and POMC promoter activity was increased.

In mice, blocking EGFR activity with gefitinib, an EGFR tyrosine kinase inhibitor: ●● ●●

●● ●●

attenuated POMC expression inhibited corticotroph tumor cell proliferation and induced apoptosis decreased both tumor size and corticosterone levels reversed signs of hypercortisolemia, including ele­ vated glucose levels and excess omental fat.

These study results indicate that inhibiting EGFR signal­ ing may be a novel strategy for treating Cushing disease.

Box 7.3  Common clinical signs of PDH in dogs and cats Polyuria and polydipsia Polyphagia Abdominal distension Bilaterally symmetric endocrine alopecia Panting Hypertension Urinary tract infections Additional dermatologic signs: – Thin skin – Pyoderma – Calcinosis cutis

Box 7.4  Common laboratory findings of PDH Hematologic abnormalities “Stress” leukogram (uncommon in cats): – Neutrophilic leukocytosis – Lymphopenia – Eosinopenia Mild thrombocytosis Mild erythrocytosis Serum biochemical abnormalities Increased serum alkaline phosphatase Milder increased in alanine aminotransferase Hypercholesterolemia Hypertriglyceridemia Hyperglycemia Urinalysis

­History and Clinical Signs Clinical signs, as well as laboratory abnormalities, seen in patients with PDH are secondary to the effects of ster­ oid excess, well recognized, and similar in scope to those seen with exogenous glucocorticoid supplementation (Boxes 7.3 and 7.4). The clinical signs of polyuria and polydipsia occur as the result of excessive cortisol interfering with pituitary release of antidiuretic hormone (ADH) or the binding of ADH to receptors in the renal tubules. Abdominal distension and thinning of skin occur due to the cata­ bolic effects of cortisol on tissues such as muscle and connective tissue. Hepatomegaly steroid‐induced vacu­ olar hepatopathy also contributes to the “pot‐bellied” appearance. The endocrine alopecia mirrors the known distribution of sex hormone receptors in the skin with  endocrine alopecias often sparing the head and extremities. Panting occurs in  both dogs and humans

Decreased urine specific gravity osmoreceptors sense decrease in osmolality > decrease AVP release > compensatory polyuria

It is important to note, however, that patients with hypoadrenocorticism will have a low sodium level due to mineralocorticoid deficiency despite primary polyuria. Plasma osmolality can also be used to determine if a patient has primary polydipsia. In clinical practice, this can be estimated by calculating serum osmolality using the following formula: 2 × [Na] + [BUN in mg/dL]/2.8+ [Glu in mg/dL]/18 Low osmolality is suggestive of primary polydipsia and further evaluation for hepatic disease or hyperthyroidism are indicated. Second‐Tier Diagnostic Tests Further diagnostics will depend on physical and laboratory examination findings. For example, an abdominal ultrasound should be considered for patients with flank/abdominal pain in whom pyelonephritis is suspected. For patients in whom adrenal disease is suspected, an adrenocorticotrophic hormone (ACTH) stimulation test should be considered. It is important to remember that approximately 15% of dogs with pituitary dependent and 40% of dogs with adrenal‐dependent hyperadrenocorticism can have a normal ACTH stimulation test. If hyperadrenocorticism is strongly suspected and the dog is otherwise healthy, a low‐ dose dexamethasone suppression test is a better option. Other tests that may be considered, depending on laboratory findings, clinical signs, and signalment, include pre‐ and postprandial bile acids and imaging studies of the chest and abdomen. Imaging will be particularly useful for patients suspected of a neoplastic process and in any patient with hypercalcemia. Abdominal ultrasonography may be useful in looking for masses, enlarged abdominal lymph nodes, or adrenal enlargement.

In addition to being time‐consuming and difficult to perform, for patients with underlying undiagnosed illness, this test is contraindicated and may increase the risk for complications, including death. This test should never be performed in patients who are azotemic or dehydrated. In addition, diseases such as hyperadrenocorticism may show some positive response, thereby complicating the diagnosis or resulting in misdiagnosis. Steps for Conducting the Modified WDT

Phase 1 Patients that have had PU/PD for a prolonged period of time may have wash‐out of their renal medullary concentration gradient and therefore, may be unable to concentrate their urine in the face of water deprivation. For this reason, it is recommended that once the pet’s water intake has been quantified, the total amount of water given should be gradually restricted over 3–5 days until the goal of 60–80 mL/kg/day is reached. To avoid prolonged periods of water restriction, the total daily water should be calculated and provided in frequent small portions. A dry food diet should be fed during this phase. This testing will require that the owners pay very close attention to the pet for any signs of illness.  Patients with severe polyuria are at increased risk for hypertonic dehydration and hospitalization during this phase of the WDT should be considered. A USG should be measured at the conclusion of this phase in case the patient has already reached a USG >1.030 (supportive of a diagnosis of psychogenic polydipsia) and the WDT is no longer required. Phase 2 ●●

●●

●●

●● ●● ●●

Modified Water Deprivation Test (WDT) or DDAVP trial A modified WDT should only be considered for patients in whom more common causes of PU/PD have been ruled out and central DI, primary NDI, or psychogenic polydipsia is strongly suspected. Fortunately, very few cases of PU/PD require performing this test.

●●

●●

The patient should be admitted to the hospital for a complete physical examination and assessment of hydration and mentation status. An indwelling urinary catheter is placed. The bladder should be emptied completely, and the  USG measured. A blood urea nitrogen (BUN), creatinine, packed cell volume (PCV), total solids (TS), and electrolytes should be measured. Obtain patient’s body weight. Patient should not have access to food or water. Patient should have the following performed every 1–2 hours. –– Physical examination to assess hydration and mentation –– Completely empty bladder and check USG –– Obtain weight Every 4–6 hours check BUN, creatinine, PCV, TS, and electrolytes. The test should be stopped when any of the following occur:

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Table 8.1  Currently available DDAVP preparations and common dosages Preparation

Strength

Dose

Intranasal

100 μg/mL (1 drop = 1.5–4.0 μg desmopressin)

1–2 drops in conjunctival sac one to two times daily Or 2 μg SC one to two times daily (can be administered SC if sterilized through a bacteriostatic filter)

Injectable

4 μg/mL

2 μg SC one to two times daily

Oral

0.1–0.2 mg tablets (each 0.1 mg = approximately 4 μg desmopressin)

0.05–0.2 mg PO twice daily Bioavailability of the oral preparation may be a concern

PO, orally (per os); SC, subcutaneously.

–– The patient loses 5% of body weight. Maximal stimulation of AVP release occurs at this point but the time required to reach dehydration may be a few hours or up to 12 hours in some patients. –– There is evidence of clinical dehydration or any vomiting, lethargy, or altered mentation is noted. –– Any evidence of azotemia. –– The USG is >1.030. Phase 3 ●●

●●

If the patient is 5% dehydrated and has not produced an adequately concentrated urine, desmopressin acetate (DDAVP, a synthetic AVP analog) should be administered (see Table 8.1 for a list of desmopressin formulations). –– Assess mentation and measure weight, BUN, creatinine, PCV, TS, electrolytes. –– Bladder should be emptied. –– Administer injectable DDAVP (2-10    μg IV). Alternatively, an intranasal DDAVP formulation may be administered into the conjunctival sac (1–4 drops/dog). Empty the bladder in 30 minutes and measure USG. Repeat every 30 minutes for up to two hours until USG >1.015. Bladder should be emptied at each time point. If USG is still below 1.015, empty bladder and measure USG every hour for up to 8 hours. At the conclusion of the test, water can be reintroduced slowly  (10-20 mL/ kg every 30 minutes for 2 hours).

Interpretation

If the patient achieved an adequate urine concentration during phase 1 of the test, then a diagnosis of primary (psychogenic) polydipsia can be made. AVP secretion and responsiveness are intact.

If concentrated urine was not noted at the end of phase 1, then the patient has primary polyuria – either CDI or primary NDI (causes of secondary NDI should have been ruled out prior to the WDT). Administration of DDAVP differentiates between these two possibilities. A patient with CDI will achieve an increase in USG (>1.015) after DDAVP administration whereas a patient with NDI will show little to no response. Again, patients with secondary causes of NDI (i.e., hyperadrenocorticism) may also fail to show an elevation in USG after DDAVP administration, which is why it is imperative that secondary causes be ruled out prior to the WDT. Interpretation may also be complicated in cases of pati­ ents with concurrent primary polydipsia and CDI or NDI. DDAVP Response Test

In cases where more common causes of PU/PD have been ruled out and where there is a high index of suspicion for CDI, a DDAVP response test may be conducted in lieu of a WDT. This can be conducted at home as the owner continues to monitor water intake. ●●

●●

Measure USG on day 1 and administer DDAVP intranasal preparation (1–4 drops) into the conjunctival sac twice daily for 5–7 days. Alternatively, the owner may administer oral DDAVP (tablet) at a dose of 0.1–0.2 mg every 8 hours for 5–7 days. After 5–7 days, the USG is measured again.

Animals with CDI should demonstrate an increase in the USG as well as a decrease in water intake. An animal with NDI or PP will show no response. Patients with primary polydipsia may continue to drink excessively despite the DDAVP and there is a risk of water intoxication in these patients, which is why appropriate patient selection and owner vigilance are imperative.

­Therapy Treatment will need to address the underlying disorder. For patients with CDI, DDAVP will need to be administered daily. DDAVP can be administered topically in the conjunctival sac or a subcutaneous injection. An oral preparation is also available but may be more costly and has been shown to be less effective than parenteral administration in some patients. See Table  8.1 for recommended dosing schedules. Regardless of the route of administration, the DDAVP dose should be titrated up gradually as needed to control the PU/PD as the effects last 8–24 hours. For patients with primary psychogenic polydipsia, gradual water restriction and behavioral modification therapy may be warranted. Consultation with or referral to a behaviorist may be considered.

8  Polyuria and Polydipsia

­Further Reading DiBartola SP. Disorders of sodium and water: hypernatremia and hyponatremia. In: DiBartola SP, ed. Fluid, electrolyte, andacid-base disorders. 4th Edition. St. Louis: Saunders Elsevier; 2012. Feldman EC, Nelson RW. Canine and Feline Endo­ crinology and Reproduction, 4th ed. St Louis, MO: Elsevier, 2015. Guillaumin J, DiBartola SP. Disorders of sodium and water homeostasis. Vet Clin North Am Small Anim Pract. 2017 Mar; 47(2). Harb MF, Nelson RW, Feldman EC, et al. Centraldiabetes insipidus in dogs: 20 cases (1986–1995). J Am Vet Med Assoc. 1996 Dec1; 209(11).

Nichols R, Peterson ME. Clinical use of vasopressinanalog desmopressin for the diagnosis and treatment of diabetes insipidus. In: Bonagura JD, Twedt D, ed’s. Kirk’s Current Veterinary Therapy XV. Philadelphia: Elsevier Saunders; 2014. Rothuizen J, Biewenga WJ, Mol JA. Chronic glucocorticoidexcess and impaired osmoregulation of vasopressin release in dogs with hepaticencephalopathy. Domest Anim Endocrinol. 1995 Jan; 12(1). Teshima T, Hara Y, Taoda T, et al. Central diabetes insipidus after transsphenoidal surgeryin dogs with Cushing’s disease. J Vet Intern Med. 2011 Jan; 73(1).

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9 Canine Hypothyroidism David S. Bruyette, DVM, DACVIM (SAIM) Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology and Epidemiology Canine hypothyroidism, while a common endocrinopathy in the dog, may be overdiagnosed due to confusion/ inconsistencies in establishing a definitive diagnosis. Hypothyroidism is due to decreased thyroidal production of the thyroid hormones thyroxine (T4) and triiodothyronine (T3). Greater than 90% of cases are primary and are due to acquired immune‐mediated destruction of the thyroid gland which is preceded by thyroiditis, idiopathic atrophy or, less commonly, neoplasia. Secondary forms of the disease include thyroid‐stimulating hormone (TSH) deficiency, pituitary neoplasia, and cystic Rathke’s pouch. Tertiary hypothyroidism with thyrotropin‐releasing hormone (TRH) deficiency has not been documented in dogs. Congenital cases have been reported in both dogs and cats. Hypothyroidism most commonly occurs in young to middle‐aged dogs with an average age of 7 years. Dogs with autoimmune disease tend to develop hypothyroidism at a younger age. While thyroid values decrease within the reference range in senior dogs, hypothyroidism is very uncommon and other factors (see later) are likely responsible for the observed decreased thyroid concentrations in euthyroid older patients. Spayed females and neutered males are at increased risk when compared to sexually intact animals. Breed predispositions have been reported for golden retrievers and Doberman pinschers. Thyroiditis is heritable in the beagle, borzoi, golden retriever, Great Dane, Irish setter, Doberman pinscher, and old English sheepdogs. No known environmental factors have been identified.

­History and Clinical Signs As thyroid hormone regulates the metabolic rate and influences the functions of many organs, clinical signs are often nonspecific and insidious in onset. Many other diseases can have similar clinical signs to hypothyroidism, which may lead to an incorrect diagnosis. As such, laboratory testing of thyroid function is often performed as part of the diagnostic work‐up in animals with nonthyroidal illness. Common clinical signs include lethargy, mental dullness, weight gain, exercise intolerance, alopecia, and obesity. Many metabolic, infectious, neoplastic, congenital, degenerative, and inflammatory diseases can cause similar clinical signs and biochemical abnormalities seen with hypothyroidism.

­Diagnosis Laboratory Diagnosis Thyroxine is the major secretory product of the thyroid while the majority of T3 is derived from extrathyroidal sources. Both T4 and T3 are highly protein bound to serum carrier proteins such as thyroid‐binding globulin, transthyretin, and albumin. Only unbound (free) hormone is able to penetrate cell membranes, bind to receptors, and result in biologic activity. Protein‐bound hormone acts as a reservoir to maintain steady concentrations of free hormone in the plasma despite rapid alterations in release and metabolism of T3 and T4 and changes in the plasma protein concentrations.

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Serum Total T4

Serum T4 is a sensitive (>90–95%) but not specific test (70–75%) for the diagnosis of canine hypothyroidism. The vast majority of dogs with hypothyroidism have a serum T4 below normal, but some normal dogs and those with a variety of other problems may have a low serum T4. A diagnosis of hypothyroidism can be ruled out if the T4 is in the upper 50% of the reference range. Autoantibodies to T4 occur in about 15% of hypothyroid dogs, and these antibodies may falsely increase the serum T4 concentration from below normal into or above the normal range. In‐house testing of total T4 is not recommended. Serum Total T3

Serum T3 concentration is an unreliable test for evaluation of thyroid function. Serum Free T4 (fT4)

Thyroxine is highly (99.9%) protein bound in the circulation. Protein binding can be altered by many nonthyroidal illnesses and by certain drugs. Measurement of the unbound or free hormone can provide a more accurate assessment of thyroid function in these cases (sensitivity >95%, specificity >97%). The sensitivity of fT4 is equivalent to or slightly better than total T4 in diagnosing hypothyroidism in routine cases. More importantly, fT4 is more specific, particularly when nonthyroidal factors that can influence total T4 are present. Free T4 is less affected by most nonthyroidal illness and drugs, but still can be altered in cases of moderate to severe illness. In addition, fT4 by equilibrium dialysis is not affected by the presence of T4 autoantibodies that will falsely elevate total T4. Measurement of fT4 by equilibrium dialysis should be performed when uncommon clinical signs of hypothyroidism are present, the dog is being treated with a drug that may affect thyroid function, when nonthyroidal illness is present, and if autoantibodies to T4 are detected. Serum TSH

Primary hypothyroidism results in a decrease in T4 and thus decreased negative feedback on the pituitary gland. In response, the pituitary secretes more TSH and plasma TSH levels increase. In humans, TSH is elevated prior to any decrease of T4 or fT4 outside the normal range. In the dog, TSH concentration is elevated in only 65–75% of cases of hypothyroidism, and thus it lacks sensitivity for use as a screening test. The combination of decreased total T4 or fT4 with an elevated serum TSH is diagnostic of hypothyroidism (specificity >95%). Therefore, a normal TSH does not rule out hypothyroidism, but an

e­ levated TSH combined with a low T4 or fT4 provides a definitive diagnosis. Diagnosis of Thyroiditis

Antibodies against either T4 or T3 or both are sometimes present in dogs with thyroiditis with or without hypothyroidism. The presence of these antibodies does not indicate that the dog is hypothyroid, but suggests that autoimmune thyroid disease is present. These antibodies frequently cause false elevation of T4 or T3 concentrations that can result in marked elevation of the hormones. Autoantibodies to T4 are present in about 10–15% of hypothyroid dogs. Dogs with autoimmune thyroiditis may have circulating antibodies to thyroglobulin, the primary protein in the colloid of the thyroid gland. This is not a test of thyroid function but rather a marker for the presence of autoimmune thyroiditis. In one long‐term study at Michigan State University, 20% of asymptomatic, antithyroglobulin‐positive dogs with normal thyroid function progressed to hypothyroidism in one year. The presence of these antibodies in a dog with borderline laboratory evidence of hypothyroidism and clinical signs supports a diagnosis of hypothyroidism. Additional Considerations Breeds

Certain breeds have normal ranges of thyroid hormones that are different from most other breeds. Few have been evaluated but greyhounds have serum total T4 and fT4 concentrations that are considerably lower than most other breeds. Scottish deerhounds, salukis, and whippets also have total T4 concentrations that are well below the mean concentration of dogs in general. Alaskan sled dogs have serum T4, T3, and fT4 concentrations below the reference range of most pet dogs, particularly during periods of intense training or racing. Time of Day

In one study 50% of normal dogs had a low serum T4 concentration at some time during the day. Medications

The drugs that are known to commonly alter thyroid function tests are glucocorticoids, phenobarbital, sulfonamides, clomipramine, aspirin, and some other NSAIDs. Glucocorticoids suppress total T4 and sometimes fT4 as well. Phenobarbital causes decreased total T4 and mild increases in TSH. Sulfonamides can induce

9  Canine Hypothyroidism

overt primary hypothyroidism with clinical signs and thyroid function tests that support the diagnosis. The changes may be reversible when the medication is discontinued. There are dozens of drugs that affect thyroid function and thyroid function tests in humans, so many others likely affect the dog as well. Nonthyroidal Illness

Illness not involving the thyroid gland can alter thyroid function tests and has been labeled “nonthyroidal illness” or “euthyroid sick syndrome.” Any illness can alter thyroid function tests, causing a fairly consistent decrease in total T4 and T3 concentrations in proportion to the severity of illness. Serum TSH concentration is increased in 8–10% of dogs with nonthyroidal illness. Serum fT4 measured by equilibrium dialysis is less likely to be  affected, but can also be increased or decreased. However, in dogs with substantial nonthyroidal illness, the fT4 is likely to be decreased. It is recommended that testing of thyroid function be postponed until the nonthyroidal illness is resolved. If this is not possible, measurement of T4, TSH, and fT4 is indicated. Ancillary Testing Thyroid Gland Ultrasound

Although rarely necessary, ultrasound of the thyroid glands (by an experienced ultrasonographer) can aid in differentiating dogs with primary hypothyroidism from those with nonthyroidal illness. Thyroid glands of hypothyroid dogs tend to be smaller, less homogeneous, and hypoechoic than those of euthyroid dogs. There is considerable overlap with the ultrasonographic appearance and size of the thyroid glands of euthyroid and hypothyroid dogs. Thyroid ultrasound can only be used to help support a diagnosis of hypothyroidism if the thyroid glands are quite small.

­Treatment Levothyroxine is the only hormone that appears necessary for treatment of hypothyroidism. The frequency of levothyroxine dosing is controversial, and the only study to closely evaluate the response to treatment showed that once‐daily treatment is adequate. However, in clinical practice some dogs seem to respond better to twice‐ daily treatment. The initial starting dose is 0.02 mg/kg PO q24h. In general, you will never have to exceed 0.8 mg as an

initial daily dosage even in very large dogs. If the dog has significant cardiovascular disease, diabetes mellitus, or hypoadrenocorticism, treatment should be instituted at 25% of the standard dose, with the ­dosage  increased by 25% every two weeks based on clinical response and postpill testing. Most dogs show  improvement within the first 1–2 weeks, with increased activity, improved attitude, and partial or complete resolution of neurologic signs. The cutaneous manifestations of hypothyroidism may take several weeks to months to resolve. Posttreatment monitoring may be carried out but clinical response is the most important monitoring tool. Peak T4 concentrations generally occur 4–6 hours after administration of levothyroxine and should be in the high normal to slightly above normal range (40– 70 nmol/L). However, the bioavailability of thyroxine ranges from 13% to 87% in the same dog from day to day, bringing into the question the utility of random postpill monitoring of TT4. It is likely more meaningful (though more expensive) to measure TSH (especially if the TSH concentration was elevated pretreatment) or fT4 concentrations after replacement therapy has been started, especially in animals that show a poor clinical response to therapy. Serum TSH concentrations should be in the normal range or undetectable and fT4 concentrations should be in the normal range. Serum concentrations of TSH and fT4 should not be performed until  the patient has been on supplementation for at least two weeks. If the patient was initially started on twice‐daily therapy, treatment can be reduced to once‐ daily treatment when a good clinical response has been obtained. Hyperthyroidism is the most common complication of treatment with levothyroxine, but it is rare in dogs. Clinical signs are similar to those of hyperthyroidism in cats and the diagnosis is confirmed by documenting a substantial elevation of serum T4. Treatment consists of stopping levothyroxine treatment for 2–3 days, then instituting treatment at a lower dose.

­Prognosis Response to therapy should be observed in the first 4–8 weeks post treatment. Improvements in mentation and physical activity may be noted within the first week though some abnormalities, especially dermatologic signs, may take several months to resolve. An absent or incomplete response to therapy may be due to an incorrect diagnosis, poor owner compliance, inadequate dosing, or poor absorption.

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­Further Reading Bellumori TP, Famula TR, Bannasch DL, Belanger JM, Oberbauer AM. Prevalence of inherited disorders among mixed‐breed and purebred dogs: 27,254 cases (1995–2010). J Am Vet Med Assoc 2013; 242(11): 1549–55. Lewis VA, Morrow CMK, Jacobsen JA, Lloyd WE. A pivotal field study to support the registration of levothyroxine sodium tablets for canine hypothyroidism. J Am Anim Hosp Assoc 2018; 54: 201–8. Panciera DL, Purswell BJ, Kolster KA, Werre SR, Trout SW. Reproductive effects of prolonged experimentally

induced hypothyroidism in bitches. J Vet Intern Med 2012; 26(2): 326–33. van Dijl IC, Le Traon G, van de Meulengraaf BD, Burgaud S, Horspool LJ, Kooistra HS. Pharmacokinetics of total thyroxine after repeated oral administration of levothyroxine solution and its clinical efficacy in hypothyroid dogs. J Vet Intern Med 2014; 28: 1229–34. Ziglioli V, Panciera DL, Troy GC, Monroe WE, Boes KM, Refsal KR. Effects of levothyroxine administration and withdrawal on the hypothalamic‐pituitary‐thyroid axis in euthyroid dogs. Vet Intern Med 2017; 31: 705–10.

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10 Feline Hyperthyroidism David S. Bruyette, DVM, DACVIM (SAIM) Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology and Epidemiology Two recent large studies have looked at possible environ­ mental or dietary factors involved in the pathogenesis of hyperthyroidism. One of the studies with a case–control design looked at 100 cats with hyperthyroidism and 163 control cats. The cats’ medical records were reviewed and the owners were asked to complete a mailed ques­ tionnaire. Data included demographic variables, envi­ ronmental exposures, and diet to include the preferred flavors of canned cat food. In this study, housing, expo­ sure to fertilizers, herbicides, regular use of flea prod­ ucts, and the presence of a smoker in the house were not associated with an increased risk but cats that preferred fish or liver and giblets flavors of canned cat food had an increased risk. The results suggested that cats that prefer to eat certain flavors of canned cat food may have a sig­ nificantly increased risk of hyperthyroidism. In the second case–control study, owners of 379 hyper­ thyroid and 351 control cats were questioned about their cats’ exposure to potential risk factors, including breed, demographic factors, medical history, indoor environ­ ment, chemicals applied to the cat and environment, and diet. The association between these hypothesized risk factors and outcome of disease was evaluated. Two genetically related cat breeds (Siamese and Himalayan) were found to have diminished risk of developing hyper­ thyroidism. Cats that used litter had higher risk of devel­ oping hyperthyroidism than those that did not. Use of topical ectoparasite preparations was associated with increased risk of developing hyperthyroidism. Compared with cats that did not eat canned food, those that ate commercially prepared canned food had an approximate twofold increase in risk of disease. When these four vari­ ables (breed, use of cat litter, consumption of canned cat food, and use of topical ectoparasite preparations) from

the univariate analysis were selected for further, a persis­ tent protective effect of breed (Siamese or Himalayan) was found. In addition, results suggested a 2–3‐fold increase in risk of developing hyperthyroidism among cats eating a diet composed mostly of canned cat food and a threefold increase in risk among those using cat litter. In contrast, the use of commercial flea products did not retain a strong association. The results of this study indicate that further research into dietary and other potentially important environmental factors (cat litter) is warranted. Altered G protein expression was found in thyroid gland tissue from hyperthyroid cats compared to normal control cats. Adenomatous thyroid glands obtained from eight hyperthyroid cats and thyroid glands obtained from four age‐matched euthyroid cats were examined for expression of G inhibitory protein (Gi) and G stimulatory protein (Gs). Expression of Gi was significantly reduced in thyroid gland adenomas from hyperthyroid cats, com­ pared with normal thyroid gland tissue from euthyroid cats. Expression of Gs was similar between the two groups. A decrease in expression of Gi in adenomatous thyroid glands of cats may reduce the negative inhibition of the cAMP cascade in thyroid cells, leading to autono­ mous growth and hypersecretion of thyroxine. What we don’t know is what causes the reduction in Gi in hyper­ thyroid cats. The factors mentioned above in the studies of environmental and dietary risk factors may play a role in altering the G protein expression found in this study. Oncogenes and the tumor suppressor gene p53 were examined in cats with hyperthyroidism. Formalin‐fixed, paraffin‐embedded thyroid glands from 18 cats diag­ nosed with hyperthyroidism were evaluated immunohis­ tochemically for overexpression of the products of oncogenes c‐ras (a mitogenic oncogene) and bcl2 (an apoptosis inhibitor) and the tumor suppressor gene p53.

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Fourteen thyroid glands from euthyroid cats without his­ tologically detectable thyroid lesions were examined similarly as controls. Results from these investigations showed that all cases of nodular follicular hyperplasia/ adenomas stained positively for overexpression of c‐Ras protein using a mouse monoclonal antihuman pan‐Ras antibody. The most intensely positively staining regions were in luminal cells surrounding abortive follicles. Subjacent thyroid and parathyroid glands from euthy­ roid cats did not stain immunohistochemically for pan‐ Ras. There was no detectable staining for either Bc12 or p53 in any of the cats. These results indicated that over­ expression of c‐ras was highly associated with areas of nodular follicular hyperplasia/adenomas of feline thy­ roid glands, and mutations in this oncogene may play a role in the etiopathogenesis of hyperthyroidism in cats. As with the study on G protein abnormalities, c‐ras mutations could either be an initiating cause of hyper­ thyroidism or simply mediate the effects of an as yet uni­ dentified dietary or environmental initiator. Alterations in the thyrotropin (TSH) receptor were also examined in cats with hyperthyroidism. The authors used the polymerase chain reaction (PCR) to amplify codons 480–640 of the previously uncharacterized feline thyrotropin receptor (TSHR) gene, and determined the DNA sequence in this transmembrane domain region. They then analyzed single‐stranded conformational pol­ ymorphisms in thyroid DNA from 11 sporadic cases of feline thyrotoxicosis and leukocyte DNA from two cases of familial feline thyrotoxicosis. They also determined the DNA sequence of this region of the TSHR in five of the cases of sporadic feline thyrotoxicosis and the two familial thyrotoxic cats. The normal feline TSHR sequence between codons 480 and 640 is highly homolo­ gous to that of other mammalian TSHRs, with 95%, 92%, and 90% amino acid identity between the feline receptor and canine, human, and bovine TSHRs, respectively. Thyroid gland DNA from 11 cats with sporadic thyro­ toxicosis did not have mutations in this region of the TSHR gene. Leukocyte DNA from two littermates with familial feline thyrotoxicosis did not harbor mutations of this region of the TSHR gene. These studies suggested that TSHR gene mutations are likely not involved in feline hyperthyroidism. Since its first description in 1979, the incidence of hyperthyroidism has dramatically increased, prompting veterinarians and researchers to hypothesize whether exposure to environmental thyroid‐disruptor chemicals or other environmental, genetic or dietary factors are  involved in the pathogenesis of hyperthyroidism. Potential exposure to several substances has been impli­ cated, including organohalogen compounds such as pol­ ychlorinated biphenyls and polybrominated diphenyl ethers, fertilizers, soy isoflavones, bisphenol‐A primarily

released from “pop‐top” canned cat food lids, and con­ sumption of commercial canned food. One theory as to why the number and percentage of unexpected outliers becomes accelerated over the age of 9 years is that it may take several years of exposure to such environmental, dietary, and genetic factors before they express them­ selves clinically and hyperthyroidism ensues, although this topic requires further investigation.

­History and Clinical Signs With time, we have seen both an increase in the diagno­ sis of hyperthyroidism and a decrease in the severity of the clinical signs associated with thyrotoxicosis. This is most likely due to an increased awareness on the part of the pet owner and the veterinarian as well as the increased use of T4 concentrations as an integral part of routine feline health screening. We have also seen addi­ tional work on some of the less obvious manifestations of hyperthyroidism such as hypertension which may be clinically silent and/or present initially with ocular signs, as well as the effects of hyperthyroidism on the cardio­ vascular and renal system (to be discussed later). As stated earlier, the clinical signs associated with hyperthyroidism have been decreasing in severity over the years (Box  10.1). A paper examined the electrocar­ diographic and radiographic changes seen in hyperthy­ roid cats today versus those seen 10–12 years ago. Two populations (1992–1993 and 1979–1982) of confirmed hyperthyroid cats were compared to determine whether the incidence of certain cardiovascular specific manifes­ tations of feline thyrotoxicosis had experienced similar changes. Sinus tachycardia, which is the most commonly recognized cardiac manifestation of feline thyrotoxico­ sis, was not as prevalent in the 1993 group when com­ pared to the 1982 group. This was also true for the finding of an increased R‐wave amplitude on lead II elec­ trocardiography. Both groups had a similar low inci­ dence of atrial and ventricular dysrhythmias; however, the 1993 group had a significantly higher occurrence of right bundle branch block. Thoracic radiographs were deemed necessary in a larger proportion of the 1982 group compared to the 1993 group. Although there were Box 10.1  Clinical signs of feline hyperthyroidism Weight loss and poor hair coat Aggressive or “cranky” behavior Periodic vomiting Polyuria and polydipsia Increased appetite, activity, restlessness, and heart rate Occasionally, difficulty breathing, weakness, and depression

10  Feline Hyperthyroidism

no significant differences in radiographically defined cardiac size between the two groups, a larger number of cats in the 1982 group had evidence of congestive heart failure. These findings suggest that feline hyperthyroid­ ism is being diagnosed earlier and with less severe clini­ cal signs than when studied a decade ago.

­Diagnosis Diagnosis most often is based on the presence of one or more typical clinical signs and increased serum total thyroxine (T4) concentration. However, up to 10% of all hyperthyroid cats and 40% of those with mild dis­ ease have serum T4 values within reference range. The diagnosis of hyperthyroidism should not be excluded on the basis of a single normal serum T4 value, espe­ cially in a cat with typical clinical signs, a palpable thy­ roid nodule and serum T4 in the upper half of the normal range. In these cases, serum free T4 (fT4), measured by equilibrium dialysis, may provide an alternative means of diagnosing hyperthyroidism in cats with normal serum total T4 values. Studies docu­ ment that up to 20% of sick euthyroid cats can have increased fT4 concentration. Therefore, it is most appropriate and reliable to interpret the two values together. Mid‐to‐high reference range total T4 and increased fT4 concentration are consistent with hyper­ thyroidism. In contrast, low total T44and increased fT4 values are usually associated with nonthyroidal illness. In cases where a strong index of suspicion exists for hyperthyroidism and test results are equivocal, thyroid imaging can be of benefit.

­Treatment Once hyperthyroidism has been diagnosed, all manage­ ment options (thyroidectomy, radioactive iodine, antithyroid drugs, nutritional management) should be discussed with pet owners. All options can be ≥90% effective for controlling hyperthyroidism when used appropriately. The selected management option will differ for each cat based on several considerations (Table  10.1). Radioactive iodine therapy is considered the gold standard for treatment of hyperthyroidism but most pet owners currently opt for medical manage­ ment. Until recently, this included oral or transdermal antithyroid drugs. Now nutritional management using a limited‐iodine food is another option for cats with hyperthyroidism. Radioactive Iodine Radioiodine treatment is often considered the best option for many hyperthyroid cats because: ●●

●●

●● ●●

it has the potential to eliminate a benign thyroid tumor or abnormal thyroid tissue with a single treatment it treats extrathyroidal thyroid tissue, which may occur in 10–20% of hyperthyroid cats no general anesthesia is required reported side‐effects are minimal.

Cats should be stable prior to radioiodine therapy; those with clinically significant cardiovascular, renal, gastroin­ testinal, or endocrine (e.g., diabetes mellitus) disease may not be very good candidates, especially because of the time necessary for boarding after treatment.

Table 10.1  Advantages/disadvantages of options for managing cats with hyperthyroidism Option

Advantages

Disadvantages

Thyroidectomy

Cures current tumor

High initial costs Requires anesthesia Hospitalization required Risk of postoperative hypocalcaemia Irreversible

Radioactive iodine

Cures current tumor Single treatment Effective for ectopic tissue Side‐effects uncommon

High initial costs Limited availability Hospitalization required Irreversible

Antithyroid drugs

Routinely available Reversible Costs spread over time

Not curative (controls T4 and signs) Daily administration needed Drug side‐effects

Limited‐iodine food

Routinely available Reversible Costs spread over time

Not curative (controls T4 and signs) Cat can only eat a single food

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After administration, radioactive iodine is actively concentrated by the thyroid gland and has a half‐life of eight days. It emits both beta‐particles and gamma‐radi­ ation; the beta‐particles are responsible for the majority of tissue destruction, but are only locally destructive, traveling a maximum of 2 mm. Therefore, no significant damage to adjacent parathyroid tissue, atrophic thyroid tissue, or other cervical structures is expected. The main limitation to widespread use of radioactive iodine is the requirement for special licensing and the isolation of the cat for variable periods after treatment. This can range from several days to several weeks depending on state or local radiation regulations and the dose administered. The goal of treatment is to restore euthyroidism with  the smallest possible single dose of radioactive iodine,  while avoiding development of hypothyroid­ ism. Controversy exists as to the best method of calcu­ lating the optimum dose for individual cats. Based on the majority of reported cases, posttreatment hypothy­ roidism is transient and generally uncommon (2–7% of cases); even fewer cats have clinical signs or appear to require thyroid hormone replacement. However, up to 30% (50 of 165 cats) were hypothyroid three months after radioactive iodine therapy in one study; of these, 56% (19 of 34 hypothyroid cats with available informa­ tion) had clinical signs of hypothyroidism and 52% (23 of 44 cats) were given thyroid hormone supplementa­ tion. Thyroid hormone replacement may be needed in some cats, especially those with concurrent kidney dis­ ease, since hypothyroidism has been associated with azotemia and decreased survival time in previously hyperthyroid cats. Owners should be advised of this possibility, particularly if their motivation is to avoid long‐term oral medication. Antithyroid Drugs Antithyroid drugs (e.g., methimazole, carbimazole) are commonly used for treatment of hyperthyroidism in cats. If administered appropriately, they reliably inhibit the synthesis of thyroid hormones and thereby lower serum thyroid hormone concentrations. These drugs do not affect the thyroid gland’s ability to trap inorganic iodide or release preformed hormones. They are widely recommended to stabilize hyperthyroid cats prior to surgery and are the only drugs that can be used chroni­ cally for management of hyperthyroidism. Almost all cats are potential candidates unless thyroid carcinoma is suspected. Antithyroid drugs used most often in cats include methimazole and carbimazole; both can be given orally or formulated for transdermal application. Custom for­ mulation of transdermal products may increase the expense of therapy and stability of the product is not

guaranteed. Results of a recent prospective study con­ ducted in New Zealand showed that once‐daily treat­ ment for 12 weeks with transdermal methimazole in a novel lipophilic vehicle was as effective as twice‐daily carbimazole administered orally. While many cats have been successfully managed long term with antithyroid drugs, it is important to monitor for potential side‐effects that have been associ­ ated with their use. In the study with the largest number of cats, 18% had side‐effects associated with methima­ zole; a more recent study revealed that 44% of 39 cats had side‐effects. In 44 cats receiving carbimazole for one year, 44% had associated side‐effects, with gastroin­ testinal signs (decreased appetite, vomiting, diarrhea) being most common. In another study, 13% of 39 cats treated with carbimazole experienced side‐effects. It is difficult to determine what percentage of side‐effects are caused by the drug versus something else such as concurrent disease. Most adverse reactions occur within the first few weeks to months after beginning therapy and include depression, inappetence, vomiting, and self‐induced excoriations of the head and neck (facial pruritus). Gastrointestinal signs are less common with transder­ mal administration of methimazole. Mild to serious hematologic complications, including agranulocytosis and thrombocytopenia either alone or concurrently, and more rarely immune‐mediated hemolytic anemia may also occur. Hepatic toxicity with marked increases in bilirubin concentration and hepatic enzyme activi­ ties has been described in less than 2% of cats treated with methimazole. Cessation of therapy is required if either serious hematologic or hepatic reactions develop. Serum antinuclear antibodies develop in approximately 50% of cats treated with methimazole for longer than six months, usually in cats on high‐dose therapy (>15 mg/ day). Although clinical signs of a lupus‐like syndrome have not been reported, decreasing the daily dosage is recommended. Nutritional Management Production of thyroid hormone requires uptake by the thyroid gland of sufficient amounts of iodine, which is provided by dietary intake. The only function for ingested iodine is for thyroid hormone synthesis. This observa­ tion led to the hypothesis that limiting dietary iodine intake could be used to control thyroid hormone pro­ duction and potentially manage hyperthyroidism in cats. After more than a decade of research and development, a limited‐iodine therapeutic food (Hill’s® Prescription Diet® y/dTM Feline) containing 90% of the adrenal cortex. Clinical signs are due to both cortisol and aldosterone deficiency in the majority of patients, although some patients only have signs of cortisol deficiency (atypical hypoadrenocorticism). Aldosterone is the major mineralocorticoid secreted by the outermost layer of the adrenal cortex, the zona glomerulosa. Its main activities are the conservation of sodium and water, and excretion of potassium and hydrogen ions (acid), from the distal renal tubules. Aldosterone secretion is primarily regulated by the renin‐angiotensin‐ aldosterone system, but secretion can also be stimulated by hyperkalemia. In patients with hypoadrenocorticism and subsequent aldosterone deficiency, hyperkalemia, hyponatremia, and hypovolemia are common. Cortisol is the major glucocorticoid produced by the innermost layers of the adrenal cortex, the zonae fasciculata and reticularis. Its functions include stimulation of gluconeogenesis and erythropoiesis, suppression of the inflammatory response, and maintenance of gastrointestinal mucosal integrity. Cortisol also helps the body deal with stress. In dogs with HOAC, cortisol deficiency often results in lethargy, gastrointestinal signs, hypoglycemia, and anemia. Cortisol release is regulated by the hypothalamic‐ pituitary‐adrenal axis (HPAA). During times of stress (emotional, physiologic, other), corticotropin‐releasing hormone (CRH) is secreted from the hypothalamus, and stimulates the release of adrenocorticotropic ­hormone (ACTH) from the pituitary gland. ACTH then stimulates cortisol secretion from the adrenal cortex. Cortisol has a negative feedback effect on both the

­ ituitary and hypothalamus, such that ACTH and CRH p production are decreased when adequate cortisol concentrations are present. Most naturally occurring cases of primary hypoadrenocorticism in dogs are idiopathic, likely due to immune‐ mediated destruction of the adrenal cortex. Rarely, infiltration of the adrenal cortex by fungus, neoplasia, other granulomatous disease, or amyloidosis has been reported. Trauma, hemorrhage, and infarction may also lead to Addison disease, as can drugs used to treat hyperadrenocorticism. Although proper use of mitotane usually leads to necrosis of only the zonae fasciculata and reticularis, improper monitoring, or use in a particularly sensitive patient, can result in destruction of all three layers of the cortex, causing both cortisol and aldosterone deficiency. Trilostane is an enzyme inhibitor that decreases cortisol concentrations, but also decreases aldosterone concentrations to a lesser extent. It can lead to cortisol deficiency and, occasionally, aldosterone deficiency. Additionally, trilostane has been reported to cause idiosyncratic adrenocortical necrosis, resulting in both cortisol and aldosterone deficiency. Secondary hypoadrenocorticism can be caused by either a pituitary lesion, resulting in ACTH deficiency, or iatrogenically. Iatrogenic secondary hypoadrenocorticism is caused by abrupt cessation of exogenous glucocorticoids following long‐term use. Long‐term use of glucocorticoids suppresses production of ACTH. Without the trophic effects of ACTH, the zonae fasciculata and reticularis atrophy. When the exogenous glucocorticoid is abruptly discontinued (not tapered), ACTH production occurs, but the adrenal cortex is atrophied and incapable of producing adequate amounts of cortisol. Thus, signs of cortisol deficiency occur. Aldosterone deficiency is not present in cases of secondary hypoadrenocorticism, since ACTH has little regulatory control of aldosterone production.

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Section 2  Endocrine Disease

Epidemiology Hypoadrenocorticism affects approximately 1/2000 dogs. Dogs with other immune‐mediated endocrinopathies, such as hypothyroidism and diabetes mellitus, are at increased risk. Signalment

Young to middle‐aged female dogs are predisposed to Addison disease, although dogs at any age may be affected, with some studies revealing a more equal distribution between sexes. Poodles of all sizes, West Highland white terriers, Great Danes, bearded collies, Portuguese water dogs, Leonbergers, and Nova Scotia duck‐tolling retrievers (NSDTRs) are predisposed. Notably, NSDTRs may develop clinical signs at a very young age (as young as 2 months). Heritability has been proven in standard poodles, bearded collies, and NSDTRs. History and Clinical Signs The clinical manifestation of hypoadrenocorticism is highly variable in presenting complaint, chronicity, and severity. Whereas some dogs present with chronic signs, others present acutely, in an “Addisonian crisis.” The pathophysiology for chronic and acute hypoadrenocorticism is the same, and these presentations represent a continuum of disease progression. If not diagnosed and treated early in the course of disease, dogs with chronic signs may present in crisis. Addison’s disease often causes vague, nonspecific clinical signs that can be confused with other diseases, thus its nickname “The Great Pretender.” Nonspecific signs such as lethargy, inappetence, and weight loss occur in most patients. A waxing and waning pattern,  with exacerbations following stressful events or improvement following fluid or steroid administration, is often noted. Gastrointestinal signs, including vomiting and diarrhea, are common and likely due to the loss of “trophic” effects of cortisol on the gastrointestinal mucosa. Melena and/or hematochezia, and abdominal pain, may be reported by the owners or found on rectal examination. Exacerbation or onset of gastrointestinal (GI) signs following a stressful event is often reported in dogs later diagnosed with Addison disease, so HOAC should be considered in patients with recurrent “stress colitis,” particularly if accompanied by other GI signs, lethargy, or weakness. Renal loss of sodium and water, and the resulting decreased renal medullary concentration gradient due to hyponatremia, sometimes lead to polyuria and polydipsia. Rarely, seizures due to hypoglycemia occur, and HOAC should be ruled out in all dogs with unexplained hypoglycemia.

Less common clinical signs include generalized or hindlimb muscle weakness, megaesophagus (ME), and muscle cramping. The etiology of these abnormalities is unclear, but is probably due to aberrant neuromuscular function caused by deranged electrolyte concentrations and/or hypocortisolemia. Muscle weakness can be severe, and the patient may present for inability to rise. Generalized debility could explain the weakness, but some of these dogs are specifically weaker in the hindlimbs. Regurgitation due to ME is a rare presentation for hypoadrenocorticism, but since the ME usually resolves with appropriate treatment, HOAC should be ruled out in all cases of ME. Hair loss or a change in hair coat color are observed occasionally in dogs with hypoadrenocorticism, but the cause is unknown. Hypothyroidism should be ruled out in these patients following initial management of HOAC. Approximately 30% of dogs with Addison disease present in hypovolemic shock, or Addisonian crisis. In addition to the aforementioned clinical signs, especially vomiting, diarrhea, and GI bleeding, these dogs sometimes experience collapse and/or severe generalized weakness. Classic signs of hypovolemic shock are usually present, including weak pulse, pale mucous membranes, and prolonged capillary refill time. Some dogs are also hypothermic. Heart rate is variable. Whereas most hypovolemic non‐Addisonian dogs are tachycardic (>160 bpm), patients in an Addisonian crisis often have a normal to decreased heart rate. This is due to the effects of hyperkalemia on cardiac conduction. The presence of a decreased or normal heart rate in a patient in hypovolemic shock (“relative bradycardia”) should raise suspicion of hyperkalemia and hypoadrenocorticism. In patients with cardiac changes associated with hyperkalemia, rapid treatment and correction are critical for the survival of the patient. Gastrointestinal bleeding, including melena and hematochezia, is frequently seen during crisis. It may be ­present initially or appear within 2–3 days of presentation. Progressively decreasing hematocrit during treatment should increase suspicion of melena, and the possibility of GI bleeding should not be excluded despite the absence of melena or hematochezia on the initial examination. Hospitalization is recommended until the hematocrit stabilizes or increases. GI blood loss is occasionally severe enough to require blood transfusion. Diagnosis Since dogs with HOAC often present with nonspecific signs, most of the diagnostics are performed early in the work‐up, prior to significant suspicion of hypoadrenocorticism. A complete blood count, serum biochemistry analysis, and urinalysis should be performed in each

11  Hypoadrenocorticism in Dogs and Cats

patient with clinical signs consistent with hypoadrenocorticism. Electrolyte analysis must be included in the minimum database, as sodium and potassium abnormalities are often the first specific indicators of HOAC, and electrolyte disturbances are common in patients with gastrointestinal signs of any etiology, and need to be addressed during treatment. Serum Biochemistry, Urinalysis, and Complete Blood Count

Most dogs with HOAC are hyperkalemic and hyponatremic at diagnosis. Dogs with “atypical” hypoadrenocorticism are not (see Atypical Hypoadrenocorticism later). Potassium concentrations usually remain below 8 mEq/L, but may be as high as 11 mEq/L. Sodium concentrations are usually 120–140 mEq/L, but may be as low as 100 mEq/L. Some dogs present with one abnormality without the other (e.g., hyponatremia without hyperkalemia). Hypochloremia follows hyponatremia in about half the patients. Metabolic acidosis is also common, due to the inability to excrete H+ ions from the distal tubule. Electrolyte abnormalities are not always detected early in the course of the disease, and may appear as disease progresses. Emphasis is sometimes placed on calculation of the Na+/K+ ratio. The lower the sodium and the higher the potassium concentration, the lower the ratio. Although lower ratios make HOAC more likely, a high ratio does not rule it out, and dogs with low ratios do not always have HOAC. Therefore, the utility of the ratio is questionable. Any dog with hyponatremia and/or hyperkalemia should be considered a suspect for HOAC, regardless of the ratio. Most patients experience some degree of azotemia, with increased BUN (90% of dogs) and creatinine (65%). This is due to a combination of decreased renal perfusion from hypovolemia, and GI blood loss (increased BUN). Phosphorus is usually increased, as well. Although the azotemia is primarily pre-renal in nature, most dogs with HOAC have a urine specific gravity 50% suppression of the pretest UACR. This test has its limitations, including the availability of urine aldosterone measurement, but may be useful in PHA suspects with PAC within reference range. Therapy Initial treatment of PHA is aimed at controlling hypertension and hypokalemia and medical stabilization is recommended prior to adrenalectomy for adrenal tumors. Spironolactone is a competitive aldosterone receptor antagonist that helps control both hypertension and hypokalemia. The starting dose is 2 mg/kg q12h but this dose may be increased to 4 mg/kg q12h. Spironolactone has been reported to cause facial dermatitis in Maine Coon cats treated for hypertrophic cardiomyopathy, but this has not been reported in cats with PHA. Amlodipine, a calcium channel blocker, is given at 0.625–1.25 mg/cat q12–24h, to help control hypertension. Potassium gluconate is administered at a dose of 2–6 mEq PO q12h to alleviate hypokalemia. While adjusting medication doses, blood pressure and potassium concentration should be measured weekly. Myopathy is expected to resolve, although normokalemia may not be achieved. Hypertension usually resolves with appropriate therapy, but the vision loss is almost always permanent. Patients with idiopathic adrenal hyperplasia should be treated medically. Following medical stabilization, adrenalectomy is ideal for patients with unilateral adrenocortical neoplasia. However, surgery is associated with a high mortality rate (30% in one study), particularly if the vena cava is affected. Additionally, some owners have financial limitations that prohibit surgery, and some cats have concurrent conditions or metastasis. If the owner elects not to pursue surgical therapy, medical management can be pursued long term. The most common complication associated with adrenalectomy is hemorrhage; in one study, 4/10 cases experienced intraoperative or perioperative hemorrhage. One patient was stabilized with autotransfusion and another laparotomy, but the other three patients were euthanized following failed attempts at surgical hemostasis. Postsurgical monitoring includes blood pressure and  potassium monitoring, at least daily for 2–3 days. Amlodipine and spironolactone are discontinued postoperatively, and potassium supplementation is tapered until the potassium concentration normalizes.

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Prognosis Cats with unilateral adrenocortical neoplasia that ­survive the perioperative period have good long‐term survival (up to 1803 days); patients with adenocarcinomas appear to have the same prognosis as cats with

adenomas. Medically managed cats have reported survival times from several months to years. The prognosis for cats with adrenocortical hyperplasia is not well defined, but  most cats eventually succumb to chronic kidney disease.

­Further Reading Djajadiningrat‐Laanen S, Galac S, Kooistra H. Primary hyperaldosteronism: expanding the diagnostic net. J Feline Med Surg 2011; 13: 641–50. Gunn‐Moore D, Simpson K. Hypoadrenocorticism in cats. In: Rand J, ed. Clinical Endocrinology of

Companion Animals. Ames, IA: Wiley‐Blackwell, 2013, pp. 22–7. Lathan P. Hypoadrenocorticism in dogs. In: Rand J, ed. Clinical Endocrinology of Companion Animals. Ames, IA: Wiley‐Blackwell, 2013, pp. 1–21.

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12 Diabetes Mellitus in Dogs and Cats Jacquie S. Rand, BVsc, DVSc, MANZVS, DACVIM (SAIM) School of Veterinary Science, University of Queensland, Gatton, Queensland, Australia; Australian Pet Welfare Foundation, Kenmore, Queensland, Australia

­Etiology/Pathophysiology Glucose is the primary source of energy in the body. Blood glucose concentrations are tightly controlled by a feedback system of endocrine hormones. Insulin is produced by the pancreatic beta‐cells in response to increased blood glucose, and promotes storage of glucose, fatty acids, and amino acids as glycogen, fat, and protein. Insulin promotes uptake of glucose into most body cells for use, promotes storage of glucose as ­glycogen in liver and muscle, and inhibits gluconeogenesis. Cortisol, glucagon, epinephrine, and growth hormone oppose the actions of insulin, promoting catabolism of stored carbohydrate into glucose, stored triglycerides into free fatty acids and ketone bodies, and stored protein into amino acids. Because they oppose insulin’s effects, these four hormones are called “counterregulatory hormones” (Table 12.1). Diabetes mellitus is a disease characterized by hyperglycemia and associated clinical signs. It is classified into four broad types in human and veterinary medicine. ●●

Type 1 diabetes is a result of immune‐mediated beta‐ cell destruction, usually leading to an absolute insulin deficiency. The majority of neutered diabetic dogs have type 1 diabetes, but this form appears to be very rare in cats. The etiology of type 1 diabetes in dogs is multifactorial, and likely involves genetic factors and poorly understood environmental factors, which trigger beta‐ cell injury and inflammation. In type 1 diabetes, following nonspecific beta‐cell injury, autoimmunity then propagates continued destruction of beta‐cells, preventing regeneration after injury. Factors which trigger the gastrointestinal immune system are implicated in human type 1 diabetes, and in dogs, pancreatitis is a potential triggering factor. As in humans, there is a seasonal influence, with the incidence peaking in winter. Environmental factors such as chronic insulin r­ esistance

●●

●●

●●

secondary to glucocorticoid administration or obesity, and diseases which antagonize insulin’s actions would be expected to hasten onset of clinical signs when superimposed on a reduced capacity to secrete insulin as a result of immune‐mediated beta‐cell destruction. Type 2 diabetes is characterized by insulin resistance with concomitant beta-cell dysfunction. In developed countries, the majority of feline diabetics are type 2. Insulin resistance is multifactorial and associated with genetic factors, obesity, physical inactivity, male gender, and glucocorticoid steroids. Beta‐cell destruction is initiated and propagated by a variety of factors, not yet fully understood, that lead to a decline in insulin secretory capacity. When the increased demand to produce insulin as a result of insulin resistance is superimposed on beta‐cell dysfunction, chronic hyperglycemia ensues. Chronic hyperglycemia is toxic to beta‐cells and in turn causes injury and death of the beta‐cells, reducing insulin production even further. “Other specific types of diabetes” includes most other forms in companion animals, including drug‐induced diabetes (usually steroid use), endocrinopathies that antagonize insulin action (acromegaly, hyperadrenocorticism), or exocrine pancreatic disease. Chronic pancreatitis is the most common cause of canine diabetes in this category, accounting for approximately 30% of cases. Approximately 25–30% of poorly controlled diabetic cats have acromegaly. Gestational diabetes is not a significant cause of diabetes in companion animals in the US, as most companion animals are desexed. Pregnancy and diestrus are associated with increased progesterone concentrations, which uniquely in the bitch stimulate the mammary glands to produce growth hormone. Both hormones oppose insulin’s glucose lowering effects, leading to periodic insulin resistance in intact female dogs, and diestrus‐associated diabetes. Concurrent obesity would be expected to exacerbate this insulin resistance.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 12.1  Effects of insulin and counterregulatory hormones on carbohydrate metabolism Hormone

Cortisol

Effect on carbohydrates

Increases blood glucose ↓ Glucose uptake by cells ●● ↑ Gluconeogenesis ●● ↑ Glycogenolysis ●● Inhibits insulin ●●

Glucagon

Increases blood glucose ↑ Gluconeogenesis ●● ↑ Glycogenolysis ●● Inhibits insulin ●●

Epinephrine

Increases blood glucose ↑ Gluconeogenesis ●● ↑ Glycogenolysis

­History and Clinical Signs Diabetes is characterized by hyperglycemia resulting from insufficient insulin production or function. Hyperglycemia causes the hallmark clinical signs of polyuria, polydipsia, polyphagia, and weight loss that  distinguish diabetes. Some dogs and up to 50% of cats present with decreased appetite. Other clinical signs include hepatomegaly, lethargy, cataract for­ mation (dogs), and diabetic neuropathy (mainly cats). Signs are usually slowly progressive over weeks to months. Polyuria and Polydipsia

●●

Growth hormone

Insulin

Increases blood glucose ●● ↓ Glucose uptake by cells ●● ↑ Gluconeogenesis Decreases blood glucose ↑ Glucose uptake by all cells except liver, brain, red blood cells ●● ↑ Glycogen formation ●● ↓ Gluconeogenesis, glycogenolysis ●●

­Epidemiology Diabetes mellitus is the most common disorder of the endocrine pancreas in dogs and cats. The incidence in dogs is approximately 0.13%, and the i­ncidence in cats is approximately 0.5%, depending on the study population, and is influenced by geographical location and type of veterinary practice (referral or primary accession).

­Signalment Diabetes mellitus is a disease of middle‐aged to older companion animals, with peak prevalence of 7–12 years of age in dogs and 10–13 years in cats. Intact female dogs and male cats are predisposed. Various breeds of cat and dog are overrepresented, and predisposed breeds vary with geographic area. For example, in the USA, dog breeds at increased risk include miniature schnauzer, Samoyed, and miniature poodle, whereas in the UK, in addition to miniature schnauzer and Samoyed, breeds more commonly seen include Tibetan, cairn, border and Yorkshire terriers and Labrador retrievers. In cats, Burmese are overrepresented in Australia, New Zealand and the UK, and in the USA, Maine Coon, domestic longhair, Russian blue and Siamese. Norwegian forest cats and Burmese are at increased risk in Scandinavia.

Relative or absolute insulin deficiency in diabetes causes reduced blood glucose uptake by liver, muscle, and adipose tissues, and unchecked glucose production in the liver, which result in hyperglycemia. In the glomerulus of the kidney, nearly all the blood glucose is passively filtered into the ultrafiltrate. In the proximal tubule of the kidney, receptors are capable of resorbing only a fixed amount of glucose from the ultrafiltrate. This fixed amount is called the renal threshold, and is approximately 200 mg/dL in dogs and 250–290 mg/dL in cats. When the amount of glucose in the blood exceeds this renal threshold, glucose remains in the ultrafiltrate, ­acting as an osmotic diuretic and causing polyuria and compensatory polydipsia. Polyphagia Animals with uncomplicated diabetes mellitus may have an increased appetite. This is governed by central mechanisms, including increased hunger triggered by nutrient loss (predominantly glucose), and lack of insulin‐mediated uptake of glucose into cells of the satiety center, which normally inhibits hunger. Weight Change Obesity is a common co‐morbidity in middle‐aged and senior diabetic dogs and cats. It reduces insulin sensitivity, and when superimposed on beta‐cell loss from any cause, will hasten onset of hyperglycemia and clinical signs. However, untreated diabetes mellitus leads to weight loss, due to malassimilation of nutrients absorbed from the gut, and urinary loss of glucose and amino acids. Dogs in which pancreatitis is either a complicating co‐morbidity or a cause of their diabetes may later progress to exocrine pancreatic insufficiency (EPI), which exacerbates weight loss, and may be first evident as increased frequency of defecation.

12  Diabetes Mellitus in Dogs and Cats

Hepatomegaly

sorbitol occurs more rapidly than its degradation into fructose. In the diabetic eye, a large amount of sor­bitol is produced and acts as an osmotic agent within the lens fibers, drawing water into the lens, leading to lens swelling, lens fiber collapse, and ultimately cataract formation. Lens‐induced uveitis is common in diabetic dogs due to the rapid onset of the cataracts. Lens‐induced uveitis occurs when the lens becomes swollen and the stretched lens capsule allows leakage of cataract lens proteins into the anterior and posterior chambers. This, in turn, causes iris and ciliary body vasculitis, resulting in serum protein leakage into the aqueous humor and prostaglandin release from the iris, which continue the uveitis. Diabetic cataracts are found in 80% of diabetic dogs within 16 months (Figure 12.1). Diabetic cats are rarely clinical for cataracts, though recent studies suggest that mild diabetes‐associated lens opacity is more common than previously thought. Topical administration of

Insulin promotes storage of lipid in adipose tissue. Relative or absolute lack of insulin promotes lipid catabolism into fatty acids, which are transported to the liver. The liver uses this excess of fatty acids to produce and store triglycerides (lipids), leading to hepatic lipidosis and hepatomegaly. In the diabetic liver, excess fatty acids are also used for gluconeogenesis, which worsens hyperglycemia, and are converted into ketone bodies, which cause ketoacidosis. Cataracts The pathogenesis of cataracts in diabetics is multifac­torial and incompletely understood. The enzyme aldose reductase catalyzes the conversion of glucose to sorbitol, and the enzyme sorbitol dehydrogenase catalyzes the conversion of sorbitol to fructose. In the eye, the formation of (a)

(b)

(c)

Figure 12.1  Diabetic cataracts in a dog. (a) An 11‐year‐old cross‐bred dog photographed shortly after diagnosis of diabetes mellitus. (b) The same dog three months later. Diabetic cataracts had developed rapidly and the owners reported sudden vision loss. (c) The same dog following phacoemulsification surgery to remove the cataract from the right eye. Source: Fleeman and Rand 2000. Reproduced with permission.

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Figure 12.2  Cat with plantigrade stance. Source: Rand 2013. Reproduced with permission of John Wiley & Sons.

aldose reductase inhibitors three times daily retards progression of ocular changes. Diabetic Neuropathy The majority of diabetic dogs, cats, and people have microscopic evidence of nerve injury, including demye­lination, remyelination, and axonal degeneration. However, less than 50% of cats with diabetes mellitus have clinical neuropathy with hindlimb paraparesis, a plantigrade stance, and decreased ability to jump, as well as reduced reflexes and proprioceptive deficits (Figure  12.2). Weakness and muscle wasting are present in 50% of diabetic cats. Clinical diabetic neuropathy is rarely recognized in dogs.

­Diagnosis of Diabetes Mellitus Diagnosis of diabetes mellitus is based on demonstration of persistently increased blood glucose concentration, and is typically associated with glucosuria and classic clinical signs of polyuria and polydipsia. In dogs, blood glucose concentrations above the renal threshold of 200  mg/dL (11  mmol/L) are considered diabetic. In humans and cats, blood glucose concentrations persistently above normal but below that considered diabetic (>117 to < 180 mg/dL; >6.5 to 400 μmol/L; reference range varies by laboratory) or glycated hemoglobin (A1c) (reference range depends on assay used) in symptomatic pets with hyperglycemia and glucosuria helps confirm the diagnosis of diabetes mellitus. However, fructosamine is not consistently increased above the reference range in cats until blood glucose is approximately >300 mg/dL (>17 mmol/L), making it unsuitable for differentiating stress hyperglycemia from diabetes when blood glucose is 300 mg/dL (17 mmol/L) or less. In contrast, unpublished data suggest the new Baycom A1c is more sensitive at distinguishing transient hyperglycemia secondary to stress from persistent hyperglycemia of diabetes or prediabetes. Fructosamine is formed when blood glucose ­irreversibly binds plasma proteins (mainly albumin). Fructosamine concentration is roughly proportional to the average blood glucose concentration over the preceding 7–14 days, which is the half‐life of albumin in dogs and cats. Therefore, while a single blood glucose measurement reflects blood glucose at one moment in time, fructosamine is used as an indicator of blood glucose over a longer period of time and is used to document persistent hyperglycemia. Similarly, glycated hemoglobin is formed when blood glucose irreversibly binds hemoglobin (especially hemoglobin A1), and its concentration reflects the average blood g­lucose concentration over approximately 70 days (erythrocyte lifespan). A new test (Baycom A1c) validated for cats is available, and may be useful for differentiating cats with blood glucose in the normal range from those with a persistent mild hyperglycemia, for example prediabetic cats with  glucose concentrations of 130–180 mg/dL (7.2–9.9 mmol/L) or cats with subclinical diabetes (>180 mg/dL up to the renal thres­ hold of 250–290 mg/dL; >10 mmmo/L to 14–16 mmol/L). In cats, stress hyperglycemia may be confused with diabetic hyperglycemia on initial presentation. Stress hyperglycemia is usually (but not always) lower than ­diabetic hyperglycemia, with median blood glucose in stressed cats around 305 mg/dL (16.9 mmol/L) with a wide range (126–505 mg/dL; 7–28.1 mmol/L). Median blood glucose in newly diagnosed diabetic cats is around 475 mg/dL (26.4 mmol/L) with a wide range of 180– 1100  mg/dL (10–61.  mmol/L) that overlaps with the range of stress hyperglycemia. Diabetic hyperglycemia and glucosuria will be persistent over time, and will always be accompanied by typical clinical signs if blood glucose is above the renal threshold. In contrast, stress hyperglycemia typically resolves in 3–4 hours but may take 24 hours in a very few cats. Therefore, in most cats, an easy way to rule out persistent hyperglycemia of diabetes or prediabetes, is to test a repeat sample from the ear after the cat has been sitting quietly in a cage for 4 hours; in the majority of cats with stress hyperglycemia, blood glucose will be 360 mg/dL; 20 mmol/L) for at least 3–5 days. However, a normal fructosamine concentration does not rule out prediabetes or diabetes associated with more moderate hyperglycemia, whereas Hb1c appears more sensitive. Home blood glucose monitoring with a meter calibrated for feline blood may also assist in identifying prediabetic cats with persistent mild elevation of glucose (>117–162 mg/dL; 6.5–9 mmol/L). A typical work‐up for dogs or cats with suspected ­diabetes mellitus will include a complete blood count, chemistry profile with electrolytes, urinalysis, and urine culture via sterile collection. Some veterinarians routinely use fructosamine or, less commonly, glycated hemoglobin to document persistent hyperglycemia. In  dogs, if ­hypothyroidism or hyperadrenocorticism is suspected, it is best tested after diabetes is regulated, because of the effect of poorly regulated diabetes on interpretation of tests. In intact female dogs, measurement of serum progesterone may also be helpful; ­elevated progesterone levels in diestrus or pregnancy increase secretion of growth hormone, a counterregulatory hormone, leading to insulin resistance. In cats, a total T4 and feline leukemia virus/feline immunodeficiency virus (FeLV/FIV) test is warranted. Middle‐aged to older cats with polyuria/polydipsia, polyphagia, and weight loss should be screened for diabetes mellitus and hyperthyroidism, as the diseases cause similar clinical signs, and may occur concurrently. For patients with atypical or additional clinical signs, consider a thyroid panel (total or free T4 and thyroid‐stimulating hormone), insulin like growth factor (IGF-1), t­ rypsinogen‐like immunoreactivity (TLI), pancreatic lipase immunoassay (PLI), cobalamin and folate, adrenal function testing, thoracic and abdominal radiographs, and abdominal ultrasound as indicated based on the patient’s history and clinical exam findings. Cats with diabetes and blood glucose above the renal threshold have weight loss, and cats with a stable weight or weight gain should be tested for acromegaly. If IGF-1 is in the normal range, but clinical signs are consistent with acromegaly, test again after 6 weeks of insulin therapy.

­Therapy Nearly all veterinary patients with diabetes mellitus are managed with a combination of insulin therapy, diet, and weight management. Human and veterinary insulins are categorized both by onset and duration of action, and by protein source. Based on onset and duration of action, insulins are short acting, intermediate acting, and long acting.

Short‐acting insulins are generally used in hospital for intensive blood glucose control, while intermediate‐ and long‐acting insulins are used both in hospital and at home for long‐term, daily glycemic control. Based on protein source, insulins are human, human analog (synthetic insulin that is altered from the natural form but retains biological function), bovine, or porcine. Porcine and canine insulin have an identical amino acid sequence, while bovine and feline insulin differ by just one amino acid. Traditionally, veterinarians have managed canine and feline diabetes with a variety of veterinary or human insulins (Table 12.2). Veterinary insulins include porcine lente (Vetsulin/Caninsulin®, Merck), and protamine zinc human recombinant insulin (ProZinc®, Boehringer Ingelheim Vetmedica). Human insulins or insulin analogs include human‐origin lente (no longer available), neutral protamine Hagedorn (isophane, NPH, various manufacturers), glargine (Lantus®, Sanofi‐Aventis), detemir (Levemir®, Novo Nordisk), glargine 300 U/mL (Toujeo, Sanofi-Aventis) and degludec (Tresiba®, Novo Nordisk). Premixed combinations of short‐acting lispro (Eli Lilly) or aspart (Novo Nordisk) insulin with longer‐ acting protamine insulin are also available as intermediate‐acting insulins (Humalog® Mix 75/25 and Novolog® 70/30, respectively). The goals of treatment in canine diabetes include resolution of polyuria, polydipsia and abnormal body condition, while avoiding clinical hypoglycemia, diabetic ketoacidosis, urinary tract infections, and other complications through excellent glycemic control. In dogs, American Animal Hospital Association guidelines recommend the use of porcine lente, PZI, or NPH insulins, and a complete and balanced diet that lacks simple carbohydrates. A stable routine for meals, snacks, and exercise is crucial for good glycemic control; frequent changes in type of food or amount fed should be avoided (Box 12.1). The best diet for diabetic dogs is yet to be determined. Diets recommended include high‐fiber/restricted‐fat diets, which as a consequence are high in carbohydrate (>45%), and diets with moderate fiber and restricted carbohydrate (45% ME. If used with NPH insulin, these high‐fiber diets may exacerbate postprandial hyperglycemia in some dogs, because peak gastrointestinal glucose absorption occurs after peak insulin action, and in these cases are better fed 1–2 hours after insulin administration. High‐fiber/moderate‐fat diets may cause

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Table 12.2  Insulin types used in companion animal medicine and their characteristics Brand name

Product

Manufacturer

Concentration

Origin

Size

Humulin N

NPH

Eli Lilly

U‐100

Human

10 mL vial 3 mL pen

Vetsulin/ Caninsulin

Lente

Merck

U‐40

Porcine

10 mL vial 2. mL pen

Humalog Mix 75/25 and 50/50

Lispro (rapid acting) plus lispro protamine (intermediate)

Eli Lilly

U‐100

Recombinant human

10 mL vial 3 mL pen

Novolog Mix 70/30; 50/50

Aspart (rapid acting) plus aspart protamine (intermediate)

Novo Nordisk

U‐100

Recombinant human

10 mL vial 3 mL pen

ProZinc

PZI (protamine zinc)

Boehringer Ingelheim

U‐40

Recombinant human

10 mL vial

Lantus

Glargine

Sanofi‐Aventis

U‐100

Recombinant human analog

10 mL vial 3 mL pen

Toujeo

Glargine

Sanofi‐Aventis

U‐300

Recombinant human analog

1.5 mL pen

Detemir

Levemir

Novo Nordisk

U‐100

Recombinant human analogue

10 mL vial 3 mL pen

Tresiba

Degludec

Novo Nordisk

U‐100, U‐200

Recombinant human analog

10 mL vial 3 mL pen

Intermediate acting

Long acting

Box 12.1  Dosing protocol for dogs on lente, NPH, glargine or detemir ●●

●●

●●

●●

Starting dose is 0.5 U/kg administred BID for NPH, lente (Caninsulin/Vetsulin, Merck) and glargine (Lantus, Sanofi‐Aventis) insulin in dogs. Note starting dose for dogs on detemir (Levemir, Novo Nordisk) is 0.125 μ/kg BID. Dogs are on average four times more sensitive to detemir than other insulin. Home blood glucose monitoring is recommended using a meter calibrated for dog blood. Continuous or flash glucose monitoring systems are also very useful for assessing duration of action and adjusting insulin dose, but they measure interstitial blood glucose and are not calibrated for dogs. Their accuracy for determining actual nadir blood glucose concentration is poor. In the intitial stabilization phase, reassess dogs weekly. Feed a prescription diet formulated for diabetic dogs and keep amount fed and type of diet constant on a daily basis. Low carbohydrate is best when using lente, glargine or determir. Adjust amount of energy fed to achieve and maintain ideal body weight. If excessive weight loss occurs with a diabetic diet, swap to a premium maintenance diet with low carbohydrate and moderate fiber content. Increase insulin dose when the nadir (lowest) glucose concentration is >145 mg/dL (8 mmol/L) and when the blood glucose measurements obtained just prior to the morning and evening insulin injection are both ≥180 mg/dL (10 mmol/L).

●●

●●

●●

●●

●●

Do not change insulin dose when the nadir is 90–145 mg/ dL (5–8 mmol/L) and when the blood glucose measurements just prior to the morning and evening insulin injection are both ≥180 mg/dL (10 mmol/L). Decrease insulin dose either when the nadir is 2SD above the VHS of reference #1 (9.7 ± 0.5)

2. Jepsen‐Grant K, et al. Vet Radiol Ultrasound 2013;54:3–8

Yorkshire terrier (n = 30)

9.9 ± 0.6

VHS assessed on right lateral views − 13% of dogs had a VHS >2SD above the VHS of reference #1 (9.7 ± 0.5)

2. Jepsen‐Grant K, et al. Vet Radiol Ultrasound 2013;54:3–8

All breed dogs (n = 63) Right lateral recumbency Left lateral recumbency

9.8 ± 0.6 9.5 ± 0.8

VHS was significantly larger in right than in left lateral recumbency

3. Greco A, et al. Vet Radiol Ultrasound 2008;49:454–5

Beagle (n = 19) Right lateral recumbency Left lateral recumbency

10.5 ± 0.4 10.2 ± 0.4

VHS was significantly larger in right than in left lateral recumbency

4. Kraetschmer S, et al. J Small Anim Pract 2008;49:240–3

Boxer (n = 20)

11.6 ± 0.8

Right lateral thoracic radiographs

5. Lamb CR, et al. Vet Record 2001;148:707–11

Doberman (n = 20)

10.0 ± 0.6

Right lateral thoracic radiographs

5. Lamb CR, et al. Vet Record 2001;148:707–11

Cavalier King Charles (n = 20)

10.6 ± 0.5

Right lateral thoracic radiographs

5. Lamb CR, et al. Vet Record 2001;148:707–11

German Shepherd dog (n = 20)

9.7 ± 0.7

Right lateral thoracic radiographs

5. Lamb CR, et al. Vet Record 2001;148:707–11

Rottweiler (n = 38)

9.8 ± 0.1

Greyhound (n = 42)

10.5 ± 0.1

Comments

References

1. Buchanan JW, et al. J Am Vet Med Assoc 1995;206:194–9

6. Marin L, et al. Vet Radiol Ultrasound 2007;48:332–4 VHS was significantly higher in greyhounds than in rottweilers

Left‐Sided Congestive Heart Failure

In the case of left‐sided congestive heart failure, prominent pulmonary veins owing to venous congestion are seen, particularly in the hilar area (as they enter the left atrium) on lateral views. The pulmonary vein diameter appears greater than that of the corresponding arteries. This asymmetry is easiest to detect for cranial vessels on

6. Marin L, et al. Vet Radiol Ultrasound 2007;48:332–4

lateral views (where the pulmonary veins are ventral to the accompanying arteries), and for caudal vessels on the dorsoventral view (where the pulmonary veins are medial to the accompanying arteries). Radiographic signs of left‐sided congestive heart failure also include visualization of mild to severe ­ inter­stitial, peribronchial to alveolar pulmonary opacity

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Figure 16.6  Clock face analogy (a) applied to the canine cardiac silhouette (b) on the dorsoventral view. The aortic arch (AoA) approximately extends from 11 o’clock to 1 o’clock, the main pulmonary artery (MPA) from 1 o’clock to 2 o’clock, the left ventricle (LV) from 3 o’clock to 6 o’clock, and the right ventricle (RV) from 6–7 o’clock to 11 o’clock (with a superimposition with the RA between 9 and 11 o’clock). In cats and dogs, enlargement of the left auricular appendage (Laur) contributes to a deformation of the cardiac silhouette between 2 and 3 o’clock. In the cat, the body of the left atrium (LA) also extends from 2 to 3 o’clock, whereas in the dog the LA is rather superimposed over the caudal part of the cardiac silhouette (b). Source: Medical Imaging Unit, ENVA.

(b)

AoA

MPA RA

LAur

LA

RV

LV

(Figures  16.7c, 16.10a and 16.10b) due to pulmonary edema. Cardiogenic pulmonary edema is typically more pronounced in the dorsal perihilar area on lateral views in the dog, with a bilateral distribution on the dorsoventral view in both species. In some instances, it can be asymmetric in the dog, with the right lung fields more severely affected than the left. In the case of alveolar edema, the alveolar spaces are filled with fluid. The margins of the pulmonary vessels are therefore totally obscured and air bronchograms (corresponding to black air‐filled bronchi surrounded by white radiopaque lung tissue) are detected. Right Heart Diseases Right Atrial Enlargement

On lateral views, right atrial enlargement is characterized by: ●●

●●

enlargement of the dorsocranial part of the cardiac ­silhouette (Figure 16.11a) dorsal elevation of the trachea over the cranial portion of the heart (less commonly).

On the dorsoventral view, right atrial enlargement is characterized by bulging of the cardiac silhouette in the 9–11 o’clock position. Right Ventricular Enlargement

On lateral views, right ventricular enlargement is characterized (Figures 16.7c and 16.11a) by: ●●

●●

increased sternal contact, with widening and increased convexity of the cranioventral heart margins elevation of the apex from the sternum and, in the case of severe enlargement, dorsal elevation of the caudal vena cava as well.

On the dorsoventral view, right ventricular enlargement is characterized (Figure 16.11b) by: ●●

●●

broadening and rounding of the cardiac silhouette at the 6–7 to 9–11 o’clock position with a reverse‐D appearance a shift to the left of the cardiac apex in the case of severe enlargement.

Similar abnormal patterns are detected on the ventrodorsal view. Right‐Sided Congestive Heart Failure

Thoracic radiographic signs of right‐sided congestive heart failure include visualization of right heart enlargement associated with widening of the caudal vena cava (whose normal diameter is usually approximately that of the descending thoracic aorta) and interlobar pleural fissures secondary to pleural effusion (with lung collapse and rounded pulmonary borders in severe cases). Extrathoracic radiographic signs include hepatomegaly (extension of the liver caudally to the last rib with abnormally round margins and caudal displacement of the stomach) and ascites, which is characterized by abdominal distension and increased fluid opacity with loss of intraabdominal details. Specific Radiographic Signs Associated with Heart Diseases

The main specific radiographic signs associated with heart diseases are presented in Table  16.3 and in Figures 16.7 to 16.12.

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Table 16.3  Generalized or localized heart enlargement on thoracic radiographs: main cardiac causes, associated radiographic signs, differential diagnosis Type of heart enlargement

Main causes

Potential associated radiographic signs Differential diagnosis

Left heart enlargement

Left atrial and/ or ventricular enlargement

Congenital heart diseases: mitral dysplasia with mitral valve regurgitation, significant left‐to‐ right shunting due to VSD and PDA, subaortic stenosis and/or aortic insufficiency Acquired heart diseases: hypertrophic and restrictive cardiomyopathy (cat), degenerative mitral valve disease (dog), mitral or aortic valve endocarditis, dilated cardiomyopathy, cardiomyopathy secondary to systemic arterial hypertension and hyperthyroidism (cat)

Isolated left atrial enlargement

Congenital heart diseases: mitral valve stenosis

Enlargement of the aortic arch in the case of PDA, subaortic stenosis and systemic arterial hypertension Increased pulmonary artery prominence and increased pulmonary vessels (veins and arteries) in the case of VSD and PDA with significant left‐to‐right shunting Increased pulmonary veins in cases with left‐sided congestive heart failure (see text)

Pulmonary mass and/or adenopathy in the hilar area (left atrial enlargement)

Increased caudal vena cava on lateral views in cases of right‐sided congestive heart failure (see text) Enlargement of the main pulmonary artery (post‐stenotic dilation) with decreased pulmonary vasculature in the case of pulmonic stenosis Enlargement of the main pulmonary artery, abnormal pulmonary arteries (tortuous, larger than their accompanying veins) in the case of pulmonary hypertension Pulmonary infiltrates in the case of dirofilariosis, pulmonary embolism, respiratory diseases, etc.

Tracheobronchial adenopathy and cranial mediastinal mass (including heart base tumor)

Right heart enlargement

Right atrial and/ Congenital heart diseases: tricuspid dysplasia with tricuspid regurgitation, significant left‐to‐ or ventricular right shunting due to ASD, pulmonic stenosis enlargement and/or pulmonic insufficiency (rare), tetralogy of Fallot Acquired heart diseases: degenerative tricuspid valve disease, arrhythmogenic right ventricular cardiomyopathy, pulmonary arterial hypertension (main causes: dirofilariosis, angiostrongylosis, respiratory diseases, pulmonary embolism, reversed‐shunting congenital defects, etc.)

Isolated right atrial enlargement

Congenital heart disease: tricuspid stenosis Acquired heart disease: right atrial neoplasia (hemangiosarcoma)

Generalized heart enlargement

Severe left and/or right heart diseases Pericardial effusion, peritoneal‐pericardial hernia Intracardiac mass (tumor) “Athletic heart”

Globoid cardiac silhouette in the case of pericardial diseases, with gas‐ containing structures within the pericardial sac and loss of outline between the ventral diaphragmatic surface and the caudal ventral cardiac silhouette in the case of peritoneal‐ pericardial hernia

ASD, atrial septal defect; DV, dorsoventral; PDA, patent ductus arteriosus; VD, ventrodorsal; VSD, ventricular septal defect.

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(a) LA

Carina

(b)

(c) LB Trachea

Figure 16.7  Thoracic radiographs of three different dogs with compensated (a,b) and decompensated (c) degenerative mitral valve disease. (a) A marked left atrial enlargement (LA) is seen on this lateral radiograph. Note the tracheal and carina dorsal elevation as well as the straightening of the caudal part of the cardiac silhouette, suggesting left ventricular dilation as well. (b) On this dorsoventral view, enlargement of the left auricular appendage produces a bulge (large arrow) at the 2–3 o’clock position. Additionally, the dilated LA body (thin arrows) creates a “mass effect” in the dorsocaudal region of the cardiac silhouette. (c) This lateral radiograph shows diffuse cardiogenic pulmonary edema that is more severe in the caudodorsal perihilar area (arrow). Left heart enlargement is responsible for dorsal elevation of the intrathoracic trachea, carina, and left main bronchus (LB). Note also the abnormal increased sternal contact of the heart, suggesting right ventricular enlargement (which was further confirmed to result from pulmonary arterial hypertension). Source: Medical Imaging Unit, ENVA.

­Conventional Echocardiography Over the last 30 years, standard transthoracic echocardiography has become a major imaging tool for the diagnosis, management, and follow‐up of canine and feline heart diseases, allowing noninvasive assessment of heart morphology (ventricles, atria, cardiac valves, auricular appendages) and function, as well as examination of the anatomy of the proximal great vessels. General Principles and Technique Conventional echocardiography combines 2D and M‐ mode imaging. M‐mode echocardiography provides one‐dimensional views of the heart, while 2D echocardiography displays cross‐sectional or 2D cardiac images (in transverse and longitudinal planes) moving in real

time. M‐mode echocardiograms display motion over time of the various tissue interfaces crossed by the linear M‐mode ultrasound beam. Currently, the 2D mode is always used to guide accurate placement of the M‐mode cursor, which appears as a vertical line on 2D echocardiograms. M‐mode echocardiography has a much higher spatial and temporal resolution than 2D echocardiography, and therefore allows recording of the motion of subtle cardiac structures. Transthoracic 2D and M‐mode echocardiography can be performed on animals in lateral recumbency or in standing position, as the measurements obtained in both positions show similar variability. Some animals may not tolerate a recumbent position because of discomfort or dyspnea, making the standing position easier and more suitable. Placing the patient in lateral recumbency decreases lung interference but, on the other hand, may increase stress in nervous animals.

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(a) LB Trachea

(c)

1

(b)

2 3

ACDO

Figure 16.8  Thoracic radiographs of two dogs with patent ductus arteriosus (PDA). (a,b) Lateral radiographs of the same dog (German shepherd) respectively before and 24 hours after surgery (placement of an Amplatz canine duct occluder, ACDO). Note the marked generalized heart enlargement on (a), with dorsal elevation of the intrathoracic trachea, carina, and left main bronchus (LB), associated with vascular congestion (arrows) due to overcirculation of the lung field, and interstitial edema in caudal lungs. After ACDO placement (b), heart size (width and height) has decreased and vascular congestion disappeared. (c) Dorsoventral radiograph of a Cavalier King Charles spaniel with PDA before surgical closure. Note the elongated cardiac silhouette (double arrow), the heart base enlargement (including 1: descending aorta, 2: main pulmonary artery, 3: left auricular appendage), and the increased opacity caudally to the tracheal bifurcation, corresponding to the dilated left atrium (LA) superimposed over the caudal cardiac silhouette. Source: Medical Imaging Unit, ENVA.

When patients are echoed in a recumbent position, a special scanning table with a cutout is needed, and images are obtained from beneath the table through the table hole. The standing position does not require a specific scanning table, as the animal is gently restrained against an assistant to keep its position stable. For both animal positions, the transducer is placed on the thoracic wall in an intercostal space over the precordial impulse area, close to the ­sternum, to obtain parasternal views. However, subcostal (or subxiphoid) views may sometimes be used for Doppler assessment of the aortic outflow (as they may provide a better Doppler beam alignment than parasternal views). In most cases where manual restraint is sufficient, sedation is avoided as it can alter systolic function and may affect cardiac dimensions.

Most animals do not require clipping of hair over the area of contact on the thoracic wall (more than 90% of the patients at our unit are not clipped for conventional echocardiographic examinations or for advanced ultrasound techniques, such as tissue Doppler imaging). Acoustic gel helps to provide an optimal air‐free contact between the probe and the thoracic wall. High‐frequency transducers are characterized by low penetrating power but high resolution. Conversely, low‐ frequency transducers are characterized by high penetrating power but low resolution. In canine and feline cardiology, transducer frequencies are usually between 2.5 and 12 MHz. Transducer frequencies are commonly adjusted during a single echocardiographic examination, depending on the location and size of the cardiovascular structures tested. As most commercially

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(a)

right and left parasternal positions. Transverse views, also called short‐axis views, are obtained with the 2D ultrasound plane perpendicular to the long axis of the heart. Conversely, sagittal views, also called long‐axis views, are obtained from short‐axis views by rotating the transducer 90° counterclockwise, so that the 2D ultrasound plane becomes parallel to the long axis of the heart. Main Parasternal Short‐Axis Views

(b)

Figure 16.9  Thoracic radiographs of a cat with left‐sided congestive heart failure secondary to hypertrophic cardiomyopathy. Note on the lateral view the mild convexity of the caudal heart border and the increased cranial–caudal diameter associated with mild but diffuse interstitial pulmonary edema (a). On the dorsoventral view (b), the dilated left atrium (arrow) associated with the pointed appearance of the left apex (owing to concentric left ventricular hypertrophy) creates a “Valentine‐ shaped” heart silhouette. Source: Medical Imaging Unit, ENVA.

available transducers have a wide frequency bandwidth, frequencies can be changed while using the same probe. If possible, the ultrasound machine’s ECG should be connected to the patient during the echocardiographic examination, in order to detect potential arrhythmias and study the correlation between electrical and mechanical events. Standard 2D Echocardiographic Views and Measurements A variety of standardized 2D transverse and sagittal echocardiographic views have been defined from the

The three most commonly used 2D transverse echocardiographic views are the right parasternal transventricular short‐axis view (Figure  16.13), the right parasternal transmitral short‐axis view (Figure  16.14), and the right parasternal transaortic short‐axis view (Figure 16.15). The right parasternal transventricular short‐axis view (Figure  16.13a) provides transverse visualization of the two ventricular cavities, and is therefore used to assess left (Figures 16.13b and 16.13c) and right (Figure 16.13d) ventricular changes in size and shape. This view is also commonly used to obtain M‐mode echocardiograms at the ventricular level. The right parasternal transmitral short‐axis view (Figure  16.14) allows visualization of the mitral valve leaflets within the left ventricular cavity, and is therefore commonly used to obtain M‐mode echocardiograms at the mitral valve level. The right parasternal transaortic short‐axis view (Figure 16.15a) provides transverse visualization of the heart base, with the ascending aorta visible as a circle in the middle of the sector image. This view is commonly used to calculate the left atrium/aorta ratio, and to therefore allow detection of left atrial dilation (Figure 16.15b, Tables 16.4 and 16.5). This view is also used to evaluate the right ventricular outflow tract as well as the pulmonary cusps (Figure  16.15c). A slight rotation of the transducer provides a short‐axis view of the heart base optimized for the pulmonary arteries, which is useful for observing heartworms or blood clots within the pulmonary arteries (Figure  16.15d) and for detecting indirect signs of pulmonary arterial hypertension (Figure  16.15e). A similar view can be obtained from the left parasternal position. Main Parasternal Long‐Axis Views

Two 2D sagittal echocardiographic views can be obtained from the right side of the thorax  –  the right parasternal long‐axis four‐ and five‐chamber views (Figures  16.16 and 16.17, respectively). The right parasternal long‐axis four‐chamber view provides clear visualization of the atrial septum and the atrioventricular valves, and is therefore useful for the diagnosis of

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Figure 16.10  Thoracic radiographs of a Doberman with acute left‐sided congestive heart failure secondary to dilated cardiomyopathy. Note the tracheal and carina dorsal elevation as well as the straightening of the caudal part of the cardiac silhouette, suggesting left heart enlargement on the lateral view (a). Diffuse patchy interstitial and alveolar opacities consistent with cardiogenic pulmonary edema are also present. They are symmetrically distributed in the right and left lung fields on the dorsoventral view (b). Source: Medical Imaging Unit, ENVA.

(a)

atrial septal defects (Figure  16.16b) and mitral valve lesions (Figure 16.16c). The right parasternal long‐axis five‐chamber view is commonly used to examine the left ventricular outflow tract, the mitral valve leaflets, and the junction between the interventricular septum and the anterior aortic wall in order to respectively diagnose left ventricular outflow tract obstruction (Figure  16.17b), mitral valve lesions (Figure  16.17c), and ventricular septal defects (Figure 16.17d). Long‐axis four‐ and five‐chamber views can also be obtained from the left parasternal position (2D left apical or caudal four‐ and five‐chamber views; Figure 16.18). The left apical four‐chamber view provides good visualization of the ventricular inflow tracts and is therefore used to assess mitral and tricuspid diastolic inflows. It may also be used to calculate end‐diastolic and end‐systolic left ventricular volumes (Box 16.1). The end‐diastolic left ventricular length can also be measured from the left apical four‐chamber view and the index of sphericity can then be calculated (this index is defined as the end‐diastolic left ventricular length / left ventricular diameter assessed by M‐mode at end‐diastole; normal values are usually >1.65). The main use of the left apical five‐chamber view is for the Doppler assessment of systolic aortic outflow. Standard M‐Mode Views and Measurements The two most commonly used M‐mode echocardiograms are obtained at the ventricular level and the mitral valve level (Figure 16.19). The M‐mode echocardiogram obtained at the ventricular level is used to measure left ventricular end‐diastolic and end‐systolic diameters, left ventricular free wall thickness and interventricular

(b)

septal thickness at end‐diastole and at end‐systole (which correspond to the onset of qRs and end of T‐ wave, respectively, on concomitant ECG tracing), in order to detect and quantify left ventricular alterations (Figure  16.20a and 16.20b). The left ventricular fractional shortening can then be calculated to indirectly assess the transverse systolic myocardial function (Figure  16.19a; see also Tables  16.4 and 16.5). The M‐ mode echocardiogram obtained at the ventricular level can also be used to measure right ventricular dimensions (end‐diastolic diameter and end‐systolic myocardial wall thickness). E point to septal separation (up to 5–8 mm in the dog, depending on the size of the animal) can be measured from M‐mode echocardiograms obtained at the mitral valve level. This distance increases in the case of severely reduced systolic left ventricular function. The same M‐ mode view may be used to detect systolic anterior motion of the mitral valve, which is commonly associated with obstructive forms of feline hypertrophic cardiomyopathy (Figure 16.20c).

­Conventional Doppler Echocardiography Conventional Doppler echocardiography allows detection and analysis of blood flows within the cardiovas­cular system. The conventional Doppler examination consists of locating blood flows within the cardiac chambers and vessels, as well as assessing their direction, velocity, and duration. This technique is also used to differentiate normal laminar flows from abnormal nonlaminar (or turbulent) flows. Additionally, flow ­volumes and pressure

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mation, and has now supplanted invasive techniques such as cardiac catheterization for the diagnosis of most human and animal heart diseases. Advanced ultrasound imaging techniques beyond the scope of this chapter are summarized in Box 16.2.

(a)

Basic Principles

(b) (c)

Rib 9

Rib 9

Figure 16.11  Thoracic radiographs of two dogs with marked right heart enlargement resulting from severe pulmonic stenosis (a) and pulmonary arterial hypertension secondary to a large reversed ventricular septal defect (b,c). (a) Note on this lateral view the increased sternal contact of the cranial cardiac margin (small arrows) consistent with right ventricular enlargement, as well as protrusion of the dorsal cranial margin of the cardiac silhouette (large arrow) owing to both right atrial dilation and enlargement of the main pulmonary artery. The pulmonary vasculature is also slightly diminished. (b) On this dorsoventral view, right ventricular enlargement is characterized by a rounded right heart border bulging to the right (approximately between 6 and 9 o’clock; arrows) and a leftward shift of the cardiac apex, leading to a reverse‐D appearance. Note also the marked prominence of the main pulmonary artery (MPA) between 1 and 2 o’clock and the enlarged peripheral pulmonary arteries (PA), compared with the pulmonary veins (V, see details on (c)), which indicates pulmonary arterial hypertension. Width of peripheral pulmonary arteries should be similar to that of the corresponding pulmonary veins and should not be larger than the ninth rib diameter at the crossing point between the two. Source: Medical Imaging Unit, ENVA.

gradients across valves or between cardiac chambers can be estimated from velocity Doppler data using specific equations. Conventional Doppler echocardiography complements 2D and M‐mode echocardiography by providing accurate cardiovascular hemodynamic infor-

Conventional Doppler echocardiography is based on the Doppler principle, discovered by Christian Doppler in the 19th century, which states that a moving structure (e.g., blood cells) reflects the incident ultrasound beam back to the transducer with a higher or lower frequency than the original transmitted frequency ­ depending on whether the structure is, respectively, moving toward or away from the transducer. This difference between transmitted and received frequencies, also called the Doppler shift or frequency shift, is directly related to the velocity of the reflecting moving structures, which can thus be deduced from the frequency shift analysis. The recorded frequency shift (and therefore the calculated flow velocity) also depends on the cosine of the intercept angle (θ) between the ultrasound beam and blood flow path. Therefore, whichever Doppler mode is used, the incident ultrasound beam must be perfectly parallel to the blood flow path (θ = 0°), otherwise the blood flow velocity will not be assessed correctly, that is, it will be underestimated as compared to the actual velocity. As frequency shift values are usually within the hearing range of the human ear, an audible signal can be simultaneously associated with spectral Doppler tracings of flow velocities. Listening to this Doppler audio signal is particularly useful in practice to help in aligning the ultrasound beam parallel to blood flow: the louder the audible signal, the higher the Doppler shift, the smaller the intercept angle θ, the better the alignment between blood flow and the ultrasound beam. Standard Doppler Modes Standard Doppler echocardiography includes different complementary modes, spectral Doppler modes and color‐flow Doppler mode (see examples on Figures  16.21–16.23). Each mode has specific advantages and limitations, of which the observer must be perfectly aware in order to avoid misinterpretation of the recorded Doppler signals. Spectral Modes: Pulsed‐Wave and Continuous‐ Wave Doppler Modes

The two main spectral Doppler modes are pulsed‐wave (PW) and continuous‐wave (CW) Doppler modes.

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Figure 16.12  Lateral (a) and ventrodorsal (b) thoracic radiographs of a dog with a peritoneal‐pericardial diaphragmatic hernia. Note the marked enlargement and the globoid shape of the heart on both views, overlap of the diaphragmatic and caudal heart borders (arrow), associated with a small liver and gas‐filled intestinal loops (stars), that are superimposed on the cardiac silhouette because of their pericardial location. Source: Medical Imaging Unit, ENVA.

(a)

With these spectral Doppler modes, blood flow velocities (expressed in m/s) are displayed over time in a region of interest, that is, either all along a linear axis (or cursor line) for the CW Doppler mode or in a specific site, also called a sample gate or sample volume, for the PW Doppler mode. The position of the CW Doppler cursor as well as the location and size of the PW sample volume are carefully selected by the observer on 2D echocardiographic views. Whichever spectral Doppler mode is selected, the velocity profiles are displayed as positive (i.e., above the baseline) or negative (i.e., below the baseline), when blood cells respectively move toward or away from the transducer. The CW mode uses two transducer elements, one continuously emitting and the other continuously receiving ultrasound waves. Because this emission– reception process is continuous, there is no loss of blood flow information and therefore no maximum limit for velocity measurements. Velocities higher than 8 m/s can thus be measured. Conversely, the PW mode uses only one transducer element which discontinuously emits short bursts of ultrasound waves at a given frequency (pulse repetition frequency, PRF = number of pulses per second), and receives reflected ones only at a given time and for a limited duration. This pulsed emission–reception process explains why the main limitation of the PW Doppler mode is its inability to measure high velocities: there is a maximum value for PW velocity recording, also called the Nyquist limit. When blood flow velocities exceed the Nyquist limit, an aliasing artifact occurs, characterized by a reversal of the velocities (Figure 16.24). The maximum velocity that can be recorded without aliasing is usually around 1.5–2.5 m/s. Its value depends on both the transducer frequency and the sample gate depth: the lower the transducer frequency and the lower the sample gate depth, the higher the maximal velocity that can be recorded.

(b)

Nevertheless, the advantage of the PW Doppler mode is that blood flows can be analyzed at very specific locations, whereas with the CW Doppler ­ mode it is impossible to determine the accurate ­location of recorded blood flow velocities on the line of interrogation. Color Flow Doppler Mode

The color flow Doppler mode provides a color‐coded map of the velocity and direction of blood flows, which is superimposed in real time on black‐and‐ white 2D‐mode images. Blood flow velocities towards the transducer are coded in red whereas those away from the transducer are coded in blue (Blue Away, Red Toward or “BART”), with lighter colors for higher velocities. Many ultrasound s­ ystems also use variance color maps, adding green to the above‐mentioned colors when the tested flow is turbulent. As color flow imaging provides numerous color sites of flow information, it may be considered as a form of PW Doppler mode with numerous sample gates located on many scan lines. It therefore suffers from the same limitation, the aliasing artifact, which occurs when blood flow velocities exceed the Nyquist limit. This artifact is characterized by a color reversal (blue instead of red, red instead of blue). The maximum value for color‐coded velocity without aliasing is dependent on blood flow depth and transducer frequency, as with the PW Doppler mode. Normal Transvalvular Flows General Characteristics

Transvalvular flows include two arterial flows (pulmonary and aortic flows) and two atrioventricular flows (mitral and tricuspid flows). These four flows are typically laminar, which means that all blood cells are moving in the same

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direction and at approximately the same velocity throughout the entire cardiac cycle. Normal PW Doppler transvalvular flow profiles therefore appear as hollow (Figures  16.21c, 16.21d, 16.22b, 16.22c, 16.23c), because

the blood flow velocities in the selected sample gate are very similar. Conversely, normal CW Doppler flow profiles appear as completely filled because of the various velocities recorded all along the CW scan line (Figure 16.21b).

(a) Right parasternal transventricular short-axis view

Figure 16.13  Two‐dimensional right parasternal transventricular short‐axis view in a normal dog (a) and in three dogs with heart diseases (b–d). (a) The top image shows spatial orientation of the ultrasound beam, with the transducer placed on the right side of the thorax. As shown on the middle image, the ultrasound plane goes first through the right ventricle (RV) and then the left ventricle (LV). Therefore, the real‐time two‐dimensional right parasternal transventricular short‐axis view shows the crescent‐shaped RV at the top of the sector image and the mushroom‐shaped LV below, with the curved interventricular septum (IVS) between the two. Note the two symmetric papillary muscles (Pm) within the LV cavity and the left ventricular free wall (LVFW) at the bottom of the image. Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005. (b) In this dog with dilated cardiomyopathy, the right parasternal transventricular short‐axis view taken in early systole shows an abnormal dilated and rounded (instead of mushroom‐shaped) LV with thin myocardial walls and atrophied left Pm. (c) Unlike (b), in this dog with lipid storage myopathy and associated hypertrophic cardiomyopathy, note the reduced LV cavity, the hypertrophied myocardial walls and left Pm, associated with hyperechoic focal lesions (arrows) due to myocardial fibrotic remodeling. (d) In this dog with pulmonic stenosis, the IVS and the RV myocardial wall (RVW) are severely thickened. Note also the hypertrophied right Pm, the flattened IVS, and the reduced LV cavity.

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(b)

(c)

(d)

Figure 16.13  (Continued)

Normal Pulmonary and Aortic Systolic Flows

Pulmonary flow is assessed from the right or left parasternal transaortic short‐axis views, whereas the left apical five‐chamber view and the subcostal view are commonly used to analyze aortic flow. The subcostal view sometimes provides a better alignment than the  left apical five‐chamber view for imaging aortic outflow. Both arterial flows are systolic (starting from the qRs complex and lasting until the end of the T‐wave) and move away from the transducer. Pulmonary and aortic systolic flows are therefore encoded in blue with color flow Doppler mode, and the corresponding CW and PW systolic flow profiles are negative, with a symmetric

or  asymmetric appearance, respectively (Figures  16.21 and 16.22). Peak arterial flow velocities are usually less than 2 m/s (Tables  16.4 and 16.5). Nevertheless, owing to left ­ventricular outflow tract morphology or high sympathetic tone, the laminar aortic flow profiles of some ­normal dogs may show higher peak values (between 2 and 2.5 m/s). Normal Mitral and Tricuspid Diastolic Inflows

Mitral and tricuspid inflows are both assessed from left parasternal views (the left parasternal apical, or caudal, four‐chamber view and the left parasternal cranial view optimized for right cavities, respectively).

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Right parasternal transmitral short-axis view

Figure 16.14  Two‐dimensional right parasternal transmitral short‐axis view. The top image shows spatial orientation of the ultrasound beam, with the transducer placed on the right side of the thorax. As shown on the middle image, the ultrasound plane goes first through the right ventricle (RV) and then the left ventricle (LV) at the mitral valve level. Therefore, the real‐time two‐dimensional right parasternal transmitral short‐axis view shows the crescent‐shaped RV at the top of the sector image and the rounded LV below, with the curved interventricular septum (IVS) between the two, and the two mitral valve leaflets within the LV cavity. amvl and pmvl: anterior and posterior mitral valve leaflets, respectively. Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005.

Unlike the above‐described arterial flows, mitral and tricuspid flows are both diastolic (starting from the end of the T‐wave and ending at the beginning of the qRs complex) and move toward the transducer. They are therefore encoded in red using the color flow Doppler mode, and the corresponding CW and PW diastolic flow profiles are positive, with a similar biphasic appearance (Figure  16.23) owing to an early

rapid ventricular filling wave (E) and a late filling wave (A) resulting from atrial contraction. Peak early and late diastolic flow velocities are usually less than 1.5 m/s in dogs and cats (Tables 16.4 and 16.5), with an E/A ratio >1 except for healthy aged  animals (as E/A ratio decreases with age). If the heart rate increases, E and A waves can fuse in a single summated signal, and this is commonly observed in the feline species.

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(a) Right parasternal transaortic short-axis view

Figure 16.15  Two‐dimensional right parasternal transaortic short‐axis view in a normal dog (a) and in four dogs with heart diseases (b–e). (a) The right parasternal transaortic short‐axis view is a transverse view of the heart base. The top image shows spatial orientation of the ultrasound beam, with the transducer placed on the right side of the thorax. As shown on the middle image, the ultrasound plane goes first through the right cavities including the right ventricular outflow tract (RVOT) and the right atrium (RA), then the aorta (Ao), and lastly the left atrium (LA). Therefore, the real‐time two‐dimensional right parasternal transaortic short‐axis view shows the RVOT at the top of the sector image with the RA on the left, the circular‐shaped Ao including the more or less visible aortic cusps at the center (appearing like the Mercedes car insignia), and the LA with the left auricle (Laur) at the bottom of the image. Note also the pulmonary trunk (PT) at the bottom right of the image. Measurements of the Ao and LA diameters can be obtained from this view, as follows: the internal short‐ axis Ao diameter is measured along the commissure between the noncoronary and left coronary aortic valve cusps, and the LA is measured in a line extending from and parallel to the commissure between the noncoronary and left coronary aortic valve cusps (double arrows). The LA/Ao ratio is then calculated (see Tables 16.4 and 16.5). Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005. (b) In this dog with end‐stage degenerative mitral valve disease, the right parasternal transaortic short‐axis view shows severe dilation of the LA and a markedly enlarged Laur (arrow). (c) In this dog with valvular pulmonic stenosis, the right parasternal transaortic short‐axis view taken in systole shows thickened immobile pulmonic cusps (arrow) remaining in the center of the PT (instead of being “pushed” against the PT walls). (d) In this dog with severe heartworm infestation, the right parasternal transaortic short‐axis view optimized for the pulmonary arteries shows several worms within the PT as well as the right and left pulmonary arteries (RPA and LPA, respectively). Heartworms appear as linear echoes (arrow) including two parallel lines separated by a thin hypoechoic line. (e) (same view as (d), taken in diastole) In this dog with severe pulmonary arterial hypertension associated with a right‐to‐left PDA, note the severe RPA, LPA and PT dilation (compared with the Ao) and the abnormal doming of the pulmonic cusps (arrow).

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(b)

(c)

(d)

(e)

Figure 16.15  (Continued)

Physiologic Regurgitations

Many healthy dogs display trivial to mild physiologic regurgitations of the pulmonary and tricuspid valves. Physiologic systolic tricuspid regurgitation is also common in cats. Such physiologic regurgitations are  commonly used to noninvasively assess pulmonary arterial pressure, and thus diagnose pulmonary arterial hypertension (see example in Figure 16.25). High regurgitant jet velocities of the tricuspid valve (>2.5 m/s in the absence of pulmonic stenosis) and the pulmonary valve (>2.0 m/s at end-diastole), detected by CW Doppler mode, are usually considered diagnostic for systolic and diastolic pulmonary arterial hypertension, respectively.

Standard Quantitative Doppler Echocardiographic Measurements

Conventional Doppler echocardiography using spectral Doppler modes provides accurate assessment of the direction and peak velocity (m/s) of blood flows. These velocity measurements can be used to estimate the maximum pressure gradients (∆P in mmHg) across valves or between cardiac chambers with the modified Bernoulli equation: 2 P 4 V

where V is the peak velocity (m/s) of blood flow distal to the orifice assessed by spectral Doppler modes.

16  Imaging in Cardiovascular Disease

Table 16.4  Standard echocardiographic and pulsed‐wave Doppler data obtained from 100 healthy dogs of various breeds Variable

Mean ± SD

Range

Heart rate (beats/min)

109 ± 20

70–171

Fractional shortening (%)

36.2 ± 3.5

30.1–49.0

Left atrium/aorta ratio at end‐diastole (2D method)

0.90 ± 0.11 0.52–1.13

Table 16.5  Standard echocardiographic and pulsed‐wave Doppler echocardiographic data obtained from 100 healthy cats of various breeds Variable

Systolic peak aortic flow velocity (m/s) 1.29 ± 0.22 0.92–1.88 Systolic peak pulmonary flow velocity (m/s)

1.05 ± 0.19 0.50–1.50

Mitral E wave velocity (m/s)

0.87 ± 0.13 0.58–1.17

Mitral A wave velocity (m/s)

0.61 ± 0.12 0.39–0.86

Mitral E/A ratio

1.46 ± 0.35 0.92–2.72

Tricuspid E wave velocity (m/s)

0.72 ± 0.11 0.50–0.98

Tricuspid A wave velocity (m/s)

0.43 ± 0.09 0.29–0.70

Tricuspid E/A ratio

1.75 ± 0.34 1.09–2.80

Source: Chetboul V, Carlos Sampedrano C, et al. Use of quantitative two‐dimensional color tissue Doppler imaging for assessment of left ventricular radial and longitudinal myocardial velocities in dogs. Am J Vet Res 2005; 66: 953–61.

Mean ± SD

Range

Left atrium/aorta ratio at end‐diastole (2D method)

0.9±0.1

0.5–1.2

Left ventricular end-diastolic diameter (mm)

15.9±2.3

9.7–21.2

Left ventricular end-systolic diameter (mm)

8.1±1.8

4.1–12.7

Left ventricular end-diastolic free wall (mm)

4.3±0.7

2.4–5.8

Left ventricular end-systolic free wall (mm)

7.5±1.1

4.2–10.3

Interventricular end-diastolic septum (mm)

4.6±0.6

2.9–5.9

Interventricular end-systolic septum (mm)

7.4±1.3

4.6–12.1

Interventricular subaortic septum in end-diastole (mm)

4.1±0.8

2.3–5.7

49±7

33–66

Systolic peak aortic flow velocity (m/s)

1.1±0.2

0.8–1.9

Systolic peak pulmonary flow velocity (m/s)

0.9±0.2

0.5–1.6

Morphologic parameters

Systolic functional parameter

Fractional shortening (%)

Several indices of diastolic function can also be ­calculated. These include the mitral E/A ratio, E wave deceleration time, and the isovolumic relaxation time (time interval between end of aortic flow velocity and onset of transmitral flow), which is an index of active myocardial relaxation (Figure 16.23e). Lastly, the volume of blood flowing through an orifice can be estimated by multiplying the area under the spectral Doppler curve (velocity time integral) by the cross‐sectional orifice area assessed on 2D echocardiographic images.

Common Abnormal Flow Patterns Abnormal Pulmonary Flow Patterns Alterations in Systolic Pulmonary Flow

The most common causes of abnormal systolic pulmonary flow patterns are pulmonary arterial hypertension and congenital pulmonic stenosis. Elevated pulmonary arterial pressures may induce an abnormal asymmetric PW Doppler pulmonary velocity profile with a short acceleration time and a delayed deceleration time, similar to the normal aortic flow profile. A midsystolic notch (Figure  16.26a) may also be seen, most commonly in the case of severe pulmonary arterial hypertension (prevalence of notched pulmonary velocity profiles of 38% for systolic pulmonary arte-

Systolic maximal ejection velocities

Diastolic Doppler parameters in cats with distinct mitral E and A waves

(n = 89)

Mitral E wave (m/s)

0.7±0.1

0.5–1.1

Mitral A wave (m/s)

0.5±0.1

0.3–0.9

Mitral E/A ratio

1.5±0.3

1.1–2.9

Isovolumic relaxation time (ms)

43±9

34–56

Heart rate (beats/min)

184±33

100–261

Time parameters

Source: Chetboul V, Carlos Sampedrano C, et al. Quantitative assessment of velocities of the annulus of the left atrioventricular valve and left ventricular free wall in healthy cats by use of two-dimensional color tissue Doppler imaging. Am J Vet Res 2006; 67: 250–8.

rial pressures >40 mmHg). Peak systolic flow velocity may also be diminished. Pulmonic stenosis is responsible for an increase in systolic pulmonary flow velocity (Figures  16.26b and  16.26c), which can be assessed without aliasing artifact using the CW Doppler mode (Figure  16.26c).

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When the PW Doppler mode is used, the velocity spectrum is broadened, as the flow is turbulent (instead of laminar). Measurement of peak velocity using the CW Doppler mode and subsequent calculation of the maximum systolic pressure gradient across the stenotic lesion provides an accurate and noninvasive estimate of

lesion severity (Figure  16.26c). Pulmonic stenosis is considered as severe when the pressure gradient across the stenotic orifice exceeds 80–100 mmHg. Color flow Doppler mode shows narrowing of the color‐coded systolic pulmonary flow map, and is therefore useful to accurately localize the stenotic lesion (Figure 16.26b).

(a) Right parasternal long-axis 4-chamber view

Figure 16.16  Two‐dimensional right parasternal long‐axis four‐chamber view in a normal dog (a) and in two dogs with heart diseases (b,c). (a) The top image on the left shows spatial orientation of the ultrasound beam, with the transducer placed on the right side of the thorax. As shown on the top image on the right, the ultrasound plane goes first through the two right cavities, i.e., the right ventricle (RV) and the right atrium (RA), then the two left cavities, i.e., the left ventricle (LV) and the left atrium (LA). Therefore, the real‐time two‐dimensional right parasternal long‐axis four‐chamber view shows the RV and the RA separated by the tricuspid valve at the top of the sector image, and the LV and the LA separated by the mitral valve below. The interventricular septum (IVS) and interatrial septum (IAS) are also clearly seen. Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005. (b) In this dog presenting with exercise intolerance, a large defect (arrow) is located in the middle of the IAS ( ostium secundum type atrial septal defect). (c) Two‐dimensional right parasternal long‐axis four‐chamber view obtained in systole from a dog with degenerative mitral valve disease and chordae tendinae rupture. A portion of the ruptured chordae tendinae (arrow) still attached to the prolapsed anterior mitral valve leaflet is seen within the severely enlarged LA.

16  Imaging in Cardiovascular Disease

(b)

cardiomyopathy, respectively. Whatever the cause, aortic stenosis produces an accelerated turbulent systolic aortic flow, with similar spectral and color flow Doppler characteristics to those described for pulmonic stenosis (Figure 16.27). Aortic Insufficiency

(c)

Common causes of aortic insufficiency include infectious endocarditis, congenital abnormal aortic valve (most often in association with subaortic stenosis), and degenerative valvular disease. Color flow Doppler mode criteria indicating severe aortic regurgitation are large insufficiency jet size, compared to the left ventricular outflow tract, and high extension of the diastolic jet within the left ventricle, that is, beyond the mitral leaflets tips. Severe aortic insufficiency also causes a rapid deceleration of the diastolic Doppler CW signal, owing to the marked decrease in the diastolic pressure gradient across the abnormal aortic valve over time. Examples of mild and severe aortic regurgitation jets are presented in Figures  16.27c and 16.28. Abnormal Mitral Flow Patterns Alteration in Diastolic Transmitral Inflow

Figure 16.16  (Continued)

Nonphysiologic Pulmonary Regurgitations

Pulmonary regurgitation is often associated with congenital pulmonic stenosis, because of the abnormally thic­ kened and irregular pulmonary leaflets (Figure  16.26c). Its severity may be mildly to moderately increased in patients with pulmonary stenosis that have undergone balloon valvuloplasty. In contrast, pulmonary regur­ gitation caused by infectious endocarditis is a very rare condition. Physiologic diastolic pulmonary regurgitation becomes turbulent as well as of increased velocity, duration and extension in the case of diastolic pulmonary hypertension (see Figure 16.25 and above paragraph). Abnormal Aortic Flow Patterns Alterations in Systolic Aortic Flow

Aortic stenosis may result from infectious endocarditis. However, the two most common causes of systolic aortic flow obstruction in the dog and cat are cong­ enital aortic stenosis and obstructive hypertrophic

Alteration in left ventricular diastolic function may induce transmitral inflow changes, including delayed relaxation and restrictive transmitral flow patterns (Figures  16.29a and 16.29b). Impaired relaxation may cause lower E wave velocity and higher A wave velocity (E/A 1.5 m/s) suggests elevated left  atrial pressure and therefore severe mitral regurgitation. Mitral Regurgitation

One of the methods commonly used to assess mitral regurgitation severity in dogs with mitral valve dysplasia or degenerative mitral valve disease consists of

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c­ alculating the maximal ratio of the regurgitant jet area signal to left atrium area (ARJ/LAA ratio) using color flow Doppler mode (Figure 16.30). The major advantage of this color Doppler mapping method is the rapidity and ease of data acquisition. Nevertheless, this technique presents several limitations: the maximal value for the ARJ/LAA ratio is 100%, which precludes accurate discrimination between dogs with “significant” mitral

regurgitation. Additionally, the ARJ/LAA ratio may be influenced by several factors including systemic arterial blood pressure, left atrial pressure, and spatial orientation of the regurgitant jet. Note that it may also be possible to quantify (rather than “semi‐quantify”) mitral valve regurgitation (in mL) using another Doppler technique, called the Proximal Isovelocity Surface Area (PISA) or flow convergence method.

(a)

Right parasternal long-axis 5-chamber view

Figure 16.17  Two‐dimensional right parasternal long‐axis five‐chamber view in a normal dog (a) and in three dogs with heart diseases (b–d). (a) The top image on the left shows spatial orientation of the ultrasound beam, with the transducer placed on the right side of the thorax. As shown on the top image on the right, the ultrasound plane goes first through the two right cavities, i.e., the right ventricle (RV) and the right atrium (RA), then the aorta (Ao), and the two left cavities, i.e., the left ventricle (LV) and the left atrium (LA). Therefore, the real‐time two‐dimensional right parasternal long‐axis five‐chamber view shows the RV and the RA separated by the tricuspid valve at the top of the sector image, the LV and the LA separated by the mitral valve below, and a long‐axis image of the Ao too. The interventricular septum (IVS) is continuous with the anterior aortic wall. Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005. (b) A muscular bulge of the IVS into the LV outflow tract (arrow) creating outflow obstruction is clearly visible in this boxer dog with subvalvular aortic stenosis. (c) In this dog suffering from bacterial endocarditis, large vegetative lesions (arrow) are visible on the mitral valve leaflets. (d) In this dog presenting with exercise intolerance, a large ventricular septal defect (arrow) is seen just below the aortic valve. Note also the markedly enlarged right pulmonary artery (RPA).

16  Imaging in Cardiovascular Disease

(b)

(c)

(d)

Figure 16.17  (Continued)

Abnormal Tricuspid Flow Patterns

Abnormal tricuspid flow patterns are similar to those described above for mitral inflow. In the case of s­ ystolic pulmonary hypertension, tricuspid regurgitation demonstrates increased velocity, duration, and extension. The CW Doppler recording of this high peak s­ ystolic tricuspid regurgitant velocity can be used to calculate the systolic right ventricle‐to‐right atrium pressure gradient across the tricuspid valve by applying the modified Bernoulli equation. The systolic pulmonary arterial pressure (equivalent to the systolic right ventricular pressure in the absence of pulmonic stenosis) is then calculated by adding the estimated right atrial pressure (5, 10 or 15 mmHg in patients with a normal

sized right atrium, a dilated right atrium, or right‐sided congestive heart failure, respectively) to the systolic right ventricle‐to‐right atrium pressure gradient. Nonvalvular Abnormal Flows Patent Ductus Arteriosus

Color flow Doppler shows the turbulent jet flowing from the ductus into the pulmonary trunk throughout the cardiac cycle (Figure  16.31a). Continuous‐wave Doppler mode confirms continuous systolic and ­diastolic turbulent flows of high velocity into the pulmonary artery (>5 m/s in systole, around 4 m/s in diastole), and allows estimation of the aorta–pulmonary artery pressure gradient (Figure 16.31b). When the pulmonary

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Section 3  Cardiovascular Disease

Left apical (or caudal) 4-chamber view

Slight rotation to the left

Slight rotation to the left

Left apical (or caudal) 5-chamber view

Figure 16.18  Two‐dimensional left apical (or caudal) four‐ and five‐chamber views. The top image on the left shows spatial orientation of the ultrasound beam, with the transducer placed on the left side of the thorax, to obtain the left apical four‐chamber view. As shown in the top image on the right, the ultrasound plane goes first through the apex, then the right (RV) and left (LV) ventricles, and lastly the left and right atrial cavities (LA and RA, respectively). Therefore, the real‐time two‐dimensional left apical four‐chamber view shows the apex and the two ventricles at the top of the sector image, and the two atrial cavities below, with the mitral valve opened during diastole and closed from end‐diastole to end‐systole (middle and right bottom images, respectively). The left apical four‐ chamber view can be used to measure maximum end‐systolic and end‐diastolic LV length as well as LV volumes. An example of calculation of end‐diastolic LV volume using the Simpson’s derived method of discs is provided on the bottom right image. The endocardial border has been traced and closed around the mitral annulus, thus delimiting the end‐diastolic LV area. The end‐diastolic LV volume, considered as the summation of parallel cylinders, whose diameters are derived from endocardial border tracing, is then automatically calculated using a specific software (62 mL). Another method, also called the length‐area method, relies on the following simple formula: LV volume = 0.85A2/L, where A is the LV area, and L is the LV length measured on the same view. From the left apical four‐chamber view, a slight rotation of the transducer to the left allows visualization of the left ventricular outflow tract and a long‐axis image of the aorta (Ao), thus defining the left apical five‐chamber view (bottom image on the left). See also Figures 16.22 and 16.27 as examples for use of the latter view. Source: Tessier-Vetzel D and Chetboul. In Chetboul et al. 2005.

Box 16.1  Measurement of ventricular volumes Echocardiography can be used to measure ventricular volume using a variety of methods, including the Simpson’s derived method of discs or length‐area method (see explanation in the legend of Figure 16.18). The left ventricular ejection fraction (expressed as a percentage) can then be calculated according to the

f­ollowing formula: 100*(EDV‐ESV)/EDV, with EDV and ESV corresponding to the end‐diastolic and end‐systolic left ventricular volumes. The end‐systolic volume index, defined as the end‐systolic left ventricular volume divided by the body surface area, may be used as an index of systolic myocardial function.

16  Imaging in Cardiovascular Disease

(a)

(a)

(b)

(b)

(c)

Figure 16.19  M‐mode echocardiograms obtained from two normal dogs at the ventricular level (from the right parasternal transventricular short‐axis view, (a)) and the mitral valve level (from the right parasternal transmitral short‐axis view, (b)). (a) This ventricular M‐mode echocardiogram displays the right ventricular myocardial wall (RVW) and the right ventricular cavity (RV) at the top of the image, the left ventricle (LV) and the LV free wall (LVFW) below, with the interventricular septum (IVS) between the two ventricular cavities. The M‐mode cursor is placed perpendicular to the IVS and the LVFW between the two left papillary muscles. The LV end‐diastolic (LVd) and end‐systolic (LVs) diameters can be measured (double arrows), and the left ventricular fractional shortening (expressed in percentage and defined as the difference between the LVd and LVs divided by LVd) can then be calculated. (b) This M‐mode echocardiogram at the mitral valve level shows the M‐shaped motion of the anterior mitral valve leaflet during diastole. The E point and the smaller A point represent the maximum valve opening during the rapid ventricular filling phase and the atrial contraction, respectively. Closure of the mitral valve occurs after atrial contraction, at end‐diastole.

v­ascular resistance increases, lower velocities are recorded across the ductus. Septal Defects (Figures 16.31c and 16.31d)

Color flow Doppler echocardiography associated with spectral Doppler modes may identify small defects that cannot be visualized on 2D mode images. High‐velocity

Figure 16.20  Abnormal M‐mode echocardiograms obtained from three cats with heart diseases at the ventricular level (a,b) and the mitral valve level (c). (a) In this cat with taurine deficiency‐induced dilated cardiomyopathy, the ventricular M‐mode echocardiogram shows marked dilation of the left ventricular cavity (LV) with almost no difference between the end‐diastolic and end‐systolic LV diameters. (b) This ventricular M‐mode echocardiogram in a Maine Coon cat with hypertrophic cardiomyopathy shows severe symmetric hypertrophy (end‐diastolic interventricular septum and LV free wall >6 mm, double arrows). (c) This M‐mode echocardiogram obtained at the mitral valve level in a cat with hypertrophic cardiomyopathy shows a significant systolic anterior motion of the mitral valve (arrows), which is characterized by an abnormal mitral valve septal contact during diastole, leading to LV outflow tract obstruction. IVS, interventricular septum; LVFW, left ventricular free wall.

m/s) flow across the ventricular septal defect (>5  ­associated with normal peak systolic pulmonary flow ­velocity is indicative of “restrictive” ventricular septal shunt, that is, hemodynamically not significant, with conservation of the left ventricle–right ventricle pressure gradient (at least 100 mmHg). Flow velocities across the atrial septal defect are typically low (0.4  mV, >0.04  s

>0.04  s

Left atrial enlargement

II

P‐wave

Notched Right atrial enlargement P‐wave

>0.4  mV

>0.2  mV

>2.5  mV in lead II, aVF

>0.9  mV in lead II

Left ventricular enlargement

III

R‐wave

>3.0  mV in large‐breed dogs >1.5  mV in lead I QRS duration*

aVR

aVL

>0.06  s

>0.04  s

>0.35  mV in lead II

S‐wave in leads I, II,

>0.05  mV in lead I

III and aVF (>0.5  mV)

Right ventricular enlargement S‐wave

S‐wave in leads I, II III and aVF Electrical axis

Right shift (>+100°)

Right shift (>+160°)

Source: Adapted from Tilley and Smith 2008. * QRS duration >0.08 s in the dog and >0.06 s in the cat can be associated with left or right bundle branch blocks. See text for more detail.

aVF

Figure 17.10  Six‐lead ECG example of a right axis shift and right bundle branch block in a cat with right ventricular cardiomyopathy. The leads are I, II, III, aVL, aVR, aVF from top to bottom. The conduction disturbance is identified by the prolongation of the QRS complex (0.07 s, red bar) and a right axis deviation. The right axis shift is denoted by deep S‐waves in leads I, II, III, and aVF (blue arrows) (50 mm/s; 10 mm/mV).

is blocked or slowed within either of the branches and block results in characteristic QRS waveform changes. In order to diagnose a bundle branch block, the MEA has to be determined, which requires a six‐lead ECG

recording. RBBB (see Figure 17.10) is characterized by a right shift of the MEA and markedly prolonged QRS duration (>0.08 s in the dog and >0.06 s in the cat), while LBBB (Figure 17.12) is characterized by a normal MEA and markedly prolonged QRS duration (>0.08 s in the dog and >0.06 s in the cat) (normal values listed in Table 17.3). The QRS morphology with bundle branch block patterns may be confused with a ventricular rhythm. Impulse origin in most cases of bundle branch block is the sinus node, hence the rhythm is supraventricular, despite the marked change in QRS waveform morphology, and there is a P‐wave associated with each QRS complex. RBBB may be a benign finding in both dogs and cats, or associated with right ventricular cardiomyopathy whereas LBBB is usually associated with significant underlying heart disease, such as dilated cardiomyopathy.

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Section 3  Cardiovascular Disease

Figure 17.11  Atrial enlargement is manifest as a P‐wave of increased amplitude in a poodle with degenerative AV valve disease and atrial enlargement. The blue arrow shows the tall P‐ wave (amplitude 0.6 mV) (50 mm/s; 10 mm/mV).

Figure 17.12  ECG example of a left bundle branch block in a rottweiler with dilated cardiomyopathy. The conduction disturbance is identified by the prolongation of the QRS complex (0.08 s; blue bar) but a normal MEA (50 mm/s; 10 mm/mV).

­Further Reading Fogoros RN. The electrophysiology study in the evaluation of supraventricular tachyarrhythmias. In: Electrophysiologic Testing, 3rd edn. Oxford: Blackwell Science, 1998. Kraus MS, Moise NS, Rishniw M. Morphology of ventricular tachycardia in the boxer and pace mapping comparison. J Vet Intern Med 2002; 16: 153. Meurs KM. Boxer dog cardiomyopathy: an update. Vet Clin North Am Small Anim Pract 2004; 34: 1235. Meurs KM, Spier AW, Wright NA, et al. Comparison of the effects of four antiarrhythmic treatments for familial ventricular arrhythmias in Boxers. J Am Vet Med Assoc 2002; 221(4): 522. Moise NS. Diagnosis and management of canine arrhythmias. In: Fox PR, Sisson D, Moise NS, eds.

Textbook of Canine and Feline Cardiology, 2nd edn. Philadelphia, PA: WB Saunders, 1999. Moise NS, Gilmour RF Jr, Riccio ML, et al. Diagnosis of inherited ventricular tachycardia in German shepherd dogs. J Am Vet Med Assoc 1997; 210(3): 403. Oyama MA, Kraus MS, Gelzer AR, eds. Evaluation of the electrocardiogram. In: Rapid Review of ECG Interpretation in Small Animal Practice. Oxford: Taylor and Francis Group, 2014. Tilley LP, Smith WK. Electrocardiography. In: Tilley LP, Smith WK, Oyama MA, Sleeper MM, eds. Manual of Canine and Feline Cardiology, 4th edn. St Louis, MO: Saunders Elsevier, 2008.

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18 Pathophysiology of Heart Failure Barret J. Bulmer, DVM, MS, DACVIM (Cardiology) Tufts Veterinary Emergency Treatment & Specialties, Walpole, MA, USA

Heart failure represents a potential sequela of nearly all congenital and acquired cardiac diseases in dogs and cats. Without an appropriate understanding of the pathophysiologic alterations that produce heart failure, and the knowledge of which therapies can successfully mitigate their consequences, veterinarians will be unable to treat this life‐impairing and often life‐threatening condition. Therefore, the goals of this chapter are to highlight the classification schemes for heart failure, discuss normal cardiovascular physiology along with the mechanisms that contribute to the maladaptive hemodynamic and neurohormonal alterations precipitating heart failure, and outline how understanding of these mechanisms leads to targeted medical therapy.

­Components of the Circulatory System The circulatory system is composed of the heart and its associated arteries, capillaries, and veins. Appropriate function of the circulatory system further requires all of the constituents of normal circulating blood. The heart, blood vessels, and blood carry essential substances to metabolizing tissues and effectively remove unneeded by‐products of metabolism. Additional functions of the circulatory system include regulation of body temperature, humoral communication, and appropriate adjustments of blood delivery depending on the underlying physiologic condition.

­ eterminants of Normal Cardiac D Function Mechanical function of the heart requires coordinated filling of the cardiac chambers (diastolic function) ­followed by an effective contraction (systolic function)

to propel blood from the right and left heart to the pulmonary and systemic circulations, respectively. ­ Impaired mechanical function may occur with disorders of the myocardium, cardiac valves, or pericardium. Effective filling and emptying, along with an appropriate increase or decrease in the rate of contraction, ­further rely on coordinated activity driven by pacemaker cells within the sinoatrial (SA) node and the heart’s conduction system. Therefore, perturbed ­systolic or diastolic function produced by alterations in  myocardial function, valvular integrity, pericardial confinement, volume loading, increased resistance to ejection or electrical instability may all produce cardiac dysfunction with subsequent development of heart ­failure (Box 18.1). Origin of Ventricular Contraction The electrical impulse that produces ventricular systole begins within the SA node and propagates along the specialized conduction system and cell walls to produce atrial and ventricular myocyte depolarization. Calcium influx through the L‐type calcium channels within the sarcolemma along the T‐tubules contributes to a larger calcium release from the sarcoplasmic reticulum (so‐ called calcium‐induced calcium release). The rise in cytosolic calcium enhances binding to troponin‐C and enables configurational changes in the actin and myosin molecules so that cross‐bridge cycling (and hence myocardial contraction during systole) can occur. Diastole (myocardial relaxation) occurs as calcium influx through the L‐type calcium channel and release from the sarcoplasmic reticulum terminate, and calcium is resequestered by the sarco(endo)plasmic reticulum calcium ATPase (SERCA) into the sarcoplasmic reticulum. The fall in cytosolic calcium concentration during diastole results in inhibition of the interaction of

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 3  Cardiovascular Disease

Box 18.1  Disease conditions that may produce heart failure Impaired cardiac filling

Increased resistance to ejection

Pericardial disease

●●

●● ●● ●●

Pericardial effusion with tamponade Constrictive pericarditis Uncommon forms of congenital pericardial disease (e.g., pericardial cysts)

Inflow obstructions ●● ●●

●●

Tricuspid/mitral valve stenosis Uncommon forms of congenital heart disease (e.g., cor triatriatum, supravalvular mitral stenosis) Neoplasia (e.g., obstructive heart base tumors)

Myocardial disease that has a primary component of diastolic dysfunction ●● ●●

Hypertrophic cardiomyopathy Restrictive cardiomyopathy

●●

●●

Pulmonic/aortic valve stenosis Pulmonary/aortic thromboembolism or obstruction (e.g., tumors) Pulmonary hypertension

Volume overload Altered blood flow resulting in volume overload ●●

●●

Valvular insufficiencies (congenital dysplasia, degenerative or infective) Left to right shunting (e.g., patent ductus arteriosis, ventricular septal defect, atrial septal defect, arteriovenous fistulas)

Disease conditions associated with chronically elevated cardiac output ●●

Anemia Hyperthryoidism

Impaired cardiac ejection

●●

Myocardial disease that has a primary component of systolic dysfunction

Disorders of impulse formation or conduction

●● ●●

●●

Dilated cardiomyopathy (idiopathic) Ischemic, nutritional (e.g., taurine deficiency), toxic (e.g., adriamycin), or infectious (e.g., Chagas disease) myocardial disorders Arrhythmogenic right ventricular cardiomyopathy

actin molecules and myosin heads that is required for contraction to occur. Determinants of Ventricular Systolic Function The primary determinants of ventricular systolic function are preload, afterload, and contractility. Heart rate and ventricular synchrony are additional factors that influence systolic function. It has long been recognized that increased cardiac filling enhances systolic performance. In the normal heart, increased preload, which represents the end‐diastolic distending force of the ventricular wall, produces this enhanced performance as defined by Starling’s law of the heart. Surrogates for preload include the end‐diastolic volume, diameter, or end‐diastolic pressure. Therefore, preload is dependent on venous return, total blood volume, and distribution of the blood volume within the vascular system. The recognition that the ascending limb of the Starling curve in skeletal muscle is dictated by the extent and “optimization” of actin‐myosin filament overlap does not appear to be solely responsible for enhanced

Sustained tachyarrhythmias (e.g., supraventricular tachycardia, atrial fibrillation) Sustained bradyarrhythmias (e.g., high‐grade AV block, sinus bradycardia) Wolff–Parkinson–White syndrome

systolic performance in cardiac muscle. An alternative mechanism, termed length‐dependent activation, seems to account for augmented ventricular systolic function with diastolic stretch. The exact mechanisms behind length‐dependent activation remain controversial but the basic premise is that up to a maximum length, the myofilaments become sensitized to calcium as sarcomere length increases. The force, tension or stress experienced by the ventricular myocytes during contraction is termed the afterload. Afterload is the degree of interference to ventricular ejection and is determined primarily by the systemic (or pulmonary) vascular resistance with a contribution from aortic (or pulmonary arterial) impedance. Impedance is dependent on the physical properties of the vascular wall and the blood (e.g., blood density and viscosity, arterial wall diameter, and viscoelasticity) and represents the ratio of aortic pressure to flow. The Anrep effect dictates that an acute increase in afterload enhances ventricular function via stimulation of sarcolemmal stretch receptors and increased cytosolic calcium levels.

18  Pathophysiology of Heart Failure

Contractility is the inherent ability of the cardiac ­ yocytes to shorten and generate force independent of m preload, afterload, or heart rate. Increased contractility represents either increased calcium movement or sensitization of the contractile proteins at a given level of cytosolic calcium. Although criticisms of this concept include the lack of noninvasive measures of isolated contractility and the impossibility of separating the molecular mechanisms of enhanced function from preload and afterload from contractility, this concept is still useful as we discuss the pathophysiology of heart failure. The Bowditch staircase effect (also known as the Treppe effect or the positive force‐frequency relationship) defines that increased heart rate not only increases cardiac output via more frequent contractions, it also enhances the force of ventricular contraction. It is hypothesized that an increased rate of depolarization and the subsequent increased sodium and calcium current across the sarcolemma produce overload of the sodium– potassium pump. The activity of the sodium–calcium exchanger is therefore augmented in the “reverse” mode, extruding sodium from the cell in exchange for increased calcium into the cell. Cross‐bridge cycling is supported by the higher cytosolic calcium level. A final determinant of ventricular systolic function is the coordinated activation (and hence contraction) of the ventricular walls, termed ventricular synchrony. An altered activation sequence and the lack of coordinated mechanical contraction, as witnessed with ventricular arrhythmias, right ventricular apical pacing, or in humans with markedly prolonged QRS durations, can contribute to functional and clinical deterioration. Cardiac (pacemaker) resynchronization therapy has been shown to offset some of these negative consequences in people.

or when the heart can do so only with an elevated filling pressure. The heart’s inability to pump a sufficient amount of blood to meet the needs of the body tissues may be due to insufficient or defective cardiac filling and/or impaired contraction and emptying. Compensatory mechanisms increase blood volume and raise cardiac filling pressures, heart rate, and cardiac muscle mass to maintain the heart’s pumping function and cause redistribution of blood flow. Eventually, however, despite these compensatory mechanisms, the ability of the heart to contract and relax declines progressively, and the heart failure worsens. Although common long‐standing cardiac conditions typically contribute to volume overload and development of pulmonary (left‐sided) or systemic (right‐sided) congestion, there are conditions (e.g., cardiac tamponade, acute pulmonary thromboembolism, acute severe valvular insufficiency) that may severely impair the ability to provide vital organs with their necessary cardiac output without fluid accumulation. Therefore, the term heart failure is often preferred over congestive heart failure. Classification of Heart Failure The American College of Veterinary Internal Medicine (ACVIM), for animals, and the ACCF/AHA, for humans, recognize a functional classification scheme, and a separate, complementary classification system that emphasizes the risk, development, and progression of heart disease. The modified New York Heart Association (NYHA) functional classification is defined as follows. ●●

­The Failing Heart ●●

Definition of Heart Failure The 2013 American College of Cardiology Foundation and American Heart Association (ACCF/AHA) guideline for the management of heart failure defines heart failure broadly as “a complex clinical syndrome that results from structural or functional impairment of ventricular filling or ejection of blood.” The biochemical mechanisms behind this complex syndrome continue to be elucidated but the macroscopic view and description of heart failure has not varied significantly from the conclusion of a panel of the National Heart, Lung and Blood Institute in 1994. They stated: Heart failure occurs when an abnormality of ­cardiac function causes the heart to fail to pump blood at a rate required by the metabolizing tissues

●●

●●

Class I describes patients with asymptomatic heart disease (e.g., heart disease is present but there are no clinical signs even with exercise). Class II describes patients with heart disease that causes clinical signs only during strenuous exercise. Class III describes patients with heart disease that causes clinical signs with routine daily activities or mild exercise. Class IV describes patients with heart disease that causes severe clinical signs even at rest.

A newer veterinary system outlined in the ACVIM Consensus Statement detailing canine valvular heart disease highlights four basic stages of heart disease and failure. Because this system was devised for valvular heart disease, it is not always easily incorporated into discussion of other forms of congenital and acquired heart disease in dogs and cats, but nonetheless it is still useful for discussion of heart failure. The stages of this classification scheme are defined as follows.

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●●

●●

●●

Stage A identifies patients at high risk for developing heart disease but that currently have no identifiable structural disorder of the heart (e.g., every Cavalier King Charles spaniel without a heart murmur, boxers without evidence of arrhythmia or myocardial dysfunction). Stage B identifies patients with structural heart disease but that have never developed clinical signs caused by heart failure. This stage is further subdivided. –– Stage B1 refers to asymptomatic patients that have no radiographic or echocardiographic evidence of ­cardiac remodeling. –– Stage B2 refers to asymptomatic patients that have hemodynamically significant cardiac disease as evidenced by radiographic or echocardiographic findings of heart enlargement. Stage C denotes patients with past or current clinical signs of heart failure associated with structural heart disease. Stage D refers to patients with end‐stage disease with clinical signs that are refractory to “standard therapy.”

The ABCD system is useful because it enables veterinarians to educate owners that their animal is at risk for developing heart disease even in the absence of an audible murmur, gallop or arrhythmia. This system is also useful because treatment and subsequent resolution of signs attributable to heart failure alters patient classification with the modified NYHA model whereas animals that have previously displayed heart failure always remain a Stage C or D with the newer system. Phases of Heart Failure Discussion of heart failure can center around three distinct phases. Phase 1 constitutes the initial cardiac injury and in most veterinary patients, this phase passes silently, without detectable clinical signs, because the disease is mild and slowly progressive (e.g., valvular disease, feline cardiomyopathies). However, some conditions (e.g., development of cardiac tamponade subsequent to bleeding of right atrial hemangiosarcoma or pulmonary thromboembolism) can produce severe life‐threatening signs commensurate with the initial cardiac injury. Phase 2 is the stage wherein short‐term and ultimately long‐term compensatory mechanisms are activated for stabilization of cardiac output and normalization of afterload. Unfortunately, cardiac disease is often progressive and the heart’s ability to hypertrophy may ultimately be ­overwhelmed. A heart murmur, gallop rhythm, cardiac arrhythmia, or cardiomegaly is more likely to be identified in this second phase despite the absence of significant or activity‐limiting clinical signs. The hallmarks of phase 3 of heart failure are the emergence of clinical signs

(e.g., exercise intolerance, lethargy, coughing, tachypnea) at rest or with minimal activity. Phase 1 – Initial Insult

Cardiovascular disease is encountered commonly in daily veterinary practice. Patent ductus arteriosus, subaortic stenosis, and pulmonic valve stenosis are common congenital conditions in dogs while ventricular septal defect and atrioventricular valve dysplasia are more common in cats. Chronic degenerative valvular disease (CDVD) and dilated cardiomyopathy (DCM) are the most common acquired cardiac diseases in dogs while hypertrophic cardiomyopathy (HCM) is the most common acquired cardiac disease in cats. Although identification of disease at its exact onset is next to impossible, it is useful to highlight some of the available data on disease prevalence. Congenital cardiac malformations are uncommon, with data from several veterinary schools suggesting a prevalence of 0.46–0.85%. Feline congenital heart disease is even less common with a reported prevalence of 0.2%. Although these data are several decades old, they still highlight that even at large referral centers, the prevalence of congenital heart disease is low. However, identification of acquired cardiovascular disease is much more common and represents the majority of dogs and cats that ultimately develop heart failure. Dilated cardiomyopathy is encountered most often in large‐ and giant‐breed dogs with a reported prevalence of 0.16–1.1%, depending on the study cited and the population investigated. In some breeds and age groups, the prevalence of DCM is much higher. For example, a study of Doberman pinschers in Europe found that only 3.3% of Dobermans 1 to 170–180  bpm. Ventricular tachycardia is ­considered sustained if it lasts >30 seconds and nonsustained if it lasts 90 ms in cats). Second‐ degree AV block is diagnosed when some P‐waves are not followed by a QRS complex on the surface ECG. Second‐degree AV block is said to be high grade when the number of atrial impulses that fail to be conducted (P‐waves not followed by a QRS complex) to the ven­ tricles are more than the number of impulses that are conducted (P‐waves followed by a QRS complex). Third‐degree or complete AV block is characterized by an absence of conducted P‐waves to the ventricles. The ECG displays independent atrial and ventricular activities (Figure 21.6). Cardiac output is dramatically reduced. In response, the atrial rate, which is under adrenergic tone, is ele­ vated. Electrical activation of the ventricles is dependent on an escape rhythm beyond the site of block. In dogs, the QRS complexes are generally wide and bizarre at rates around 20–60 bpm. In cats, the escape rhythm is usually seen as narrow QRS complexes, indicating their origin in the area of the His bundle, and it occurs at a rate of 80–140 bpm. Finally, the ventricular rate is regular, unless ventricu­ lar premature beats originating from ischemic areas of myocardium are present. Echocardiography

An echocardiogram is indicated to identify concomi­ tant structural cardiac disease, which is frequent in cats with complete AV block. It is also essential to identify cardiac neoplasia that can infiltrate and dis­ rupt the AV nodal tissue, to assess systolic function in the presence of myocarditis, and to determine the degree of volume overload secondary to chronic bradycardia. Other Methods

Serum cardiac troponin I can be used to assess the degree of myocardial damage and raise a suspicion of myocardi­ tis. Although a mild elevation of plasma cardiac troponin I level is common in dogs with complete AV block, marked increases suggest myocarditis as the cause for the AV block.

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(a) II

(b) II

(c)

Figure 21.6  ECG strips of dogs with atrioventricular block. (a) First‐degree atrioventricular block: there is a prolongation of the PR interval (180 ms, recording speed at 50 mm/s) and all the P‐waves are associated with a QRS complex. (b) Second‐degree atrioventricular block: the third and sixth P‐waves (arrows) are not followed by a QRS complex, indicating an intermittent failure of the impulse to propagate to the ventricles. (c) Third‐degree atrioventricular block: there is complete dissociation between the P‐waves and the QRS complexes. The ventricles are activated by an escape rhythm at a rate of 37 bpm (recording speed at 50 mm/s).

Therapy Medical Therapy

Sympathomimetic chronotropes increase heart rate by beta‐adrenergic stimulation. Agents with beta‐2 effects cause systemic vasodilation, whereas drugs with associ­ ated alpha stimulation cause vasoconstriction. Dopamine (5–10 μg/kg/min IV) and dobutamine (2–10 μg/kg/min IV) may contribute to an increase in heart rate and ­systolic function. They are usually administered as a con­ tinuous rate infusion and the dose is increased to effect. They are particularly indicated in the management of beta‐blocker overdose. Isoproterenol, a pure beta‐ago­ nist, improves conduction in the AV node and the His– Purkinje system, which may result in the partial or complete resolution of AV block. It may also increase the rate of a ventricular escape rhythm in complete AV block, but usually with limited success. It is administered as a continuous rate infusion and its dose adjusted to effect. However, it causes a significant decrease in diastolic blood pressure via beta‐2 stimulation. Finally, respiratory and metabolic acidosis decrease its effectiveness.

Most dogs will usually die of noncardiac‐related causes, unless concurrent progressive degenerative valvular dis­ ease or cardiomyopathy is present. Pacemaker implantation may not be required in the vast majority of cats with complete AV block. Indeed, these ani­ mals have a median survival greater than one year without pacemaker therapy. Finally, the presence of underlying structural heart disease or congestive heart failure at the time of diagnosis may not significantly alter their prognosis.

­Atrial Fibrillation Etiology/Pathophysiology

Permanent ventricular pacemaker implantation is the treat­ ment of choice for dogs and cats with clinical AV block.

Atrial fibrillation is characterized by uncoordinated atrial activation from multiple simultaneous electrical wavelets resulting in inadequate mechanical contraction and an irregular ventricular response rate. Atrial dilation is a risk factor for developing AF. However, it commonly occurs in large‐breed dogs with only mild to moderate atrial dilation, and it is rarely present in small‐breed dogs with degenerative mitral valve disease despite extreme atrial dilation. It is also rarely diagnosed in cats. Fibrosis, inflammation, and autonomic imbalances are other important contributors to the initiation of AF.

Prognosis

Epidemiology

In dogs, following permanent pacemaker implantation, estimated survival at one year is approximately 85%.

In dogs, AF is one of the most common types of tachyar­ rhythmias that require treatment. Most dogs with AF

Pacemaker Therapy

21  Supraventricular Arrhythmias

have underlying primary cardiac disease, in the form of dilated cardiomyopathy or degenerative mitral valve dis­ ease. Less commonly, “lone atrial fibrillation” occurs in giant‐breed dogs with structurally normal hearts. Atrial fibrillation is rarely identified in cats but most cats with AF have underlying structural cardiac disease with atrial enlargement. Many of these cats also show signs of con­ gestive heart failure. Signalment The vast majority of dogs with AF are giant‐ and large‐ breed dogs. In the giant‐breed group, Irish wolfhounds, mastiffs, Newfoundlands, rottweilers, and Great Danes are overrepresented, with a male predisposition. Cats with AF are usually older males. History and Clinical Signs Most dogs with “lone” atrial fibrillation are asymptomatic. Conversely, some dogs with compensated cardiac disease and AF show signs of lethargy, exercise intolerance, and rarely anorexia, cough or syncope. When AF occurs in the presence of congestive heart failure, generalized weak­ ness, dyspnea, cough, and abdominal distension from ascites are usually present. Cats frequently show signs of an underlying cardiac disease, including dyspnea and arterial thromboembolism. However, AF can also be an incidental finding on auscultation in this species. Diagnosis Physical Examination

On physical examination, AF is identified as a sustained and irregularly irregular rhythm accompanied with pulse deficits. A murmur may be ausculted, as severe underly­ ing cardiac disease is common in dogs with AF. Electrocardiography

On the surface ECG, AF is an irregular tachyarrhythmia, with usually narrow QRS complexes and no P‐waves that can be replaced by an undulation of the baseline, or F‐waves. It is also permanent, indicating that sinus rhythm never sponta­ neously resumes following initiation of AF. The average heart rate is typically above 180–200 bpm (Figure 21.7). The rate is much lower with “lone” AF, usually approaching 100–120 bpm at rest. Although atrial fibril­ lation is a SVT, concurrent lesions of the ventricular con­ duction system (bundle branch block) may result in widening of the QRS complexes that may resemble those of ventricular tachycardia. Ambulatory 24‐Hour Holter Recording

Determination of the average ventricular rate during AF is critical information in deciding upon a treatment

strategy. Unfortunately, brief ECG recordings obtained in an unfamiliar environment do not adequately reflect the daily variations of heart rate. Heart rate is therefore better assessed from a 24‐hour Holter recording. For ­reference, the average heart rate of healthy dogs varies between 85 and 100 bpm. Conversely, it is approximately 120 bpm in dogs with “lone” AF, 155 bpm in dogs with underlying cardiac and 200 bpm in dogs with congestive heart failure. Therapy Treatment Strategy

A rate‐control strategy is usually applied to the treat­ ment of AF. This approach aims at slowing ventricular rate in response to the rapid supraventricular impulses which are bombarding the AV node. Control of the num­ ber of impulses, which are able to conduct through the AV node, will help in alleviation of clinical signs and pre­ vention of further deterioration of ventricular function. However, it does not terminate the arrhythmia. Rate‐control is based on using drugs to decrease the ability of the AV node to conduct impulses. The need for pharmacologic control of the heart rate is determined from the heart rate distribution obtained from a baseline 24‐hour Holter recording. The effect of rate‐control drugs is then assessed on a follow‐up Holter two weeks after treatment initiation. What constitutes adequate rate control has not been precisely defined and may depend on the animal’s underlying myocardial function. Usually, an average heart rate of no greater than 120– 140  bpm is considered adequate to observe clinical improvement. If the average heart rate is considered too high, drug dosages are increased by small increments with close monitoring of the animal’s clinical status, as response to antiarrhythmics varies between patients. In contrast, medical therapy is usually not needed in dogs with “lone” AF. “Lone” AF can be successfully terminated via DC electrical cardioversion. However, there does not appear to be any benefit in converting the rhythm when adequate/appropriate rate‐control therapy is effective. Medical Therapy Calcium Channel Blockers

Diltiazem is widely used for the management of AF. A graded dose of diltiazem results in a decrease in ven­ tricular response rate, because of slower action potential propagation in the AV node. Oral formulations include an extended‐release form. When evaluated for long‐ term management of AF, a dose of 3–5 mg/kg q12h PO of the extended‐release formulation usually achieves satisfactory rate‐control. Better rate‐control, especially during periods associated with high adrenergic tone, can be obtained by adding digoxin to the treatment regi­ men. Calcium channel blockers decrease ventricular

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21  Supraventricular Arrhythmias

ventricular response rate usually approaches the heart rate recorded in cats with sinus rhythm and heart failure.

flutter results from an electrical impulse circling rapidly and continuously around a large area of myocardium. Focal junctional tachycardia is caused by the rapid dis­ charge of cells in the area of the AV node. Finally, atrio­ ventricular reciprocating tachycardias result from the presence of an accessory pathway, which is a small strand of muscle tissue bridging the atria and ventricles through the cardiac skeleton and forming an alternative route of atrioventricular or ventriculoatrial electrical conduction besides the atrioventricular node. When conduction is also possible from atrium to ventricle, preexcitation may be identified during periods of sinus rhythm as short PR intervals and a widening of the initial portion of the QRS complexes (Figure 21.8). This finding confirms the exist­ ence of an accessory pathway. During episodes of atrio­ ventricular reciprocating tachycardia, the impulse typically descends along the atrioventricular node to the ventricles and returns to the atrium using the accessory pathway. Sustained tachyarrhythmias can lead to the develop­ ment of heart failure, a phenomenon known as tachycar­ diomyopathy. It is likely that a sustained rate above

­ ther Forms of Supraventricular O Tachyarrhythmia Etiology/Pathophysiology Initiation of tachyarrhythmias requires a suitable sub­ strate and precipitating factors. Interstitial fibrosis, inflammation, ischemia or atrial chamber dilation, com­ bined with adrenergic stimulation and electrolyte abnor­ malities promote the risk for tachyarrhythmia. It is therefore expected that SVTs are commonly diagnosed in pets with heart failure. Supraventricular tachyarrhythmias encompass focal atrial tachycardia, atrial flutter, focal junctional tachycar­ dia, and atrioventricular reciprocating tachycardia. Focal atrial tachycardia is caused by the rapid and repeated acti­ vation of a small area of diseased atrial myocytes. Atrial (a)

(b) I

E 6.0~ 0.5 – 4.0 H:W

SA node AP

II AV node

III

aVR

aVL

aVF

Figure 21.8  Ventricular preexcitation. (a) When a dog has an accessory pathway (AP), the sinus impulse may conduct simultaneously through the atrioventricular node and the abnormal muscle bundle. Ventricular activation is initiated without delay from the accessory pathway, resulting in a short PR interval and widening of the QRS complex. (b) Six‐lead ECG of a dog with ventricular preexcitation.

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250  bpm can induce severe myocardial dysfunction within 3–4 weeks in dogs. Epidemiology Supraventricular tachycardias other than AF are rare in both dogs and cats. Signalment Large‐ and giant‐breed dogs, in addition to brachyce­ phalic breeds, are more commonly diagnosed with SVTs. Tachyarrhythmias associated with an accessory pathway have more frequently been reported in Labradors. History and Clinical Signs Signs of SVTs include weight loss, lethargy, exercise intolerance, excessive panting, and dyspnea. However, it is not uncommon for dogs to be perceived as being non­ clinical. Owners of dogs with SVTs may report “seeing” their pet’s heart pounding in the chest during episodes of tachycardia. Weakness and transient loss of conscious­ ness are less common with SVT. Additionally, SVTs may only be recognized at the time of physical examination in dogs with signs of congestive heart failure. Diagnosis Physical Examination

Cardiac auscultation is a useful diagnostic tool to calcu­ late heart rate, detect occasional ectopic beats, and dis­ tinguish between paroxysmal and sustained arrhythmias. While paroxysmal SVTs are usually regular, SVTs may also be irregular. Some dogs experiencing episodes of paroxysmal SVT will be in sinus rhythm at the time of presentation. Femoral pulses may be weaker during bouts of tachycardia. Electrocardiography

It is critical to differentiate supraventricular from ven­ tricular tachycardia. However, treatment can be initiated without identifying the exact mechanism of an SVT. The first step in the diagnosis of tachyarrhythmias is recognizing that an uninterrupted and irregular (a varia­ tion >100  ms between RR intervals) tachycardia is ­typically AF. Once AF is ruled out, SVTs must be differ­ entiated from ventricular tachycardia by looking at the morphology of the QRS complexes. Ventricular tachy­ cardias have wide QRS complexes (>0.06 s in dogs; >0.04 s in cats) followed by a large T‐wave directed opposite to the QRS complex, whereas the main charac­ teristic of SVTs is narrow QRS complexes, indicating that electrical impulses, once they reach the ventricles,

propagate within specialized muscular bundles of the His–Purkinje conduction system, similar to normal sinus beats. However, on occasion, SVTs can display wide QRS complexes if a bundle branch block is present. In these challenging cases, the identification of P‐waves associated with QRS complexes confirms the origin of the arrhythmia above the His bundle. P‐waves can occur before, during or after the QRS complexes. They can be positive or negative. Oftentimes, P‐waves are buried within the T‐waves. Although rarely effective, vagal maneuvers can be done to slow AV conduction, reveal­ ing hidden P‐waves associated to QRS complexes. It is important to consider that ventricular tachycar­ dias are much more frequent than SVTs. Following care­ ful examination of the ECG, if uncertainty persists about the origin of a wide QRS complex tachycardia, it should be first treated as if it were ventricular tachycardia. Ambulatory 24‐Hour Holter Recording

The information collected from Holter recordings includes the number of episodes and their duration, the modes of arrhythmia onset and termination, and infor­ mation about the overall heart rate, especially when the animal is in a familiar environment. Evaluation of heart rate distribution over 24 hours in the light of an activity log completed by the owner during the period of record­ ing is a source of valuable information on the circadian pattern of the arrhythmia and on the contribution of adr­ energic tone to the initiation and rate of the arrhythmia. Echocardiography

An echocardiogram is useful to assess for the presence of structural cardiac disease and tachycardia‐induced cardiomyopathy. Therapy Decision to Treat

Treatment is justified when SVTs cause clinical signs, which are directly related to their rate and duration, as well as ventricular performance. In the absence of obvi­ ous clinical signs, treatment should be considered if tachycardia‐induced cardiomyopathy is suspected. When SVTs are intermittent, relating the clinical signs to the arrhythmia may be challenging. In that situation, long‐term recording of the cardiac rhythm via ambula­ tory 24‐hour Holter recording, or preferably with a wear­ able or an implantable loop event recorder, is indicated. A rate‐control or rhythm‐control strategy, which con­ sists of restoring sinus rhythm, can be applied to the treatment of SVTs. Drug selection is based on the type of arrhythmia, the risk of adverse reactions, and the degree of cardiac dysfunction. Other factors to take into account include decreased oral medication absorption, drug

21  Supraventricular Arrhythmias

metabolism and elimination in the presence of ascites, as well as decreased hepatic and renal blood flow that accompany heart failure. Moreover, drug selection may depend on the level of certainty regarding the nature of the arrhythmia. Indeed, it may be challenging to deter­ mine the origin of wide‐complex tachycardias, which are characteristic of ventricular tachyarrhythmias but on occasion correspond to SVTs with aberrant intraven­ tricular conduction. For example, drugs used for rate‐ control management of SVTs, in particular calcium channel blockers, may cause hemodynamic collapse if given in the presence of rapid ventricular tachycardia and would persist at a fast rate despite treatment. Besides the drugs used for chronic management, paroxysmal SVTs may require emergency intervention when they cause hemodynamic instability. Acute Management

Antiarrhythmic drug administration should always be pre­ ceded by attempts to identify and eliminate potential trig­ gers for the arrhythmia. This includes correcting electrolyte disturbances, hypoxemia, hypovolemia, and acidosis. Common drugs used for the management of SVTs include calcium channel blockers (diltiazem), beta‐ blockers (esmolol), sodium channel blockers (procaina­ mide, lidocaine), and drugs that combine several antiarrhythmic properties (sotalol). Diltiazem

On occasion, diltiazem terminates SVTs if the arrhythmia mechanism is dependent on the AV node. Diltiazem can be administered as intravenous boluses and constant rate infusion for acute management of tachyarrhythmias. Esmolol

Esmolol is a short‐acting intravenous beta‐1‐selective blocker that is rapidly metabolized by blood esterases. It is used for rate‐control and termination of AV node‐ dependent SVTs. However, it may be less effective than intravenous diltiazem. Procainamide

Procainamide has been used for the management of SVTs. It is also indicated for acute management of broad‐ complex tachycardias, when discrimination between SVT and ventricular tachycardia is difficult, as it is also very effective in terminating ventricular arrhythmias. Unfortunately, it is no longer available in an oral formu­ lation and the IV formulation has limited availability outside the United States. Lidocaine

While lidocaine is extensively used as first‐line therapy for acute termination of ventricular tachycardia, its use

for treatment of SVT has only been reported recently. It has been shown to terminate episodes of SVTs in the presence or absence of an accessory pathway. Lidocaine has also been used successfully to terminate recent onset of paroxysmal atrial fibrillation initiated by elevated vagal tone in large‐breed dogs with normal cardiac func­ tion. One to two boluses of lidocaine restored sinus rhythm within 30–90 seconds, while causing mild self‐ limited hypotension following the intravenous bolus. Sotalol

Although availability of the IV formulation of sotalol is limited, the oral formulation can be used for acute man­ agement of SVTs, as most of them are not immediately life‐threatening. Sotalol has potassium channel and mild beta‐blocking properties, which make it a good option to terminate and prevent/decrease the recurrence of focal atrial tachycardias, atrial flutter, and AV reciprocating tachycardia. Conversion of those arrhythmias back to sinus rhythm usually occurs 2–4 hours after oral admin­ istration of the drug. Long‐Term Management

Termination of the arrhythmia and maintenance of sinus rhythm is the preferred approach for the long‐term man­ agement of SVTs (rhythm‐control strategy). It is more likely to succeed when limited structural cardiac changes are present. However, rate‐control strategies used for the management of atrial fibrillation can also safely be applied to the control of most sustained SVTs. Among various antiarrhythmic medications, sotalol has proven to be the most effective rhythm‐control drug. Conversion to sinus rhythm usually occurs just a few hours after oral administration, but it can take up to a few days to observe a response. Sotalol is also effective at preventing arrhythmia recurrence. The negative ino­ tropic effect of sotalol is modest compared to other beta‐ blockers. Nonetheless, dogs with myocardial failure should be closely monitored while receiving sotalol. Atrioventricular reciprocating tachycardia usually responds well to sotalol, which alters the electrophysio­ logic properties of all the cardiac structures necessary for the maintenance of this arrhythmia, including the acces­ sory pathway, the atrial myocardium, the AV node, and the ventricular myocardium. In dogs that fail to respond to monotherapy, sotalol can be combined with mexile­ tine (5–7 mg/kg q8h) with better results. Nonpharmacologic Treatment of Supraventricular Arrhythmias

Synchronized DC cardioversion and radiofrequency catheter ablation have been successfully used to treat supraventricular arrhythmias. In particular, dogs with AV reciprocating tachycardia show the best results with

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radiofrequency ablation. The technique consists in iden­ tifying the location of the accessory pathway, usually around the tricuspid valve annulus, and delivering radi­ ofrequency energy to permanently damage the tissue and block electrical conduction along the abnormal muscle bundle. However, the necessary equipment is available in only a few specialized veterinary centers.

Prognosis Adequate rhythm or rate‐control prevents complica­ tions, such as tachycardia‐induced cardiomyopathy. When the arrhythmia terminates following the adminis­ tration of antiarrhythmic medications, it is critical to monitor with frequent 24‐hour Holter recordings for any signs of arrhythmia recurrence.

­Further Reading Estrada AH, Maisenbacher HW 3rd, Prosek R, Schold J, Powell M, VanGilder JM. Evaluation of pacing site in dogs with naturally occurring complete heart block. J Vet Cardiol 2009; 11: 137–9. Gelzer ARM, Kraus MS, Rishniw, M, et al. Combination therapy with digoxin and diltiazem controls ventricular rate in chronic atrial fibrillation in dogs better than digoxin or diltiazem monotherapy: a randomized crossover study in 18 dogs. J Vet Intern Med 2009; 23: 499–508. Menaut P, Bélanger MC, Beauchamp G, Ponzio NM, Moise NS. Atrial fibrillation in dogs with and without structural and functional cardiac disease: a retrospective study of 109 cases. J Vet Cardiol 2005; 7: 75–83.

Pariaut R, Moise NS, Koetje BD, et al. Lidocaine converts acute vagally associated atrial fibrillation to sinus rhythm in German shepherd dogs with inherited arrhythmias. J Vet Intern Med 2008; 22: 1274–82. Santilli RA, Perego M, Crosara S, et al. Utility of 12‐lead electrocardiogram for differentiating parosxysmal supraventricular tachycardias in dogs. J Vet Intern Med 2008; 22: 915–23. Pariaut R, Santilli RA, Moise NS. Supraventricular tachyarrhythmias in dogs. In: Bonagura JD, Twedt DC, eds. Kirk’s Current Veterinary Therapy XV. St Louis, MO: Elsevier, 2013, pp. 737–44.

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22 Systemic Hypertension Rebecca L. Stepien, DVM, MS, DACVIM (Cardiology) Department of Medical Sciences, University of Wisconsin School of Veterinary Medicine, Madison, WI, USA

­Etiology/Pathophysiology Systemic hypertension (HT) describes a situation of sustained elevation of blood pressure (BP). Systemic ­ hypertension is a clinical condition and should be considered a clinical complication of certain diseases rather than a disease in itself. In veterinary medicine, systolic blood pressure (SBP) is the value most often used to assess BP status in the conscious clinical patient, and systemic hypertension is typically diagnosed when reliable measures of SBP deliver values ≥160 mmHg. Clinically detectable damage caused by HT may be noted in the eyes, central nervous system, heart, and kidneys. Injury related to HT in these organ systems is often collectively termed “target organ damage” (TOD). Target organ damage may be clinically obvious, especially in the ­ocular or nervous systems. In other cases, TOD may be subtle and result in accelerated progression of deterioration of damaged organs (e.g., accelerated deterioration of renal function) rather than overt clinical signs. In veterinary patients, systemic hypertension has typically been identified as one of three types: situational (so‐called “white coat”) hypertension, systemic hypertension secondary to systemic disease, and idiopathic systemic hypertension. Once situational hypertension has been ruled out, systemic hypertension secondary to systemic disease is more common than idiopathic hypertension. The designation of HT as “idiopathic” in a given patient is dependent on a thorough search for common diseases associated with HT. Pathophysiology of Systemic Hypertension The pathophysiologic mechanism(s) of any individual’s elevated BP is likely related to the cause of the HT, but the specific pathways leading to sustained elevations in BP are

poorly understood for most diseases. Blood pressure in health is modulated by interaction of diverse physiologic systems affecting the heart, kidney, brain, and vasculature. These systems rely on neural modulation (via the sympathetic nervous system [SNS]), hormonal modulation (e.g., renin‐angiotensin‐aldosterone system [RAAS], natriuretic peptides), vascular resistance modulation by circulating or local vasoactive mediators (e.g., endothelin, thromboxane, prostaglandins, etc.) and any structural changes (e.g., arteriosclerosis) that may be present. Abnormalities in any of these contributing ­factors or the interaction among these factors may alter arterial BP. In health, increased BP typically leads to natriuresis in the normal kidney and lowering of systemic BP. No matter what the mechanism by which the initial elevation in BP occurs (e.g., catecholamine excess or alterations in RAAS activity), if “normalization” of elevated BP via natriuresis does not occur or is inadequate, abnormality in renal sodium handling is implied, and this may be the final common pathway for many etiologies of HT. The fact that multiple and complicated pathophysiologic pathways are involved in modulation of BP and development of HT has three clinical repercussions. First, HT due to different diseases is likely to respond to different medications, based on the pathophysiologic pathway of the development of HT in an individual patient. Second, lack of complete understanding of these pathways limits our ability to apply specific physiologic blocks to lower BP, and nonspecific vasodilation becomes the treatment of choice for many cases of HT. Last, because any decrease in BP below the individual’s current set point (normal or abnormal) is likely to stimulate reactive, BP‐increasing responses from modulating systems like the SNS and RAAS, simultaneous blockade of these systems may be required when direct vasodilators are used in order to optimize response.

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Pathophysiology of Target Organ Damage Arterial blood flow is maintained at relatively constant levels in the brain, kidneys, and eyes through a process of vascular autoregulation, wherein small resistance vessels constrict when BP is elevated and dilate when BP is decreased to maintain flow. Hypertensive damage in these organs occurs when these autoregulatory mechanisms fail. Elevated BP without adequate resistance vessel constriction causes overdistension of vessels, which leads to damage to endothelial tight junctions and allows protein and plasma leakage into interstitial tissue in affected tissue beds (i.e., tissue edema). If excessive vascular resistance vessel constriction occurs, ischemia of local tissues may result in focal hemorrhage and necrosis. Permanent structural changes such as arteriosclerosis may develop, decreasing vascular distensibility. Any combination of these mechanisms may result in clinical signs of TOD (Table 22.1).

­Epidemiology The most common diseases associated with HT in dogs  include acute or chronic renal disease, especially

protein‐losing renal disease, hyperadrenocorticism, adrenal tumors, including pheochromocytomas and aldosterone‐secreting tumors, and diabetes mellitus. The most common diseases associated with feline HT include chronic renal disease, feline hyperthyroidism, hyperaldosteronism, and diabetes mellitus. In both species, the suspected association (medical or age group related) between subclinical renal dysfunction and both diabetes mellitus and hyperthyroidism may partially account for the occurrence of HT in affected animals. Ten to 15% of hyperthyroid cats that are normotensive at the time of diagnosis may become hypertensive after successful therapy for hyperthyroidism, indicating other mechanisms of HT at work in these patients. The prevalence of idiopathic and situational HT remains unclear for both species.

­Signalment Systemic hypertension is associated with many diseases that are more common in older animals, but aging itself is not a cause of HT. In the case of secondary HT, the signalment of affected animals reflects groups at risk for the underlying disease. In cats, because two common

Table 22.1  Evidence of target organ damage. Clinical signs of target organ damage and additional recommended testing Organ system affected

Associated clinical findings

Recommended diagnostic testing

Eyes

Acute blindness due to retinal detachment or severe hyphema Retinal hemorrhage Retinal vascular narrowing or tortuosity Focal retinal transudates Focal retinal ischemic degeneration Partial or complete retinal detachment Papilledema

Fundoscopic examination Coagulation/platelet assessment if hyphema or retinal hemorrhage

Brain

Seizures (focal facial or grand mal) “Stroke‐like” intracranial neurologic deficits Decreased mentation/obtundation Photophobia Nystagmus

Complete neurologic examination Additional imaging (e.g., MRI)

Kidneys

Proteinuria Microalbuminuria Progressive decrease in renal function

Serum BUN and creatinine Complete urinalysis Urine culture Quantitation of proteinuria or albuminuria Advanced renal testing (e.g., GFR)

Heart

Left ventricular concentric hypertrophy Arrhythmia Gallop rhythm Systolic heart murmur Increased sensitivity to fluid loading (unexpected acute heart failure after fluid administration) Epistaxis

Auscultation Thoracic radiographs Echocardiography Doppler echocardiography may be required to rule out other causes of LV hypertrophy (e.g., subaortic stenosis) Assessment for coagulation/platelet abnormalities if epistaxis

Note: recommended testing list is not exhaustive, additional testing may be indicated in individual patients. BUN, blood urea nitrogen; GFR, glomerular filtration rate; LV, left ventricle; MRI, magnetic resonance imaging.

22  Systemic Hypertension

diseases associated with HT are diseases of older cats (chronic kidney disease [CKD] and hyperthyroidism), the age distribution of hypertensive cats tends to include a greater proportion of older than younger patients.

­History and Clinical Signs Patients with HT are typically identified by BP measurements taken when a suspicious underlying disease or condition (e.g., proteinuria, hyperadrenocorticism) is diagnosed or suspected. Blood pressure should also be monitored in any patient receiving drugs that can cause vasoconstriction, such as phenylpropanolamine, a ­common medication for urinary incontinence that is responsible for BP elevations in some dogs. Additional HT patients are recognized when evidence of TOD (e.g., ocular hemorrhage, neurologic signs, gallop heart sound) is identified. In these patients, further evaluation (see Table  22.1) after diagnosis of HT may reveal the underlying cause. The clinical history of hypertensive patients may be consistent with the underlying ­disease (e.g., polyuria/polydypsia with renal disease), reflect TOD (e.g., acute blindness due to retinal detachment or obtundation due to increased intracranial pressure) or reveal use of hypertensive medications ­ (e.g., phenylpropanolamine). Because of the high variability of clinical signs, and because changes in mentation and behavior due to HT may be subtle or attributed to aging changes, a high level of suspicion for HT should be maintained regarding possible reasons for and signs of HT in clinical patients.

­Diagnosis Patient Selection and Set‐Up Blood pressure should be measured in dogs and cats with diseases known to be associated with HT, and in those animals showing evidence of TOD. In both species, diagnosis or suspicion of renal disease (acute or chronic, proteinuric or nonproteinuric), adrenal neoplasias (e.g., pheochromcytoma or aldosterone‐secreting tumors) and diabetes mellitus should lead to BP assessment. In cats, evaluation of hyperthyroidism should include BP assessment and BP should be reevaluated after successful therapy for hyperthyroidism. Similarly, suspicion of or diagnosis of hyperadenocorticism in dogs should include evaluation of blood pressure. Animals with evidence of TOD (see Table  22.1) should have BP measured at the earliest opportunity regardless of underlying disease, so that intervention may ease or prevent further damage. If an animal with

TOD is diagnosed as HT, antihypertensive therapy should begin immediately. Nonspecific screening of patients for HT is not recommended. The prevalence of HT in healthy‐appearing animals without causative disease is low, so many positive diagnoses of HT in this patient group will be false positives due to situational hypertension. Normal animals may be tentatively diagnosed with HT when elevated BP is a result of agitation or anxiety during BP evaluation. These animals may be identified as anxious during measurement, and repeat measurements later in the day or the following day after a period of acclimation may ­render more accurate results. Animals without causative disease or evidence of TOD may be accepted as normotensive if a confirmatory BP measurement after an initial HT measurement is within normal range. In most cases, animals with known causative disease or TOD and initially elevated BP are confirmed as “true” HT if confirmatory measurement is performed. Commonly available methods to assess BP in dogs and cats include direct measurement (via arterial puncture or arterial cannulation) or indirect (noninvasive) methods, including Doppler sphygmomanometry and oscillometry. Although high‐definition oscillometry (HDO) is now available, results may not be comparable to other methods and repeatable measurements may be difficult to obtain in awake cats. “Standard” oscillometry (widely available in stand‐alone monitors or anesthesia monitors) or Doppler sphygmomanometry is currently recommended for clinical use. Prior to BP measurement, the patient should be allowed to relax with minimal restraint for 5–10 minutes in a quiet area. The owner may help calm the animal ­during the measurement if appropriate. Systolic BP is typically used for clinical decision making and heart rate should be recorded in all patients. Notation of heart rate and any rhythm abnormalities may assist with full evaluation of the patient; tachycardia during BP measurement may signal increased patient anxiety, or provide additional information regarding underlying disease status (e.g., sinus tachycardia or arrhythmias in thyrotoxic cats). A gradually decreasing heart rate during measurement repetitions may occur in a patient that is slowly relaxing, and additional measurements at the lower heart rate may provide more accurate results than the initial measurements at a higher heart rate. Methods of Measurement Direct Blood Pressure Measurement

Direct arterial BP measurement is performed by inserting a needle or catheter attached to a pressure transducer into a peripheral artery in a clinical patient to obtain a  pressure trace. Arterial cannulation using the dorsal

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pedal artery is most frequently used for anesthesia and postanesthesia monitoring, but is typically impractical for clinical use on conscious patients for diagnostic purposes. Direct femoral puncture may be used for acute measurement of BP in conscious animals with local anesthesia, but is time‐consuming and cumbersome in clinical practice. Doppler Sphygmomanometry

Doppler sphygmomanometry (DS) is the most common method of measuring BP in cats and is also used commonly in dogs. A piezoelectric crystal attached to an audio amplifier is applied to the skin overlying a peripheral artery and an occlusion cuff inflated with a pressure bulb is positioned proximally on the same limb. The BP cuff is premeasured and sized such that the width of the cuff is approximately 40% of the circumference of the limb at the level of cuff placement. In dogs and cats, the forelimb is the most common measurement site with the animal in lateral recumbency or sitting and the limb held at the level of the right atrium. The cuff is placed at mid‐antebrachium and attached to the sphygmomanometer. The probe (with coupling gel) is held or taped in place over an artery distal to the cuff (usually the palmar arterial arch) and the position is adjusted until a clear pulsatile audio signal can be detected. The cuff is inflated to approximately 20–40 mmHg past the point at which the sound of blood flow is occluded and then slowly deflated. The pressure at which the sound signal reappears is recorded as the systolic BP. As the cuff is further deflated, the audio signal will become muffled – this pressure may be recorded as diastolic BP but is less reliable than the systolic BP by this method. Three to five repeated measurements taken approximately 30 seconds to one minute apart can be averaged to render a representative SBP. Oscillometry (OSC)

Oscillometric BP devices deliver systolic, diastolic, and mean BP values as well as heart rate. These systems are most reliable in cats when a tailhead cuff is used in unrestrained sternal recumbency, but forelimb (radial level), hindlimb (metatarsal level or proximal to hock in recumbent animal) or tailhead (in standing or recumbent animals) cuffs may be used in dogs. The cuff width is chosen similarly to the DS technique. The cuff is positioned with the bladder of the cuff (usually identified on the cuff by a small arrow) centered squarely over the artery and secured. The OSC machine is set to read BP at approximately one‐minute intervals. The arithmetic mean of 3–5 SBP replicates is used as a representative value, with any obvious erroneous or outlying values discarded. OSC equipment may be unable to

read BP accurately at high heart rates (>180 bpm) or if an arrhythmia is present, and a Doppler or invasive measurement system should be used in these patients. Doppler and OSC devices may not deliver the exact same results in a given patient, nor will different measurements sites in the same patient deliver the exact same results for SBP. Therefore, the same technique and the same cuff site should be used consistently in a given patient when monitoring BP over time.

­Therapy Acute or chronic HT may result in life‐altering or life‐ limiting TOD and may affect the quality and therefore length of life in an affected patient. The goals of HT therapy are to stop or limit the extent of TOD in affected animals and to improve the quality of life in all patients. This may be accomplished by use of medications that decrease SBP and concurrent therapy of underlying causative disease. In all cases, control or resolution of the underlying condition is an important part of HT management. Medical Therapy The goal of therapy of HT is reduction of SBP to 15 kg), subaortic stenosis, thromboembolic disease, immune‐mediated disease, positive blood cultures not meeting major criteria, and Bartonella serology ≥1:1024. A definitive diagnosis of IE involves

Figure 25.3  Echocardiographic long axis image of the aortic valve. A hyperchoic, mobile, irregular‐shaped mass is present adherent to the valve leaflet.

the presence of two major criteria or one major and two minor criteria. The presence of one major and one minor criteria or three minor criteria is suspicious for IE. Treatment Long‐term bactericidal antibiotics are the main therapy for patients affected by IE. Broad‐spectrum combinations of antibiotics such as enrofloxacin and ampicillin should be administered empirically if blood culture is not available or pending. Treatment of Bartonella spp. involves doxycycline, fluoroquinolones, or azithromycin. In dogs requiring hospitalization, intravenous aminoglycosides such as amikacin have been recommended. Ideally, antibiotic therapy is guided by results of culture and sensitivity testing. Oral antibiotics are often continued for 6–8 weeks. Serial echocardiograms or blood ­cultures are used to monitor response to therapy. Patients with heart failure secondary to IE should be treated with standard CHF treatment. Prognosis Generally, patients with IE have a relatively poor prognosis. Outcome is related to the degree of valvular regurgitation and nature of the underlying predisposing cause. Dogs with aortic IE fare worse than dogs with mitral IE (three days versus 476 days median survival time).

­Further Reading Atkins C, Bonagura J, Ettinger S, et al. Guidelines for the diagnosis and treatment of canine chronic valvular heart disease. J Vet Intern Med 2009; 23(6): 1142–50.

Borgarelli M. Crosara S., Lamb K, et al. Survival characteristics and prognostic variable of dogs with preclinical chronic degenerative mitral valve disease

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attributable to myxomatous degeneration. J Vet Intern Med 2012; 26(1): 69–75. Borgarelli M, Savarino P, Crosara S, et al. Survival characteristics and prognostic variables of dogs with mitral regurgitation attributable to myxomatous valve disease. J Vet Intern Med 2008; 22(1): 120–8. MacDonald K. Infective endocarditis in dogs: diagnosis and therapy. Vet Clin Small Anim 2010; 40: 665–84.

Orton EC, Lacerda CMR, MacLea H. Signaling pathway in mitral valve degeneration. J Vet Cardiol 2012; 14(1): 7–17. Reynolds CA, Brown C, Rush JE, et al. Prediction of first onset of congestive heart failure in dogs with degenerative mitral valve disease: the PREDICT cohort study. J Vet Cardiol 2012; 14(1): 193–202.

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26 Canine Myocardial Disease M. Lynne O’Sullivan, DVM, DVSc, DACVIM (Cardiology) Department of Companion Animals, Atlantic Veterinary College, University of Prince Edward Island, Charlottetown, Prince Edward Island, Canada

Canine myocardial diseases are those that affect the heart muscle, as opposed to valvular, endocardial, or pericardial diseases. Cardiomyopathies are the most common myocardial diseases in dogs, and are a “heterogenous group of diseases of the myocardium associated with mechanical and/or electrical dysfunction” according to the American Heart Association. They may be classified clinically according to structural and functional phenotype: dilated, hypertrophic, restrictive, and arrhythmogenic right ventricular. Dilated cardiomyopathy (DCM) is by far the most common cardiomyopathy in dogs. Arrhythmogenic right ventricular cardiomyopathy (ARVC) is seen most commonly in the boxer, and hypertrophic cardiomyopathy (HCM) is reported in dogs but is rare. Restrictive cardiomyopathy is not recognized in dogs. Other forms of canine myocardial disease include myocarditis (infectious, inflammatory, toxic, traumatic), infiltrative disease (neoplastic, storage diseases), ischemic disease, myocardial disease secondary to systemic hypertension, and muscular dystrophies affecting the heart. The focus of this chapter is the cardiomyopathies.

­Dilated Cardiomyopathy Etiology/Pathophysiology Dilated cardiomyopathy is a primary myocardial disease characterized by dilation and systolic dysfunction of the left or both ventricles, in the absence of valvular, pericardial, congenital, coronary artery, or hypertensive heart disease. The DCM phenotype may be the result of various myocardial insults, which are often not identified, rendering many cases idiopathic. In dogs, those “insults” are in many cases genetic mutations leading to altered cardiac protein structure and function, particularly in the pure breeds predisposed to DCM. In humans,

30–50% of cases of DCM are familial or inherited, and mutations involving over 50 cardiac genes have been described to date. Mutations in two genes (PDK4 and titin) have been associated with DCM in Doberman pinschers in North America, but do not account for all cases. The PDK4 mutation was not significantly associated with DCM in a group of European Dobermans in which DCM was linked to a different chromosome ­(candidate genes yet unidentified). This highlights the complex nature of the role that genetics play in this disease and that multiple genetic factors are involved, even within one breed. Juvenile DCM in the Portuguese water dog has been linked to chromosome 8 and candidate gene identification is ongoing. Additional well‐recognized etiologies in dogs include nutritional deficiencies (taurine, carnitine) and tachycardia induced (secondary to sustained tachyarrhythmia). Infectious and toxic causes of myocarditis, characterized by myocardial inflammation and necrosis, can produce a DCM‐like phenotype (ventricular dilation, systolic dysfunction, arrhythmias) in the chronic phase, and may be worth considering as the underlying “insult” in certain clinical presentations or geographic areas. Examples include bacterial (e.g. Borrelia burgdorferi, Bartonella), viral (e.g., parvovirus, West Nile virus), protozoal (e.g., Trypanosoma cruzi [Chagas disease], Toxoplasma gondii), fungal (e.g., Aspergillus, Blastomyces, Coccidioides), and toxic (e.g., anthracyclines) agents. The above‐­ mentioned nonfamilial, nonidiopathic causes make up a very small percentage of the total DCM caseload, as ­idiopathic or familial cases are most common. On a whole‐heart level, the primary abnormality is a reduction in contractility. This results in a reduction in stroke volume (the volume ejected) and an increase in  end‐systolic volume (the volume remaining in the heart after ejection), in turn resulting in progressive increases in end‐diastolic volume. A reduction in

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blood pressure and tissue perfusion activates a number of ­neuroendocrine and cytokine systems intended to maintain stroke volume and blood pressure, primarily through sodium and water retention and vasoconstriction. Stretch receptors and remodeling pathways are triggered, leading to lengthening of individual myocytes (eccentric hypertrophy) and chamber dilation, maladaptive extracellular matrix remodeling, and cell death. Consequences of these alterations in cardiac structure and function ­eventually include congestive heart failure (CHF) and arrhythmias (see Chapter 18). Epidemiology The prevalence of DCM among canine referral populations has been reported to be 0.3–1% (equivalent to 1–3 cases per 300 patients examined). Underreporting of sudden death (SD) cases might result in underestimation of any figure, thus the true prevalence in the general dog population is unknown. Any prevalence data will be heavily influenced by the distribution of breeds and ages (prevalence increases with age) of the population under study. The prevalence is high in certain breeds based on retrospective studies and prospective screening studies. One European study of Irish wolfhounds (IW) found a prevalence of 24%. In the Great Dane, prevalence has ranged from 12% to 36% in different geographic areas. Prevalence in Dobermans has ranged from 45% to 60% in various studies. Some studies report overrepresentation of males, while in Dobermans there are gender differences in how the disease manifests and at what age, with females having a higher incidence of ventricular arrhythmias as the sole abnormality and males manifesting echocardiographic abnormalities and CHF at an ­earlier age. The mode of inheritance is known for specific breed groups. In the Doberman and the Newfoundland, DCM is inherited as an autosomal dominant trait with incomplete penetrance. Both X‐linked and autosomal dominant patterns of inheritance have been suggested in the Great Dane, whereas inheritance is autosomal recessive with sex‐specific alleles in the IW. The juvenile form described in the Portuguese water dog is autosomal recessive. The natural history of the disease is such that dogs experience a preclinical or occult phase during which echocardiographic or electrocardiographic evidence of the disease is present (see Diagnosis) but clinical signs are absent (ACVIM stage B). The length of this phase is variable and likely extends over months to years, depending on the breed. In the Doberman, previous retrospective work suggested a median preclinical phase of just over a year whereas new prospective clinical trial data suggest a median of two years with the use of pimobendan. The preclinical phase can be quite prolonged in the IW, averaging 48 months in one trial.

The overt phase follows, characterized by the onset of clinical signs including syncope, SD, or those typical of CHF (ACVIM stage C). In the Doberman, 25–30% experience SD before CHF. Survival time following the onset of CHF or syncope is again breed and etiology dependent, with Dobermans and Great Danes experiencing the shortest survival times among breeds (see Prognosis). Risk factors for the development of DCM apart from breed may include taurine deficiency (in cocker spaniels, Labrador and golden retrievers, or nontraditional DCM breeds), sustained tachyarrhythmias, myocarditis, or chemotherapy (anthracyclines). Signalment Dilated cardiomyopathy occurs predominantly in middle‐aged to older, medium‐ to large‐breed dogs. The most commonly reported breeds in North America and Europe include the Doberman pinscher, IW, Great Dane, Newfoundland, boxer, Labrador and golden retrievers, cocker spaniel, Scottish deerhound, Afghan hound, St Bernard, German shepherd, Old English sheepdog, Airedale terrier, standard poodle, Rottweiler, and ­mastiff. Small‐breed dogs are uncommonly affected but sporadic cases can be found. Age at diagnosis is typically between 4 and 8 years, depending on breed. Much younger and much older dogs may certainly be affected, regardless of breed. Specific juvenile forms of DCM are described in the Portuguese water dog, Doberman, and toy Manchester terrier, resulting in rapidly progressive CHF or SD at weeks to months of age. Sporadic cases in very young dogs of other breeds may occur, and should prompt ­consideration of viral myocarditis or primary tachyarrhythmia as an underlying cause. Many studies report a gender bias in males. While this is not corroborated in all studies of individual breeds, it is the case for the majority of studies involving multiple breeds. Male Dobermans and IWs may be affected at an earlier age than females. History and Clinical Signs History may be unremarkable and preclinical cases identified by an incidental abnormality on physical exam (such as a heart murmur, gallop, or arrhythmia) or by prospective screening in high‐risk breeds. Some of these dogs in the preclinical stage may have surprisingly profound cardiac dysfunction in the absence of clinical signs. More often, dogs are presented because of clinical signs of a fairly acute nature. Respiratory signs are the most common, including coughing and/or increased respiratory rate or effort due to pulmonary edema from left‐ sided CHF. With severe pulmonary edema, expectoration

26  Canine Myocardial Disease

of pink‐tinged fluid may be noted. Pleural effusion from right‐sided CHF can also produce increased respiratory rate and effort. Other signs include exercise intolerance, weakness, syncope, abdominal distension due to ascites, weight loss, and reduced appetite or anorexia (see Chapter 15). Sometimes SD is the first clinical sign noted. The typical clinical signs exhibited may vary by breed. For example, Dobermans tend to have a higher incidence of syncope and SD and their pulmonary edema and resultant respiratory signs are often quite severe, whereas IWs and Newfoundlands frequently present with evidence of right‐sided CHF (ascites and pleural effusion). The most common abnormalities detected on physical examination are a soft systolic murmur over the mitral or tricuspid area (often grade 1–3/6) and weak peripheral pulses. However, various studies have reported murmurs in as few as 33% and as many as 76%, so the absence of a murmur does not rule DCM out, and physical examination may be normal in the early preclinical phase. Additional findings may include a diastolic gallop (low‐ frequency third heart sound), an arrhythmia (ventricular premature contractions and atrial fibrillation most ­common), pulse deficits (if an arrhythmia is present), and jugular venous distension or pulsation. For dogs in CHF, additional findings may include pale or cyanotic mucous membranes, increased lung sounds or pulmonary crackles, tachycardia, tachypnea, dyspnea, hypothermia, and hepatomegaly (hepatic congestion) or a fluid wave (ascites) on abdominal palpation. Diagnosis The diagnosis of DCM involves a minimum of an echocardiogram and electrocardiogram (ECG). Holter

recording (24‐hour ambulatory ECG) is the most appropriate form of ECG in breeds where ventricular arrhythmias are common or an early finding such as Dobermans, for longer term heart rate (HR) assessment in atrial fibrillation, or if a cause for syncope is sought. In cases where echocardiography is not immediately available, thoracic radiographs may be useful for detecting cardiomegaly, and plasma N‐terminal pro‐B‐type natriuretic peptide (NT‐proBNP) level can be useful in Dobermans for assessing likelihood of preclinical disease. For dogs with suspected CHF, thoracic radiographs and serum biochemistry should be performed. The role for blood‐ based tests including biomarkers, taurine levels, and genetic tests is also discussed below. Screening should be performed yearly in high‐risk breeds. Electrocardiography

While arrhythmias are frequently reported on ECG (89% of cases in one large retrospective study), they can be very intermittent. Ventricular arrhythmias (VA) ­(ventricular premature contractions [VPCs] or ventricular tachycardia [VT]) and atrial fibrillation (AF) are most common, but are not specific for DCM on their own (Figures 26.1 and 26.2). There are differences between breeds in arrhythmia manifestation. VA are very common in Dobermans, boxers, and Great Danes, so their presence should always raise suspicion of DCM in these breeds. In one study in Dobermans, one or more VPCs on a 5‐min ECG was almost 97% specific for detecting a level of VA on Holter recording that would be suggestive of DCM. However, the presence of VPCs on a 5‐min ECG alone is fairly insensitive for detecting DCM. Studies report 44–64% of affected Dobermans and 54% of affected Great Danes

Figure 26.1  Lead II ECG, 12.5 mm/s, 5 mm/mV. VPCs in a Doberman pinscher with DCM. A short run of VT is seen at the beginning of the ECG followed by intermittent single monomorphic VPCs.

Figure 26.2  Lead II ECG, 25 mm/s, 5 mm/mV. Atrial fibrillation in a Golden retriever with DCM. Note the irregular rhythm, absence of P‐waves, and presence of baseline undulations or fibrillation waves.

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had at least one VPC on a 5‐min ECG and during echo exam, respectively, emphasizing that many affected dogs will have a normal in‐hospital ECG. Atrial fibrillation (AF) is more common in IWs and Newfoundlands and is often an early finding in IWs (found in 75% of occult DCM cases, >90% of overt DCM cases). In other breeds it tends to be associated with more advanced or overt disease. Importantly, giant‐ breed dogs can also have “lone” AF unassociated with gross cardiac pathology, in which case the HR is often normal. However, in the IW, it is hypothesized that AF may be a precursor to DCM in many cases. Additional ECG findings can include tall and wide QRS complexes (lead II R‐wave >3.0 mV, QRS duration >0.06 ms) associated with left ventricular (LV) enlargement, notched R‐waves associated with microscopic ischemia, wide P‐waves (lead II P duration >0.04 ms) associated with left atrial enlargement, and sinus tachycardia (particularly in CHF cases). Note that these ECG findings are neither sensitive nor specific for DCM.

severe cases). The distribution of pulmonary edema is often greatest in the caudodorsal lung fields, but can be diffuse in severe cases (Figure 26.3). In Doberman pinschers, the degree of radiographic cardiomegaly is often surprisingly modest, even in the (a)

Holter Recording

Given that arrhythmias apart from AF are typically intermittent and in‐house ECGs are relatively insensitive, Holter recordings can be very useful for diagnosing and quantifying arrhythmias, for seeking cause of collapse by coupling clinical signs with rhythm analysis, and for assessing therapeutic response to antiarrhythmics. In Dobermans, the presence of >300 VPCs/24h, or two subsequent recordings within a year with 50–300 VPCs/24h, may be suggestive of DCM. Coupling a Holter recording with a NT‐proBNP level can increase sensitivity for initial screening (see Circulating Biomarkers). Whether these Holter criteria are appropriate for breeds other than Dobermans is not known, but in one study, 70% of affected Great Danes had a similar degree of arrhythmia. Holter monitoring is also useful for monitoring HR in cases of AF as it yields a more accurate assessment of rate control. Care should be taken to utilize Holter analyses that have been manually edited and verified by qualified personnel as purely automated analyses are notoriously inaccurate.

(b)

Thoracic Radiography

Thoracic radiographs are relatively insensitive to detect the mild cardiac enlargement typical of occult cases. In instances of more severe disease, they can reveal varying degrees of cardiomegaly (left‐sided or generalized). Thoracic radiographs are always indicated if CHF is suspected. In addition to LV enlargement, findings in CHF include left atrial and possibly right‐sided enlargement, pulmonary venous distension and varying degrees of pulmonary edema (interstitial changes in mild to moderate cases and alveolar opacities with air bronchograms in

Figure 26.3  Lateral (a) and VD (b) thoracic radiographs in a Doberman with DCM and CHF. Note the cardiomegaly (including left atrium), pulmonary venous distension, and diffuse interstitial pulmonary infiltrates (worse caudodorsally but also present cranioventrally).

26  Canine Myocardial Disease

face of severe dilation on echocardiography. This ­phenomenon might be due to the Doberman’s relatively deep chest conformation and the relative lack of right heart involvement compared to other breeds. Breeds with frequent right‐sided involvement such as IWs and Newfoundlands often also have pleural effusion, as do dogs that develop AF. Echocardiography

Echocardiography is necessary for identification and quantification of the characteristic structural abnormalities, including LV chamber dilation and systolic dysfunction, and equally important for ruling out other cardiac causes apart from DCM for these findings (e.g., valvular disease, congenital shunt) (Figure 26.4). Increased LV end‐systolic dimension (LVESD) and/or volume (LVESV) is noted, along with increased LV end‐ diastolic dimension (LVEDD) and/or volume (LVEDV).

Linear dimensions (diameters) are generated from an M‐ mode of the LV in a short‐ or long‐axis view immediately below the mitral valve, and volumes are preferentially generated by tracing the LV endocardium in a right parasternal long‐axis or left apical four‐chamber view and using automated software for Simpson’s method of discs. These parameters are often indexed to body weight (BW) or body surface area (BSA) to normalize for differences in body size and conformation. In Dobermans, volume indices have been shown to be more sensitive in detection of LV dilation than diameters. Normal parameters and breed‐specific criteria suggesting the diagnosis of DCM from various studies are presented in Table  26.1. The reader is referred to the Further Reading list for more detailed information on the methods and views used. Left ventricle wall motion is globally and symmetrically reduced in most cases, but some dogs may have increased septal motion compared to the free wall owing

(a)

(b)

(c)

(d)

Figure 26.4  Two‐dimensional echocardiographic images (right parasternal long‐axis view) and M‐mode (right parasternal short‐axis view) from a Doberman with DCM. The left heart is in the far field and right heart in the near field. (a) Note the dilated LV in this systolic frame. (b) M‐mode shows reduced systolic septal and LV free wall motion. (c,d) LV volume measurement in diastole and systole by Simpson’s method of discs.

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Table 26.1  Echocardiographic parameters in normal dogs and breed‐specific criteria suggestive of diagnosis of DCM Parameter

Normal value

Normal parameters a

LVEDD index = LVEDD ÷ BW0.294

1.27–1.85 (95% PI)

LVESD index = LVESD ÷ BW

0.71–1.26 (95% PI)

LVESV index (LVESV ÷ BSA)

48 mm (M) or >46 mm (F) Or >0.1749 × BW + 40.3 mm

LVESD

>36 mm (M or F) Or > 0.1402 × BW + 35.0 mm

LVEDV index (LVEDV ÷ BSA)

>95 mL/m2

LVESV index (LVESV ÷ BSA)

>55 mL/m2

Irish wolfhoundsc LVEDD

>61.2 mm

LVESD

>41 mm

LVESV index (LVESV ÷ BSA)

>41 mL/m2

Great Danesd LVEDD

>56.1 mm (M) or >54.0 mm (F)

LVESD

>42.7 mm (M) or >41.7 mm (F)

LVESV index (LVESV ÷ BSA)

>44 mL/m2

a

 Cornell CC, Kittleson MD, Della Torre P, et al. Allomettric scaling of M‐mode cardiac measurements in normal adult dogs. J Vet Intern Med 2004; 18: 311–21. b  Wess G, Domenech O, Dukes‐McEwan J, et al. European Society of Veterinary Cardiology screening guidelines for dilated cardiomyopathy in Doberman pinschers. J Vet Cardiol 2017; 19(5): 405–15. c  Vollmar AC. The prevalence of cardiomyopathy in the Irish wolfhound: a clinical study of 500 dogs. J Am Anim Hosp Assoc 2000; 36: 125–32. d  Stephenson HM, Fonfara S, López‐Alvarez J, et al. Screening for dilated cardiomyopathy in great Danes in the United Kingdom. J Vet Intern Med 2012; 26: 1140–7. BSA, body surface area; BW, body weight in kg; DCM, dilated cardiomyopathy; F, female; LVEDD, left ventricular end‐ diastolic dimension; LVEDV, left ventricular end‐diastolic volume; LVESD, left ventricular end‐systolic dimension; LVESV, left ventricular end‐systolic volume; M, male; PI, prediction interval.

to mitral regurgitation. Indices of ejection such as fractional shortening (FS) and ejection fraction (EF) are reduced (EF 457 pmol/L (either being abnormal) yields increased sensitivity (94.5%) and specificity (87.8%). A cTnI >0.22 ng/mL (Immulite®, Troponin I, Diagnostics Products Corporation, Los Angeles) yielded 79.5% sensitivity and 84.4% specificity for detection of DCM in another study. Dobermans with NT‐proBNP or cTnI greater than these values should be evaluated with echocardiography. For the differentiation of CHF from primary respiratory disease in dogs presenting with respiratory signs, plasma NT‐proBNP >2447  pmol/L (using second‐­ generation assay) was 81.1% sensitive and 73.1% specific in a study including 8% DCM cases in a variety of breeds. This may have utility when physical exam and thoracic radiography are ambiguous and echocardiography is not available, but otherwise may not provide added value. Plasma Taurine

Whole‐blood or plasma taurine should be measured (using heparinized samples) in cocker spaniels, retrievers, Dalmatians, any atypical DCM breed, or any case with a history of lamb‐based diet, vegetarian diet, low‐protein diet, grain‐free diet (currently being investigated), or any unbalanced home‐made diet. Whole‐blood concentration 180 bpm). Typical choices for AA therapy include sotalol, a AA with both beta‐blockade and K+ channel blockade properties (initiating low doses and titrating upwards is often advised due to the beta‐blockade), mexiletine, a Na+ channel blocker, or a combination of the two. Monitoring the preclinical patient should include physical examination, echocardiography, and ECG or ideally Holter recording every 4–6 months. If AA therapy is initiated, it is best to monitor therapeutic efficacy with Holter monitoring within 1–2 weeks of initiating therapy. Due to a high degree of daily variability, an in‐ house ECG is an inadequate alternative. Overt DCM (ACVIM Stage C, CHF): Outpatient Therapy

Therapeutic goals in the CHF patient include resolution of clinical signs and improvement in quality of life

26  Canine Myocardial Disease

in addition to delay in disease progression and prolongation of survival. Strategies include intravascular volume reduction to resolve pulmonary edema or decrease cavitary effusions, improvement in hemodynamic performance through decreased systemic vascular ­ resistance and improved contractility, neurohumoral modulation, arrhythmia management, and nutritional management. The mainstay of treatment of edema and effusions remains diuretics, with furosemide the most commonly used agent. It may be administered orally or intravenously depending on the severity of CHF and whether the patient is being treated as an inpatient or outpatient. Following edema resolution, the dose may be lowered to the effective dose that maintains the patient free of ­respiratory signs and edema. Given its relatively short duration of action, dosing more than twice daily may be more effective. Substantive pleural effusion or ascites should be relieved with thoraco‐ and abdominocentesis, respectively. Angiotensin converting enzyme inhibitor is indicated for RAAS blockade and vasodilatory effects, particularly in the setting of furosemide use which heightens RAAS stimulation. The results of veterinary clinical trials using enalapril and benazepril report improved quality of life, and although the improvements in survival did not reach statistical significance for DCM dogs, there is clear ­evidence of mortality reduction in human patients and ACEI use is considered standard therapy in dogs with DCM and CHF. Three placebo‐controlled trials have found significantly longer time to treatment failure or longer survival time in DCM dogs treated with pimobendan in addition to diuretics and ACEI, so pimobendan is considered part of standard therapy for CHF due to DCM. The aldosterone antagonist spironolactone significantly reduces morbidity and mortality in humans with CHF and LV systolic dysfunction. While similar evidence is lacking in dogs with DCM, a study in Dobermans with CHF suggested a reduced risk of AF with spironolactone therapy. Spironolactone is often used in the patient that is still symptomatic despite use of triple therapy with furosemide, ACEI, and pimobendan, whereas some, including the author, advocate using it initially as part of standard therapy. Spironolactone is licensed for use in dogs with CHF in Europe. Management of the overt DCM patient with VA includes optimal management of congestion and hemodynamics to reduce sympathetic nervous system stimulation and improve perfusion. Specific AA therapy may also be indicated. Patients with VA may be managed as outlined above for the preclinical stage. DCM patients with AF and in‐hospital HR >150 bpm are candidates for rate control with AA therapy. While cardioversion to

sinus rhythm may be physiologically ideal, rate control is much more feasible. Evidence suggests that the combination of diltiazem with digoxin is more efficacious for rate control than either drug alone. A target mean HR 100 VPCs/24h, presence of couplets, and presence of R‐ on‐T phenomenon) with the presence of two out of three possibly being a more robust way to identify the disease. However, these or any other cut‐offs have yet to be validated prospectively, and to rely solely on the total VPC count may be erroneous. The complexity of the arrhythmia and the complete clinical presentation should be considered when making the diagnosis. Holter recordings can have a high degree of day‐to‐day variability (as much as 80%) and therefore may need to be repeated in dogs with high suspicion yet equivocal results. Myocardial biopsy is not typically used for diagnosis owing to anesthetic requirements and invasiveness, as lesions may be missed given the epicardial and patchy distribution of pathology, and the risk of adverse effects including provocation of arrhythmias or cardiac puncture. Unlike in humans, imaging modalities including echocardiography have low diagnostic yield as they are often normal. In few boxers will RV or LV dilation and systolic dysfunction be appreciable whereas in a case series of English bulldogs, 59% had subjective RV dilation. Cardiac MRI may have utility in detecting the fatty infiltration that characterizes the disease, but this remains to be investigated thoroughly in dogs. Circulating biomarkers including cTnI and NT‐ proBNP are not useful for detection of ARVC, based on studies in boxers. While cTnI levels did correlate with degree of arrhythmia, the degree of overlap with presumably normal boxers renders it an unsatisfactory screening tool.

26  Canine Myocardial Disease

A genetic test for the striatin mutation in boxers is available through the Veterinary Genetics laboratory at North Carolina State University (https://cvm.ncsu.edu/ nc- st ate- ve t-ho spit al/small-animal/gene tic s/). Interpretation and breeding recommendations are as for the DCM genetic tests outlined above, and genetic screening cannot replace clinical screening. Therapy While AAs have been shown to reduce VA frequency and syncope, it is uncertain whether they decrease risk of SD because studies are lacking. Nevertheless, AA therapy remains the mainstay of ARVC treatment. A consensus on arrhythmia threshold necessitating treatment is lacking, and each patient must be approached on a case‐by‐ case basis. Even asymptomatic dogs may be at risk of SD, but this must be balanced with proarrhythmic risk of AA drugs and owner factors including commitment to long‐ term therapy. Suggested candidates for AA therapy include dogs with a history of syncope, those with VT regardless of total VPC number, those with >300 VPC/24h and increased complexity including couplets, triplets, or R‐on‐T (rapid VPCs), or those with >1000 VPC/24h. For immediate treatment of VT or VA causing weakness or collapse, intravenous lidocaine is recommended. The most common and effective oral AAs are sotalol, mexiletine, or a combination of the two in more refractory cases (see Table  26.2). Note that therapy may be indicated prior to Holter recording, as in instances where there is a history of syncope and VT is identified on ECG. Therapy should be monitored by Holter examination within 1–2 weeks of AA initiation, and periodically thereafter (every six months). Therapeutic efficacy is supported by a >80% reduction in arrhythmia number and complexity. An increase in syncope post treatment may suggest a proarrhythmic effect and should be evaluated with Holter promptly. There is some evidence that oral supplementation with omega‐3 fatty acids may also be beneficial (see Table 26.2). For the minority that experience CHF (left, right, or biventricular), therapy with ACEI, pimobendan, and ­diuretics is indicated as described above for DCM, in addition to AA therapy. L‐carnitine may also be added given positive response in a small family of boxers. Prognosis While all dogs with ARVC are at risk of SD, many remain symptom free for years. In one prospective

study of boxers, the median survival time after identification of >300 VPC/24h was just over four years, and in fact overall survival of the affected group was not significantly different from the control boxer group (survival to 10–11 years of age), whereas a retrospective study of boxers with more advanced disease suggested a median survival time of 365 days (range 7–1971 days). The probability of death within a year was almost five times greater with a history of syncope, just over five times greater with >10 000 VPCs/24h, and 20 times greater with >200 runs VT/24h. In a case series of English bulldogs, a median survival time of 8.3 months was reported, but whether the prognosis is truly worse in that breed is not known, as it is likely that these dogs had more advanced disease at the time of diagnosis than other boxer studies.

­Hypertrophic Cardiomyopathy Hypertrophic cardiomyopathy is a primary myocardial disease characterized by symmetric or asymmetric concentric hypertrophy (thickening) of the LV in the absence of a stimulus for hypertrophy (e.g., stenosis or hypertension), leading to diastolic dysfunction and arrhythmias that may result in CHF or SD. In humans, 50% or more of cases are related to familial genetic mutations. While common in the cat, HCM is very rare in dogs. Like in the cat, dynamic LV outflow tract obstruction (LVOTO) caused by systolic anterior motion of the mitral valve (SAM) is described in some cases. Many cases of canine HCM appear to be in young dogs, with males overrepresented. This has raised the question as to whether the disease may be congenital and furthermore, whether LVOTO may represent a form of mitral valve dysplasia predisposing to SAM and secondary hypertrophy. Many affected breeds have been reported and include the pointer, golden retriever, Rottweiler, German shepherd, and shih tzu, amongst others. Cases are frequently evaluated for the presence of an incidental murmur which may be dynamic. Echocardiography is necessary for diagnosis. Treatment is targeted at reducing dynamic LVOTO with beta‐­ blockade (atenolol), and treating CHF or arrhythmias in severe cases. A number of cases in young dogs have been noted to resolve with maturity, while others have ­persisted and resulted in CHF or SD. It is difficult to ­estimate prognosis based on the sparse reports of this disease in dogs.

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­Further Reading Meurs KM. Arrhythmogenic right ventricular cardiomyopathy in the boxer dog: an update. Vet Clin North Am Small Anim Pract 2017; 47(5): 1103–11. Meurs KM, Stern JA, Reina‐Doreste Y, et al. Natural history of arrhythmogenic right ventricular cardiomyopathy in the boxer dog: a prospective study. J Vet Intern Med 2014; 28(4): 1214–20. Summerfield NJ, Boswood A, O’Grady MR, et al. Efficacy of pimobendan in the prevention of congestive heart failure or sudden death in Doberman pinschers with

preclinical dilated cardiomyopathy (the PROTECT study). J Vet Intern Med 2012; 26: 1337–49. Vollmar AC, Fox PR. Long‐term outcome of Irish wolfhound dogs with preclinical cardiomyopathy, atrial fibrillation, or both treated with pimobendan, benazepril hydrochloride, or methyldigoxin monotherapy. J Vet Intern Med 2016; 30(2): 553–9. Wess G, Domenech O, Dukes‐McEwan J, et al. European Society of Veterinary Cardiology screening guidelines for dilated cardiomyopathy in Doberman pinschers. J Vet Cardiol 2017; 19(5): 405–15.

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27 Feline Myocardial Disease Virginia Luis Fuentes, MA, VetMB, PhD, CertVR, DVC, MRCVS, DACVIM (Cardiology), DECVIM (Cardiology) Department of Clinical Sciences and Services, Royal Veterinary College, University of London, Hatfield, Herts, UK

Myocardial disease is extremely common in cats and will be encountered regularly by any feline clinician, but can be hard to recognize. Feline myocardial disease is very heterogeneous in presentation, clinical findings, and prognosis. Without skilled echocardiography, identifying the different subclassifications of cardiomyopathy is usually difficult. Fortunately, this is not usually necessary, as the main priorities for the clinician are to identify those cats with a specific treatable cause of myocardial disease, and to identify cats at high risk of congestive heart failure (CHF) or aortic thromboembolism (ATE).

­Etiology The term “myocardial disease” includes a range of conditions with varied etiologies. Cardiomyopathies are ­usually classified according to a phenotype based on ­cardiac structure and function (Figure 27.1). The myocardium can be affected by systemic conditions such as hyperthyroidism, systemic hypertension, and anemia, where disturbances in hemodynamic loading conditions may alter cardiac structure and function. The myocardium has a limited range of responses to injury or insult, so that different underlying causes can result in the same phenotype. For example, dietary ­taurine deficiency, an uncontrolled tachyarrhythmia or doxorubicin toxicity can all result in an identical “dilated cardiomyopathy phenotype” characterized by systolic dysfunction and ventricular dilation. In other cats, no obvious underlying cause is identified, although a genetic predisposition is suspected in some. Traditionally, feline cardiomyopathies have been classified according to contemporary human classifications, but controversy currently exists over whether human myocardial disease should be classified according to etiology or phenotype. The most commonly recognized phenotype is hypertrophic cardiomyopathy (HCM), characterized by

thickened left ventricular (LV) walls. Other phenotypes include restrictive cardiomyopathy (RCM), defined as relatively normal LV walls but with atrial enlargement and increased atrial pressures; dilated cardiomyopathy (DCM), recognized by atrial and ventricular dilation with global LV hypocontractility; and arrhythmogenic right ventricular cardiomyopathy (ARVC), which is typically manifested by fibrofatty replacement of predominantly the right heart. There is considerable overlap between phenotypes, so an additional category of “unclassified cardiomyopathy” or “non-specific phenotype” is sometimes used to describe patients with a phenotype that does not correspond with any of the above criteria. Unless a causative systemic disease or myocardial insult can be identified, the etiology is unknown for most cats with myocardial disease, although familial HCM is recognized. A genetic cause has been established for HCM in Maine Coon and ragdoll cats, where two different genetic mutations have been identified in the myosin‐binding protein C gene. In contrast, an HCM phenotype has been associated with over 1400 different mutations in humans, although a genetic cause is not identified in all human HCM patients.

­Pathophysiology The functional disturbance and consequent pathophysiology vary according to type of cardiomyopathy and severity of disease. With mild myocardial abnormalities, affected cats may not experience any obvious adverse consequences. With severe or advanced myocardial ­disease, congestive heart failure is the most commonly recognized sign. Sudden death is also a potential sequela, but the prevalence is unknown and probably underestimated. Systemic or arterial thromboembolism is recognized in many advanced cases of cardiomyopathy with left heart involvement.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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ARVC RA, RV dilation

HCM phenotype Dietary taurine deficiency

Increased LV wall thickness

Tachycardia-mediated cardiomyopathy End-stage HCM Systolic dysfunction

DCM phenotype

Chamber dilation, systolic dysfunction

RCM phenotype Normal LV, LA dilation

?Unclassified Abnormalities not fitting other categories

Hypertension Acromegaly

“Transient myocardial thickening”

Hyperthyroidism

Anemia

Figure 27.1  Myocardial disease phenotypes. There is overlap in the cardiac characteristics between the different “classic” forms of cardiomyopathy, and it can be difficult to differentiate cardiomyopathies with a genetic or unknown etiology from cardiomyopathies caused by systemic disease on the basis of cardiac characteristics alone. Hypertrophic cardiomyopathy (HCM) is defined as left ventricular (LV) hypertrophy in the absence of an obvious cause, but hypertension, acromegaly, “transient myocardial thickening,” and hyperthyroidism can all result in a phenotype that is identical to classic HCM. A minority of cats with severe (primary) HCM develop systolic dysfunction, and are described as “end‐stage HCM,” overlapping with dilated cardiomyopathy (DCM). A DCM phenotype can be produced by taurine deficiency or a tachycardia‐mediated cardiomyopathy though no underlying cause is found in most cats with dilation of all four cardiac chambers and global systolic dysfunction. The classic form of restrictive cardiomyopathy (RCM) is defined as normal LV dimensions with biatrial enlargement, but there is overlap with the above phenotypes. Not all cats fit into one of the categories listed above. Those with predominantly right heart enlargement without an obvious cause are usually presumed to be arrhythmogenic right ventricular cardiomyopathy (ARVC), but others do not fit any category and are sometimes called “unclassified cardiomyopathy” or “non-specific phenotype”.

HCM Key pathologic features of HCM include increased LV mass associated with myocyte disarray, interstitial myocardial fibrosis, and narrowing of small coronary arteries. Myocardial hypertrophy may result in delayed LV relaxation, but this is usually well tolerated under normal circumstances and is not likely to result in clinical signs. Many cats with HCM will remain clinically stable provided they are not subjected to a sudden change in hemodynamic load. Examples of such triggers include the stress of a visit to the veterinarian, which increases myocardial oxygen demand; general anesthesia, which can reduce cardiac output; or intravenous fluid therapy, which can result in volume loading. Any of these short‐ term hemodynamic stressors can cause a sudden increase

in left atrial (LA) pressures and acute pulmonary edema. The cats with subclinical HCM that are most vulnerable to these stressors are likely to be those with LA enlargement. Cats with massive LV hypertrophy may also be at increased risk, but the relationship between LV wall thickness and risk is not linear, and LA enlargement is a better global indicator of advanced disease. Many cats with well‐compensated HCM exhibit dynamic LV outflow tract obstruction, which can be either persistent or only induced by excitement/exertion. Outflow tract obstruction is usually caused by systolic anterior motion (SAM) of the mitral valve, where movement of the anterior mitral leaflet towards the interventricular septum during mid to late systole causes obstruction to ejection of blood flow, as well as mitral

27  Feline Myocardial Disease

regurgitation. Outflow tract obstruction can be severe enough to result in a pressure gradient across the LV outflow tract of over 100 mmHg for much of systole. In human HCM patients, the additional myocardial oxygen consumption associated with outflow tract obstruction can cause myocardial ischemia, which is further exacerbated by the concurrent narrowing of small coronary arteries that is a feature of HCM in both humans and cats. Clinical signs of outflow tract obstruction in human HCM patients include exertional dyspnea and chest pain. The exact clinical effect in cats is difficult to determine, as chest pain could easily pass unnoticed by owners, and the majority of affected cats appear to be subclinical. Some owners report open‐mouth breathing during play, despite an absence of documentable pulmonary edema. With progression of HCM, diastolic filling of the LV may be further impaired as interstitial fibrosis worsens, and the LV myocardium becomes stiffer. Ischemic episodes can result in focal areas of myocardial damage, resulting in replacement fibrosis. Neurohormonal activation adds to the problem by causing plasma volume expansion through sodium and water retention, and signs of chronic congestive heart failure become more likely. Signs include pulmonary edema, pleural effusion or both. Cats at this stage are also at risk of arterial thromboembolism, as LA contractile function deteriorates with chronic LA enlargement. A minority of cats will develop generalized LV systolic dysfunction, or large areas of ventricular “scarring” with regional wall thinning and replacement fibrosis. Cats with increased wall thickness but systolic dysfunction are sometimes described as “end‐stage HCM” (ES‐HCM). Although these cats may start with a nondilated hypertrophied LV, over time the LV wall thickness may decrease and LV diameter increases, so that ultimately they may bear little resemblance to the classic HCM phenotype. Occasionally, cats with HCM will present with acute signs of cardiogenic shock, including hypotension, hypothermia, and, often, bradycardia. These low‐output signs may or may not be accompanied by congestive signs. RCM Two forms of RCM have been reported: an endomyocardial form characterized by LV bridging scar (eRCM); and a myocardial form where the LV appears structurally normal but is poorly compliant (mRCM). Varying degrees and distribution of myocardial fibrosis are common to both forms. Biatrial enlargement and increased atrial pressures are also common to both. Cats diagnosed with RCM usually demonstrate clinical signs, which can include congestive heart failure, arrhythmias, arterial thromboembolism, or low‐output signs. It is difficult to differentiate RCM from the more advanced forms of

HCM with myocyte loss or ES‐HCM, and they may in fact be part of the same wide spectrum of HCM. DCM This form of cardiomyopathy is now very uncommon in cats, despite being the most common phenotype in dogs. A DCM phenotype is characterized by systolic dysfunction with normal or reduced LV wall thickness and is associated with an increase in diastolic and systolic ventricular dimensions. An association was identified in the late 1980s between DCM in cats and dietary taurine bioavailability, and since that time, a change in pet food manufacturing processes has resolved this problem. Accordingly, most cats with a DCM phenotype are found to have normal plasma taurine concentrations. Congestive heart failure, arterial thromboembolism, and low‐output signs are all common presentations for DCM. Arterial Thromboembolism All of the above forms of cardiomyopathy may lead to intracardiac thrombus formation if disease is sufficiently advanced (i.e., with left atrial enlargement and dysfunction). Arterial embolization of thrombus (ATE) into the systemic circulation causes vascular occlusion and acute ischemia of downstream tissues, resulting in severe pain, electrolyte disturbances, acute renal injury, and for those surviving the initial embolic episode, a high long‐term risk of ATE recurrence or CHF. Nevertheless, a minority of cats can survive for periods in excess of a year if they recover from the acute crisis. ARVC Arrhythmogenic right ventricular cardiomyopathy also occurs in humans and dogs, and is characterized by fibrofatty replacement of myocytes. The extent and distribution of myocyte replacement are variable, and may result in tachyarrhythmias and/or right atrial and right ventricular dilation. Affected cats may be presented with signs of syncope associated with tachyarrhythmias, but can also develop right‐sided congestive signs with ascites.

­Epidemiology The prevalence of HCM in cats is high; approximately 15% of apparently healthy cats are affected, with a prevalence increasing to nearly 30% of cats over 9 years of age. The prevalence of RCM, DCM, and ARVC is harder to gauge, but is likely considerably lower. HCM is the most common type of feline cardiomyopathy, regardless of age, breed or presenting signs. In the absence of longitudinal epidemiologic studies, it is impossible to determine

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whether RCM is a separate form of cardiomyopathy or an advanced stage variant of HCM.

­Signalment There is a male predominance in cats with HCM. Most cats with HCM are domestic nonpedigree cats, but a number of pedigree breeds are said to be predisposed, including the Maine Coon, ragdoll, Persian, British shorthair, sphynx, Cornish rex, and Norwegian forest cat. Siamese and Oriental cats may be predisposed towards eRCM. Age at diagnosis varies widely, as any age of cat can be affected with HCM, including kittens and geriatric cats, but the prevalence of LV hypertrophy increases with age.

­History and Clinical Signs Cats with advanced myocardial disease are often perceived by their owner to be healthy until shortly before the onset of CHF, ATE, syncope or sudden death. Typical presenting signs for CHF include respiratory distress and tachypnea, whereas acute limb paralysis/paresis is the most common presentation with ATE. Some cats may develop open‐ mouth breathing only with stress or exertion, and may be found to have dynamic LV outflow tract obstruction. Physical examination findings are varied. Murmurs are common in cats with HCM but also in healthy cats with normal hearts, so a murmur should not always be assumed to indicate heart disease. Murmurs are less common in the more severe forms of cardiomyopathy than in well‐compensated HCM. A minority of cats with cardiomyopathy have an audible gallop or arrhythmia, and these auscultation findings are more likely to indicate advanced disease, though can also be detected in some geriatric cats with only mild abnormalities. A prominent apical impulse may be apparent in cats with cardiomegaly or a hyperdynamic LV. Physical exam findings in cats with congestive heart failure will include tachypnea with increased respiratory effort, crackles on pulmonary auscultation with severe pulmonary edema, or absent breath sounds ventrally with a pleural effusion. The presence of a gallop sound in a cat with tachypnea and respiratory distress should prompt treatment for congestive heart failure until further confirmatory tests are possible.

­Diagnosis From the above descriptions, it is clear that establishing a diagnosis of a particular type of cardiomyopathy in an individual cat is challenging. To differentiate end‐stage

HCM, RCM, and UCM requires a high level of echocardiographic skill and experience. A precise diagnosis can be challenging even for experienced cardiologists unless a cat has a classic HCM or ARVC phenotype, due to poor consensus on the definitions of each cardiomyopathy type. Moreover, the labels “end‐stage HCM,” “RCM,” and “UCM” might not actually reflect distinct disease entities but rather different stages of a single condition. Instead, diagnostic tests should be focused on confirming the presence of heart disease; identifying any underlying ­systemic cause of cardiomyopathy that should be treated; and identifying cats at high risk of CHF and ATE. Is Heart Disease Present? In practical terms, the clinician’s first priority when presented with a cat with clinical findings suggestive of heart disease is to establish whether or not cardiac disease is present. The approach will be different for cats with ­respiratory distress versus those that are asymptomatic. Tachypneic Cats

Heart disease should always be considered in cats with tachypnea, and the presence of a murmur, gallop or arrhythmia should increase this suspicion (Figure 27.2). Differentiation of congestive heart failure from other causes of respiratory distress may require diagnostic imaging, but the potential value of radiography or echocardiography must be weighed against the possible risk of subjecting the patient to further stress that could prove fatal. Ultrasound is generally better tolerated than radiography, and has the advantage that patient positioning is less critical. Thoracic ultrasound can demonstrate pleural effusion and can even be used to indicate pulmonary parenchymal changes as with pulmonary edema (B‐line artifacts). If sufficient expertise is available, the most useful way to increase confidence of a cardiac cause of respiratory distress is to demonstrate LA enlargement with echocardiography. Thoracic radiography can be used to identify pleural effusion, pulmonary infiltrates, and assess cardiac size, but must be obtained with great care to avoid compromising patient safety. A point‐of‐care test is available to measure plasma NT‐ proBNP, providing a simple way to help differentiate cats with respiratory distress due to heart failure versus noncardiac causes. Asymptomatic Cats

Myocardial disease should be considered in an otherwise healthy cat when a murmur is detected, and should be strongly suspected when a gallop or arrhythmia is auscultated (Figure 27.3). A very loud murmur (i.e., with a precordial thrill) is usually associated with congenital heart disease.

27  Feline Myocardial Disease

Cat with tachypnea/suspected acute CHF O2, (sedation?) ?Absent breath sounds ventrally

Auscultation

Gallop or Arrhythmia Thoracocentesis

Normal or murmur

Pleural effusion

Normal or equivocal LA

Obvious LA enlargement

Fluid analysis, cytology

Echo not available

“In-house” Echo

Congestive heart failure likely

Additional tests

NT-proBNP

Furosemide

low

high

1–2 mg/kg IV to effect

Cardiomegaly & pulmonary infiltrates

Consider respiratory disease Thoracic radiography Normal heart

If BP 97% considered normal. Pulse oximetry may also be assessed after a short walk, as desaturation may be more commonly observed after exercise in patients with more mild compromise to their airway function. The 6MWT formally measures the distance that a dog can walk over six minutes; distances less than 400 meters are supportive of significant lung disease. The 6MWT may also be combined with pre‐ and post‐walk pulse oximetry to evaluate for exercise‐induced oxygen desaturation.

­Treatment Due to the lack of a definitive treatable etiology for CCB, the treatment options are limited to therapy to ameliorate known clinical signs and co‐morbid conditions. Primary treatment focuses on limiting inflammation, limiting cough, and improving exercise stamina.

Possible environmental causes should be identified and eliminated. Owners should be advised not to smoke indoors, or in closed environments with affected pets. Any air‐borne irritants should be eliminated such as perfumes, incense, scented detergents, and aerosolized cleaners. Owners should also be encouraged to remove as much dust from the environment as possible. Cloth furniture should be cleaned or removed, carpets cleaned or removed, and frequent vacuuming/dusting should be encouraged. If extensive remodeling is planned that is likely to produce potentially noxious fumes or a large amount of aerosolized particles then plans should be made for CCB patients to stay with friends or family. Exposure to potentially sick puppies should be avoided as well as trips to dog parks, grooming parlors, and boarding facilities. If a patient is obese, an appropriate weight loss program should be established. Activities that incite excess barking or strenuous exercise should be curtailed. The importance of replacing neck collars with harnesses should be emphasized. Antiinflammatory medications are typically employed to assist with reducing inflammation in the airway, and glucocorticoids are still the mainstay of treatment. Reducing inflammation can limit cough and assist in reducing the positive feedback cycle that perpetuates CCB. Glucocorticoids may be administered orally or via inhalation. Prednisone is the most commonly used glucocorticoid. Initial dosing of prednisone is 1–2 mg/kg/ day though it should be tapered to the lowest effective dose that controls clinical signs. Inhaled glucocorticoids have been widely used in people, and there is growing frequency of their use in veterinary medicine. Inhaled steroids are delivered via a spacer chamber and facemask designed specifically for dogs (e.g., AeroDawg, wwww.trudellmed.com). Inhaled glucocorticoids are more expensive than oral steroids, but the benefits include a directed therapy approach that limits systemic absorption and subsequent steroid side‐effects. One study demonstrated benefits of therapy with fluticasone at 125 μg q12h. Bronchodilators are commonly prescribed for dogs with CCB and evidence supports an improvement in clinical signs in approximately half of treated dogs. Theophylline (extended release, 10 mg/kg PO q12h) has been shown to have nonspecific effects that may benefit patients with CCB. These include decreasing diaphragmatic fatigue, increasing mucociliary clearance, and enhancing the efficacy of glucocorticoid activity. Beta‐2‐agonists, such as terbutaline, may be less effective due to the lack of reversible bronchoconstriction in some patients. The use of theophylline versus beta‐2‐agonists is still a clinical preference. It should be noted that beta‐2‐agonists might cause some anxiety and restlessness when initiated although these signs often resolve within a few day.

31  Canine Chronic Bronchitis

Box 31.3  Airway sampling techniques Cytologic samples can be obtained from the airway via a tracheal wash, blind bronchoalveolar lavage, or with bronchoscopy. Transtracheal wash (TTW) is best suited for tolerant medium‐ or large‐sized dogs. This technique should be avoided in brachycephalic breeds, and obese patients where easy palpation of the trachea is limited. ●● ●●

●●

●●

TTW is performed in unsedated or lightly sedated dogs. After aseptic preparation of a small area over the cervical trachea or larynx and a local anesthesia block is performed, a through‐the‐needle catheter is passed through the cricothyroid ligament or between tracheal rings; the catheter is fed down the trachea (beveled edge down). Two to three aliquots of 5–10 mL 0.9% sterile saline are flushed into the trachea, and promptly retrieved. Retrieval volume is typically about 50%, and the remaining fluid is rapidly absorbed from the airway. Collection of a diagnostic sample is facilitated by a patient’s ability to cough resulting in an increased likeli-

hood in retrieving a representative cytology from more areas of the airway. Endotracheal wash (ETW) is commonly performed in smaller patients, patients that are less amenable to restrain, brachycephalic breeds, and obese patients. ●●

●●

●●

●●

ETW is performed by briefly anesthetizing the dog with propofol then performing a clean intubation with a sterile endotracheal tube. Lubricating gel should not be used. In a sterile fashion, a 5–8 Fr catheter is fed through the endotracheal tube and sterile saline is infused and reaspirated. Alternatively, a modified technique using suction, a Lukens specimen container, a three‐way stopcock, and a red rubber or Argyle feeding tube can be used (Figure 31.4a,b). 0.9% saline aliquots of 3, 5, and 10 mL are used for dogs 15 kg, respectively. A sterile collection cup may be used to collect any additional samples that may be expectorated.

(a)

(b)

Figure 31.4  Equipment set‐up for a modified technique to perform bronchoalveolar lavage or endotracheal tube lavage using suction, a Lukens specimen container, a three‐way stopcock, and a red rubber or Argyle feeding tube.

309

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●●

Supplemental oxygen should be available during this procedure and for recovery.

Blind bronchoalveolar lavage (BAL) is performed similarly to an ETW with the exception of the following. ●●

●●

The flexible catheter is advanced until it is lodged in the lower airways. A larger volume of 0.9% saline is used, dependent on the patient’s size.

Antibiotics are warranted in dogs with an acute exacerbation of their clinical signs of CCB, or in those with evidence of an infection on airway cytology and/or culture. Doxycycline and azithromycin have antiinflammatory and antimicrobial properties, and should be considered in CCB patients with no specific bacterial culture and sensitivity data. Clinician preference is often influenced by cost, formulation, and availability of products. Fluoroquinolones are also a good choice for patients with CCB due to their high airway tissue concentrations although it is important to remember that concurrent administration of fluoroquinolones and theophylline may results in theophylline toxicity. If simultaneous administration of both drugs is necessary then the theophylline dose should be reduced by approximately 30–40%. Cough suppressants are typically used to reduce the positive feedback cycle of cough begets inflammation and inflammation begets cough, and to improve the quality of life of the patient and owners. Over‐the‐counter cough suppressants are rarely effective in dogs. Narcotic cough suppressants are the most effective, with hydrocodone (0.22–0.5 mg/kg PO q6–12h) the most

●●

A simple technique using a modified stomach tube has been used successfully.

Bronchoscopic bronchoalveolar lavage can be used to ­collect samples via bronchoscopy by flushing sterile 0.9% saline through the chamber of a sterilized scope and reaspirating it back through the biopsy channel.

widely used. Butorphanol (0.5 mg/kg PO q6–12h) may also be considered as a primary cough suppressant. A human study has recently reported on the efficacy of gabapentin for control of cough in humans though data in dogs are lacking.

­Prognosis The clinical course of CCB is highly variable. In the majority of patients, permanent changes are present in the airways at the time of diagnosis, and as a result treatment is not curative. Appropriate medical management can typically help ameliorate the clinical signs and palliate the disease by stopping or slowing progression of bronchial damage. Periodic relapse of cough is not uncommon, and will frequently respond to temporary or permanent adjustment in the therapeutic regimen, or treatment of a developed concurrent illness such as pneumonia. As techniques are developed to screen for and diagnose CCB at earlier stages and more directed medical therapy is established, the prognosis for and probability of cure of CCB will improve.

­Further Reading Amis TC, Kurpershoek C. Tidal breathing flow‐volume loops analysis for clinical assessment of airway obstruction in conscious dogs. Am J Vet Res 1986; 47(5): 1002–6. Bexfield NH, Foale RD, Davison LJ, et al. Management of 13 cases of canine respiratory disease using inhaled corticosteroids. J Small Anim Pract 2006; 47: 377–82. Bolongin M, Kirschvink N, Leemans J, et al. Characterization of the acute and reversible airway inflammation induced by cadmium chloride inhalation in healthy dogs and evaluation of the effects of salbutamol and prednisolone. Vet J 2009; 179(3): 443–50. Brownlie SE. A retrospective study of diagnosis in 109 cases of canine lower respiratory disease. J Small Anim Pract 1990; 31: 371–6.

Chandler JC, Lappin MR. Mycoplasmal respiratory infections in small animals: 17 cases (1988–1999). J Am Anim Hosp Assoc 2002; 38: 111–19. Creevy KE. Airway evaluation and flexible endoscopic procedures in dogs and cats: laryngoscopy, transtracheal wash, tracheobronchoscopy, and bronchoalveolar lavage. Vet Clin North Am Small Anim Pract 2009; 39: 869–80. Hawkins EC, Berry CR. Use of a modified stomach tube for bronchoalveolar lavage in dogs. J Am Vet Med Assoc 1999; 215(11): 1635–9. Hawkins EC, Basseches J, Berry CR, et al. Demographic, clinical, and radiographic features of bronchiectasis in dogs: 316 cases (1988‐2000). J Am Vet Med Assoc 2003; 223(11): 1628–35.

31  Canine Chronic Bronchitis

Hawkins EC, Clay LD, Bradley JM, et al. Demographic and historical findings, including exposure to environmental tobacco smoke, in dogs with chronic cough. J Vet Intern Med 2010; 24: 825–31. Johnson EG, Wisner ER. Advances in respiratory imaging. Vet Clin North Am Small Anim Pract 2007; 37: 879–900. Light RW, ed. Pleural Diseases. Philadelphia, PA: Lippincott Williams and Wilkins, 2007, p. 73. Lux CN, Archer TM, Lunsford KV. Gastroesophageal reflux and laryngeal dysfunction in a dog. J Am Vet Med Assoc 2012; 240(9): 1100–3. Mantis P, Lamb CR, Boswood A. Assessment of the accuracy of thoracic radiography in the diagnosis of canine chronic bronchitis. J Small Anim Pract 1998; 39: 518–20.

Oyama MA, Singletary GE. The use of NT‐proBNP assay in the management of canine patients with heart disease. Vet Clin North Am Small Anim Pract 2010; 40: 545–58. Padrid P. Chronic lower airway disease in the dog and cat. Probl Vet Med 1992; 4(2): 320–44. Padrid PA, Hornof WJ, Kurpershoek CJ, et al. Canine bronchitis: a pathophysiologic evaluation of 18 cases. J Vet Intern Med 1990; 4: 172–80. Ryan NA, Birring SS, Gibson PG. Gabapentin for refractory chronic cough: a randomized, double‐blind, placebo‐ controlled trial. Lancet 2012; 380(9853): 1583–9. Singh MK, Johnson LR, Kittleson MD, et al. Bronchomalacia in dogs with myxomatous mitral valve degeneration. J Vet Intern Med 2012; 26(2): 312–19. Swimmer RA, Rozanski EA. Evaluation of the 6‐minute walk test in pet dogs. J Vet Intern Med 2011; 25: 405–6.

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32 Pulmonary Thromboembolism Robert Goggs, BVSc, PhD, DACVECC, DECVECC Cornell University College of Veterinary Medicine Companion Animal Hospital, Ithaca, NY, USA

Pulmonary thromboembolism (PTE) refers to obstruc­ tion of a pulmonary vessel or vessels by a thrombus and encompasses both in situ thrombus formation and embolization from elsewhere in the vasculature. In small animals, PTE is the preferred term because these mecha­ nisms are difficult to differentiate clinically and both share a common pathophysiology. PTE is associated with numerous diseases and disorders in small animals (Table 32.1) and many patients with PTE have more than one predisposing condition. These underlying condi­ tions can be categorized by how they affect Virchow’s triad, although formation of thrombi in vivo typically involves more than one such effect. The true incidence of PTE in dogs and cats is unknown but it is probably more common than the lit­ erature suggests. Difficulties definitively diagnosing PTE antemortem, limited numbers of postmortem examinations being performed, and rapid postmortem fibrinolysis mean some cases are missed. In a retro­ spective case series of 29 dogs with confirmed PTE, there was antemortem suspicion in less than 40% of dogs in which it was subsequently identified at nec­ ropsy. This study suggested a prevalence of PTE in dogs of 0.9% over a 10‐year period. In cats, PTE was sus­ pected in ~25% cats with respiratory signs and in only 14% of cats in which it was subsequently diagnosed postmortem. Subsequently, a 24‐year prevalence of 0.06% was reported for PTE in cats.

­Pathophysiology Pulmonary thromboembolism frequently results in hypoxemia, hyperventilation, and dyspnea. Arterial hypoxemia in patients with PTE is secondary to abnor­ mal ventilation/perfusion (V/Q) ratios within affected lungs, potentially complicated by diffusion impairment

due to interstitial and alveolar edema. The V/Q mis­ matches occur due to small airway constriction, reduced surfactant production, and development of pulmonary edema and atelectasis. Small airway constriction occurs in both nonperfused and nonembolized areas of lung, which may lead to airway closure and alveolar collapse. Surfactant production is reduced in dogs with experi­ mental PTE, leading to fluid transudation into alveoli (pulmonary edema). Edema may also develop in nonem­ bolized regions due to increased hydrostatic pressure combined with increased microvascular permeability resulting from neutrophil activation. Rarely, pulmonary infarction and pleural effusion may result from complete occlusion of distal pulmonary vascular branches. The cardiovascular consequences of PTE are depend­ ent upon the extent of vessel occlusion. There is substan­ tial reserve capacity in the pulmonary vasculature, which likely accounts for the subclinical nature of many PTE events. In healthy dogs, >60% of the pulmonary vascula­ ture must be occluded before alterations in pulmonary vascular resistance (PVR) reduce pulmonary arterial flow. Reflex vasoconstriction secondary to alveolar hypoxia, humoral factors such as serotonin released from acti­ vated platelets, and neurogenic reflexes may also contrib­ ute. Significant pulmonary vascular occlusion leads to pulmonary hypertension and increased right ventricle (RV) afterload. Severe, acute changes in RV afterload result in dilation and dysfunction. As the RV dilates, the interventricular septum shifts leftward, impairing filling and reducing diastolic distensibility, a concept known as ventricular interdependence. Consequent reductions in left ventricular filling decrease cardiac output and may lead to signs of forward failure (hypotension, cardiogenic shock). If the patient survives an acute crisis but has residual pulmonary hypertension, then clinical signs of backward failure (hepatomegaly, ascites, pleural effusion) may develop over the medium to long term.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 32.1  Recognized risk factors for PTE disease processes with a known association with thromboembolic disease in the dog. Those conditions also associated with an increased risk in the cat are marked by an asterisk. Proposed mechanisms for these risk factors are included

Disease process risk factor

Hypercoagulable state

*Corticosteroid administration

✓

Diabetes mellitus

✓

Dirofilariasis

✓

*DIC (secondary to other disease)

✓

Endocarditis (tricuspid/pulmonic)

✓

✓

*Feline infectious peritonitis

✓

✓

Hyperadrenocorticism

✓

Hypothyroidism

✓

*IMHA

✓

*Indwelling venous catheters

Vascular flow abnormalities/stasis

Endothelial injury/ dysfunction

✓

? ✓

✓ ✓

Myocardial disease

✓

✓

*Neoplasia

✓

✓

*Pancreatitis

✓

*Protein‐losing enteropathy

✓

*Renal amyloidosis/PLN

✓

*Sepsis

✓

*Surgery/trauma

✓

✓

✓ ✓

✓

Source: Adapted from Goggs et al. [1]. DIC, disseminated intravascular coagulation; IMHA, immune‐mediated hemolytic anemia; PLN, protein‐losing nephropathy.

­Clinical Signs The clinical signs of PTE are variable, inconsistent, and nonspecific. The degree of physiologic impairment and thus the severity of clinical signs reflect both the mag­ nitude of the PTE and the patient’s ability to compen­ sate. The most common signs are dyspnea, tachypnea, and depression. Other signs include coughing, hemop­ tysis, cyanosis, syncope, collapse, and sudden death. In dyspneic patients, physical examination may reveal harsh lung sounds and crackles suggestive of pulmo­ nary edema. Occasionally, lung and heart sounds may be muffled due to pleural effusion or in very rare cases pneumothorax. In eupneic patients, lung field ausculta­ tion will likely be normal. On cardiac auscultation, tachycardia with a split second heart sound may be noted or, more commonly, a loud second heart sound  associated with pulmonary hypertension. Signs compatible with backward heart failure (jugular disten­ sion or pulsation, ascites) or forward heart failure (poor peripheral pulse quality, pallor, prolonged capillary refill time) may be present. Complicating the picture

may be clinical signs attributable to the predisposing intercurrent disease. Clinicians should suspect PTE in patients with no history of cardiopulmonary disease who develop res­ piratory distress acutely, particularly where known risk factors exist.

­Diagnosis Suggested Diagnostic Approach Pulmonary thromboembolism is commonly suspected but infrequently conclusively diagnosed, probably because of variability in clinical signs, low test specific­ ity, and limited availability of definitive diagnostics. Establishing a clear diagnosis of PTE remains a challenge in veterinary medicine, even with a logical approach. In human medicine, where definitive diagnostics are widely available, the major challenge is to identify which patients to prioritize for definitive imaging studies. This need has driven the development of clinical prediction rules and diagnostic algorithms.

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Figure 32.2  A lateral thoracic radiograph from a dog with pulmonary thromboembolism showing an area of regional oligemia (Westermark sign) affecting the right caudal lung field.

r­ epresenting zones of reduced blood flow distal to sites of vascular occlusion. This finding, known as the Westermark sign (Figure  32.2), is rare but pathogno­ monic for PTE. Pulmonary infarcts appear as distinct pleural‐based, wedge‐shaped densities. Other potential abnormalities include uneven vessel diameter, pulmonary arterial enlargement, lobar vein or artery attenuation, pleural effusion, and right‐sided cardiomegaly. Arterial Blood Gas Analysis

Typical blood gas abnormalities associated with PTE in dogs are hypoxemia, hypocapnia, and an increased A‐a gradient. In some cases, this hypoxemia may respond poorly to oxygen therapy due to intrapulmonary shunt. These findings are not specific for PTE, however, and should be interpreted in light of other data. Similarly, normal blood gas values do not rule out PTE. Using the A‐a gradient may be more sensitive than solely looking for hypoxemia, since the degree of hypoxemia is propor­ tional to the extent of thromboembolic occlusion. Arterial blood gas analysis may be useful in assessing disease severity, monitoring response to therapy, and determin­ ing prognosis. The ratio of PaO2:PaCO2 in PTE patients was highly predictive of outcome in a recent human study. Blood gas values in cats with PTE have not been reported to date but are expected to mirror those in dogs. Risk Profiling Antithrombin and D‐Dimers

Antithrombin inhibits coagulation factors IIa, IXa, Xa, XIa, and XIIa. In patients with active thrombin produc­ tion, plasma AT activity is reduced by consumption and urinary AT loss has been linked to the hypercoagulable states associated with protein‐losing nephropathy and

hypercortisolemia. Measuring patient levels of AT activity may allow thrombosis risk stratification. Based on extrapolations from humans, reductions of AT ­activity between 50% and 75% moderately increase risk, while activities below 30–50% markedly increase thrombosis risk. Increased plasma D‐dimer concentrations indicate plasmin‐mediated cleavage of factor XIII cross‐linked fibrin. Since D‐dimers require activation of both throm­ bin and plasmin for their formation, they are consid­ ered more specific for thrombosis than fibrin degradation products (FDPs). Rapid, accurate, bedside D‐dimer assays are integral to decision making in humans with possible PTE. People with high clinical probability scores (previous TE disease, deep vein thrombosis, dyspnea) undergo thoracic imaging with­ out a D‐dimer test. For all other people presenting with compatible clinical signs, rapid, quantitative D‐dimer enzyme‐linked immunosorbent assays (ELISAs) are performed prior to selection of further diagnostic test­ ing. Positive D‐dimer tests prompt thoracic imaging, while negative tests suggest no further investigation for PTE is required. Similar guidelines for small animals cannot be formu­ lated currently because we lack sufficient data and, more fundamentally, a convenient and highly accurate D‐ dimer assay is not available for small animals. Although semiquantitative latex agglutination D‐dimer assays per­ form well for disseminated intravascular coagulation (DIC) in dogs, their sensitivity and specificity for throm­ bosis vary widely depending on the cut‐off used. As such, this assay may be best suited to testing patients with a high index of suspicion for PTE, and a cut‐off of >1000 ng/ mL to minimize false positives. Unfortunately, the latex agglutination assay has limited availability and is labora­ tory based. Point‐of‐care (POC) tests allow determination of ana­ lyte values at the bedside, enabling them to be used for clinical decision making in unstable patients. Several canine D‐dimer POC tests have been evaluated for the detection of D‐dimers in patients with thromboembolic disease. Of these, the NycoCard system performed the best, but all of these assays suffer from a lack of specific­ ity for thromboembolic disease. Irrespective of the assay chosen, D‐dimers should be evaluated within 1–2 hours of the suspected embolic event because in experimental canine PTE, D‐dimers were increased by 30 minutes, peaked at >2000 ng/mL at 1–2 hours before falling back to control levels after 24 hours. Viscoelastic Coagulation Testing

Whole‐blood tests of coagulation such as thromboelas­ tography (TEG), thromboelastometry (ROTEM), and  the Sonoclot system graphically represent the

32  Pulmonary Thromboembolism

Figure 32.3  Tissue factor‐activated thromboelastography tracings from a patient with PTE secondary to PLE. The black tracing is a markedly hypercoagulable TEG run at the time of the patient’s PTE. Note the shortened reaction (R) and clot formation (K) times and the increased alpha angle (α), maximum amplitude (MA), and clot elasticity (G) values. The patient was treated with low molecular weight heparin, which significantly prolonged the R and K times and reduced the alpha and MA values (green tracing). A baseline (pink) is displayed for comparison. After stabilizing on therapy, a normal‐looking TEG tracing was produced (orange tracing).

10 millimeters

Events at time of trace At time of PTE

R time (min) 3.0–8.0

K time (min) 3.1–6.7

α angle (°) 28.0–58.9

2.3

0.8

MA (mm) 38.8–59.0

G (d/s) 3.2–7.2 k

78.6

77.1

16.8 k 2.1 k 5.3 k

On LMWH

7.2

9.5

26.1

29.8

PLE stable on Tx

4.2

2.8

54.9

51.4

v­ iscoelastic properties of clotting blood during clot for­ mation and lysis. These different systems produce com­ parable but not interchangeable results. These techniques have been validated in small animals and are now commonly used in veterinary medicine. Using TEG/ROTEM, hypercoagulability has been identified in multiple settings in veterinary patients. Hypercoagulable patients produce characteristic TEG/ROTEM tracings (Figure  32.3). These patients typically have short reaction and clot formation times, steep alpha angles, and large maximum amplitudes. In people, MA values derived from conventional TEG and from a rapid‐TEG assay have been demonstrated to predict thrombotic risk in human surgical patients, and risk of PTE following trauma. It is unclear which (if any) of these parameters best relates to thrombotic risk in veterinary patients. It is also essential to recognize that preanalytical variables such as patient hematocrit and fibrinogen concentration in addition to sample han­ dling and assay parameters contribute to the final TEG/ ROTEM tracings. To date, only one study has reported viscoelastic coag­ ulation testing in small animal PTE. That study did not identify any correlation between TEG variables and PTE, but the sample size was limiting. A recent study evalu­ ated TEG in dogs with thrombosis in various anatomic locations. This study found that although the TEG G‐ values of dogs with thrombosis were significantly greater than controls, half of the dogs’ TEG tracings were clas­ sified as normocoagulable, suggesting that TEG may not have the discriminant power necessary to diagnose PTE. These tests likely have a place in the diagnostic work‐up of possible PTE patients, but more work is needed to identify which test protocols and which

parameters are most predictive of thrombotic risk. The ability of these tests to identify hypercoagulability at the point of care suggests they help identify at‐risk patients who require further investigation, particularly once properly integrated into diagnostic algorithms. Plasma‐Based Coagulation Assays

Tests including the prothrombin time (PT), activated partial thromboplastin time (aPTT), activated clotting time (ACT), and fibrinogen concentration are of limited value in PTE because they may be normal, and any abnormalities are nonspecific. Similarly, abnormalities of FDPs have not been widely identified in small animals with PTE. These molecules indicate plasmin‐mediated degradation of fibrinogen or fibrin has occurred. Increased FDP concentrations are present in thrombo­ sis, but also occur in liver failure, dysfibrinogenemia, excessive fibrinolysis, and DIC, making FDPs less spe­ cific than D‐dimers. Complete Blood Counts/Serum Biochemistry

These tests are not discriminating for PTE, but serum biochemical testing may help identify predisposing con­ ditions such as hyperadrenocorticism, protein‐losing nephropathy, diabetes mellitus, or hypothyroidism. Complete blood counts may help identify a nonspecific inflammatory leukogram or myeloproliferative disor­ ders such as polycythemia or essential thrombocytosis that can predispose to thrombosis. Secondary thrombo­ cytosis does not predispose to PTE, although primary essential thrombocythemia may, particularly if other risk factors exist. Thrombocytopenia or schistocytosis, as markers of DIC, may increase the index of suspicion for PTE.

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artery (PA). To date, RV hypokinesis (apical sparing) has not been confirmed in dogs or cats with PTE. Cardiac Biomarkers

Figure 32.4  Long‐axis four‐chamber echocardiographic view showing a pendulous thrombus (arrowhead) adherent to the roof of the right atrium in this patient. LA, left atrium; LV, left ventricle; RA, right atrium; RV, right ventricle.

Cardiac Function Testing Echocardiography

In people, echocardiography is not a routine diagnostic test for PTE, although case registries report echocardiog­ raphy is performed in 47–74% patients. Although many echocardiography studies are normal in PTE, they may be diagnostic if cardiac or RV outflow tract thrombi are pre­ sent (Figure 32.4). Where PTE is present, echocardiogra­ phy can quantify the hemodynamic consequences of PTE and provides valuable prognostic information to guide patient management. Echocardiography can also identify differential diagnoses such as cardiomyopathy, endocar­ ditis or pericardial tamponade noninvasively. Typical echocardiographic findings in humans with PTE include right ventricular dilation and hypokinesis, septal flattening and paradoxical septal motion, diastolic left ventricular impairment, pulmonary arterial hyper­ tension, right ventricular hypertrophy, and patency of the foramen ovale. The degree of right ventricular dys­ function is related to the cross‐sectional area of the pul­ monary vasculature affected; occlusion of >30% is frequently associated with RV hypokinesis. In people with severe PTE, the apex of the RV appears to be spared the hypokinesis that affects the remainder of the right ventricular free wall (RVFW). This apical sparing (the McConnell sign) is highly specific for the diagnosis of PTE in people. There are currently few data regarding echocardiogra­ phy in canine PTE because in recent retrospective stud­ ies 50% cats, all 11 patients in the study suffered side‐effects, many attributable to reperfusion injury. The t‐PA products are fibrin specific, activating fibrin‐bound plasminogen more rapidly and more effec­ tively than plasminogen in circulation. At pharmaco­ logic concentrations, however, a systemic lytic state can still occur and all t‐PA products are associated with a risk of hemorrhage in humans. Alteplase has the short­ est half‐life of the available recombinant thrombolytics.

32  Pulmonary Thromboembolism

Alteplase has been successfully used in one dog with ATE, and experimentally induced canine PTE has been treated with t‐PA therapy. Currently, the American College of Clinical Pharma­ cology (ACCP) recommends thrombolytic therapy for  people with hypotension secondary to acute PE. Thrombolysis in PTE in the absence of hypotension is not currently recommended, but could be considered in patients with echocardiographic evidence of cardiac dys­ function or in those with high cTnI or NT‐proBNP con­ centrations. A recent trial of normotensive PTE patients with biomarker and echocardiographic evidence of myo­ cardial damage and dysfunction evaluated thrombolysis with tenecteplase. Thrombolysis in these patients sig­ nificantly reduced the rate of death due to hemodynamic collapse, but at the cost of a significant increase in major bleeding events. Where thrombolytic agents are used, the ACCP rec­ ommend they be administered over a short time period via a peripheral catheter rather than a PA catheter. Case registry data suggest thrombolytics are used in only 30% of people with massive PTE. In veterinary medicine, no consensus on their administration exists and clinicians must determine the potential benefits and risks of administration for individual patients. Recent advances in small animal interventional radiology may provide novel methods or routes for thrombolysis in future. Based on the ACCP guidelines, thrombolytic therapy should only be considered in veterinary patients with hemodynamically unstable acute PTE and where con­ tinuous hemodynamic monitoring is available. In appro­ priate cases, fibrin‐specific drugs with short half‐lives (e.g., alteplase) are preferred. Antithrombotic Strategies In PTE anticoagulants and antiplatelet agents are administered to minimize thrombus propagation, which can be stimulated by embolization, and to reduce risk of recurrence. However, the optimal antithrom­ botic strategy for small animals is not known. Some suggest that PTE should be managed with anticoagu­ lant drugs and that antiplatelet agents are not appropri­ ate. Viewing arterial and venous thrombosis as separate pathophysiologic entities is likely an oversimplification, however, and there is evidence of overlapping efficacy of antiplatelet and anticoagulant agents in the treat­ ment of both venous and arterial thromboembolic con­ ditions. Platelets are integral to the cell‐based model of hemostasis, and thrombocytosis is associated with an increased risk of PTE in people. In venous thrombosis, tissue factor‐mediated secondary coagulation precedes platelet activation, but both arms of the hemostatic sys­ tem are activated, which correlates with the architec­

tural and pathophysiologic features shared by both venous and arterial thrombi. Anticoagulants remain the mainstay of antithrombotic therapy for people with PTE, however. In people with PTE, the ACCP recommends initial parenteral antico­ agulation with low molecular weight heparin (LMWH) or fondaparinux except where renal insufficiency, throm­ bolytic therapy or poor perfusion mandate the use of IV unfractionated heparin (UFH). Ongoing anticoagulation for people with PTE is recommended for three months, although this may be extended in certain cases. Since the advent of the orally active direct Xa inhibitors and direct thrombin inhibitors, most people are now maintained on these newer medications. The direct oral anticoagulants (DOACs) have equivalent efficacy to warfarin but are associated with significantly lower bleeding risk. To date, there are no clinical trials of anticoagulants in PTE in small animals on which to base recommenda­ tions. Although there is not universal consensus on dos­ age, frequency of administration or monitoring strategy for antithrombotics in small animals, ongoing efforts by the American College of Veterinary Emergency and Critical Care led to guidelines being published in January 2019. As a result of improved understanding and the availability of the DOACs, long‐term anticoagulation in veterinary medicine is becoming increasingly common. The narrow therapeutic index of warfarin has led to increasing use of the DOACs in dogs and cats. Individual patient monitoring and dose adjustment of UFH and LMWH may enhance efficacy and improve safety and is possible using commonly available assays. Anticoagulants

Unfractionated heparin, a heterogenous mixture of poly­ saccharides, complexes with and amplifies the inhibitory activity of AT against factor IIa (thrombin) and factor Xa, although factors IXa, XIa, XIIa, and XIII are also inhibited. Thrombin inactivation prevents fibrin forma­ tion and reduces thrombin‐induced platelet activation. Variation in the size distribution of heparin molecules in UFH and unpredictable bioavailability in critical illness cause variation in heparin dose–responses, necessitating therapeutic monitoring since severe bleeding complica­ tions can result from exceeding the therapeutic range. Therapeutic monitoring can be performed using the ACT, aPTT, thrombin generation, viscoelastic testing, or anti‐FXa levels. This need for therapeutic monitoring reduces the cost benefit of UFH, but individually adjusted dosing may offer benefits over fixed dosing. Low molecular weight heparins derive from depolym­ erization of UFH and were developed to provide more consistent pharmacokinetic/pharmacodynamic (PK/PD) profiles than UFH. The reduced size of the LMWH poly­ saccharides limits their ability to simultaneously bind AT

321

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and thrombin. Thus, LMWHs are principally anti‐FXa agents and are probably best monitored with anti‐Xa assays, although tissue factor (TF)‐activated TEG can also be used. LMWHs are less protein bound than UFH, and have more predictable PK profiles and better bioa­ vailability after SC injection. The PK profiles of distinct LMWHs preparations are not interchangeable, however. LMWH PKs have been evaluated in normal healthy dogs and therapeutic anti‐Xa levels extrapolated from people are achievable. LMWH PKs in cats are less predictable, however, with frequent administration of high dosages necessary to achieve target anti‐FXa activities. A recent study evaluated the ex vivo antithrombotic effect of enoxaparin in a venous stasis model in cats, comparing it to anti‐Xa activity. The study found that at peak (but not at trough) plasma levels, enoxaparin significantly reduced thrombus formation, but in this model there was little correlation between antithrombotic effect and anti‐Xa activity. Clearly, well‐designed clinical studies evaluating LMWH in naturally occurring PTE in dogs or cats are urgently needed. Fondaparinux is a synthetic, AT‐dependent selective FXa inhibitor with no anti‐FIIa activity. In people with acute PTE, fondaparinux administration reduces PTE recurrence rate compared with UFH without increases in major bleeding, and is at least as effective as enoxapa­ rin in this setting. In dogs to date, fondaparinux has only been studied in experimental settings. In cats, a recent dose‐finding study found that SC administration of 0.06–0.20 mg/kg q12h was sufficient to achieve peak plasma anti‐Xa activity deemed therapeutic in humans. Warfarin is an oral anticoagulant that inhibits vitamin K epoxide reductase, interrupting the recycling of vita­ min K epoxide to hydroquinone and depleting the supply of carboxylated clotting factors II, VII, IX, and X. Warfarin is the mainstay for oral anticoagulation in peo­ ple but maintaining adequate anticoagulation without hemorrhagic complications is challenging. The drug has  a narrow therapeutic index, variable patient dose– response and numerous interactions with other drugs, necessitating close therapeutic monitoring. As a result, the Consensus on the Rational Use of Antithrombotics in Veterinary Critical Care (CURATIVE) guidelines pub­ lished in January 2019 advise against the use of warfarin in small animals. The difficulties associated with the clinical use of vita­ min K antagonists generated a need for effective, safe, novel oral anticoagulants. Rivaroxaban is an oral, small molecule, direct FXa inhibitor that can inhibit both free and prothrombinase complex‐associated FXa. This drug can be monitored using the PT or aPTT. Rivaroxaban was recently approved by the FDA for PTE following demonstration of equivalent efficacy to standard therapy with enoxaparin and warfarin. The pharmacokinetics of

rivaroxaban has been studied in dogs, and it is reported to be an effective anticoagulant in healthy dogs. Its clini­ cal use has been reported in a small case series, but it has not been evaluated in a randomized clinical trial. The oral prodrug dabigatran etexilate is a potent, reversible direct thrombin inhibitor. This class of drugs do not require AT as a co‐factor for their anti‐IIa activity and are also able to inhibit thrombin bound to formed clot. Dabigatran can be monitored with the PT, aPTT or the preferred ecarin clotting time (ECT) in canine plasma. Dabigatran is FDA approved for thromboembo­ lism in AF and in other countries for VTE and has a simi­ lar efficacy and safety profile to warfarin for the treatment of acute symptomatic VTE. To date, no studies have examined the effects of dabigatran in small animals. Antiplatelet Agents

In small animals, antiplatelet agents are principally used for long‐term oral maintenance therapy or as thrombo­ prophylaxis for at‐risk patients. Oral antiplatelet agents available for small animals include aspirin, ticlopidine, and clopidogrel, although concerns over myelotoxicity associated with ticlopidine have led to it being largely superseded by clopidogrel. There is limited evidence of efficacy for either aspirin or clopidogrel in veterinary medicine, however. The COX‐1 inhibitor aspirin irreversibly inhibits platelet thromboxane A2 (TXA2) synthesis, inhibiting platelet function. Aspirin is the primary antithrombotic therapy for people with atherosclerotic disease, but the populations of small animals which would most benefit from aspirin therapy are unknown. Aspirin has been rec­ ommended for dogs with protein‐losing nephropathy, nephrotic syndrome, and IMHA. The use of aspirin in IMHA is contentious, however. To date, no studies have been performed in veterinary medicine evaluating the efficacy of aspirin against objec­ tive measures of thrombosis or comparing the efficacy of aspirin against other antithrombotic therapies, but one such study is reportedly under way. The dose required to cause in vitro or ex vivo inhibition of canine platelet function varies with the assay used, but is in the range of 0.5–20 mg/kg. This wide range may be related to herita­ ble variability in canine thromboxane responsiveness. A recent study suggested that 1 mg/kg/day aspirin consist­ ently inhibited platelet function in only a third of dogs and higher dosages may be required to obtain a consist­ ent effect. The thienopyridines ticlopidine and clopidogrel are both produgs that rely on active metabolites for their in  vivo efficacy. The active metabolite of clopidogrel causes cumulative inhibition of platelet function follow­ ing repeated daily administration. In humans, clopi­ dogrel may be marginally more effective than aspirin,

32  Pulmonary Thromboembolism

while combining both agents may produce beneficial additive effects. The clinical activity and pharmacody­ namics of clopidogrel have been evaluated in cats and in dogs and significant antiplatelet effects are achievable in both species. A recent multicenter double‐blinded pro­ spective study comparing aspirin with clopidogrel in cats with ATE demonstrated superior efficacy for clopidogrel compared to aspirin, suggesting that clopidogrel should be the first‐choice antiplatelet agent in cats. The parenteral GPIIb/IIIa inhibitors abciximab, ­eptifibatide, and tirofiban, which antagonize the platelet fibrinogen/vWF receptor, have been used in canine experimental model settings of myocardial infarction and coronary stent thrombosis. Of these agents, only abciximab has been used safely in cats. Eptifibatide causes fatal cardiotoxicity in cats and should be avoided. All three drugs effectively inhibit canine platelet aggre­ gation, but none has been used in veterinary medicine. Barriers to their clinical use include cost, risk of hemor­ rhage, and the need for administration by continuous infusion.

­Conclusion Despite recent advances in human and veterinary medi­ cine, PTE remains a challenging diagnosis to confirm. A thorough, logical diagnostic approach should incorpo­ rate assessment of clinical probability, identification of risk factors and disease biomarkers, and appropriate application of diagnostic imaging, principally echocardi­ ography and multislice CTPA. Many of these diagnostic steps remain to be prospectively investigated in veteri­ nary medicine. Since PTE carries a poor prognosis and is potentially fatal, the importance of early detection cannot be over­ stated. Once we can more consistently confirm the diag­ nosis of PTE in small animals, therapeutic clinical trials of thrombolytics, anticoagulants, and antiplatelet agents will be essential in order to improve our treatment of this condition. Until such time, therapy for PTE should consist of close monitoring, good supportive therapy and judicious, individualized empiric use of thrombolyt­ ics and antithrombotic agents.

­Reference 1 Goggs R, Benigni L, Fuentes VL, Chan DL. Pulmonary

thromboembolism. J Vet Emerg Crit Care 2009; 19(1): 30–52.

­Further Reading Goggs R, Chan DL, Benigni L, et al. 2014. Comparison of computed tomography pulmonary angiography and point-of-care tests for pulmonary thromboembolism diagnosis in dogs. J Small Anim Pract. Apr; 55(4): 190–7. Jacinto AML, Ridyard AE, Aroch I, et al. 2017. Thromboembolism in Dogs with Protein-Losing Enteropathy with Non-Neoplastic Chronic Small Intestinal Disease. J Am Anim Hosp Assoc. May/Jun; 53(3):185–192.

Marschner CB, Kristensen AT, Rozanski EA, et al. 2017. Diagnosis of canine pulmonary thromboembolism by computed tomography and mathematical modelling using haemostatic and inflammatory variables. Vet J. Nov; 229:6–12. Yang VK, Cunningham SM, Rush JE, et al. 2016. The use of rivaroxaban for the treatment of thrombotic complications in four dogs. J Vet Emerg Crit Care (San Antonio). Sep; 26(5):729–36.

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33 Surgical Approaches to Thoracic Disease Raegan Wells, DVM, DACVECC Phoenix Veterinary Referral and Emergency Center, Phoenix, AZ, USA

Thoracotomy is commonly performed in critically ill small animal patients with disease of the pulmonary parenchymal structures, intrathoracic cardiovascular system, pleural space, thoracic wall, mediastinum, or esophagus. Aside from the underlying disease process, the thoracotomy procedure has the potential to induce several negative effects that can be detrimental without appropriate intensive monitoring and care. This chapter discusses the pathophysiologic effects of thoracic surgery, thoracostomy tube care, postoperative monitoring, overall case management strategies, and potential complications incurred in postoperative thoracotomy patients.

­Pathophysiology of Thoracotomy The most common surgical approaches to the thoracic cavity include intercostal thoracotomy, median sternotomy, and transdiaphragmatic incisional thoracotomy. Regardless of technical approach, some pathophysiologic consequences of entering the thoracic cavity may include atelectasis, hypoxemia, hypoventilation, hypotension, hypothermia, cardiac arrhythmias, pain, and continual pneumothorax or effusion. The four distending forces of the lungs that prevent collapse under normal conditions are transpulmonary pressure (the difference between alveolar and pleural pressure), tethering of the lungs to the surrounding structures, pulmonary surfactant, and the nitrogen skeleton. During thoracotomy, there is disruption of the natural subatmospheric pleural and transpulmonary pressure. Thoracotomy induces a positive pleural pressure which exceeds the alveolar pressure and this alteration from normal physiology rapidly leads to atelectasis. Nitrogen wash‐out occurs secondary to oxygen supplementation, which results in absorption atelectasis. This occurs as the native nitrogen skeleton is washed out by the greater than 21% FiO2, leading to a loss of the natural

nitrogen skeleton that helps maintain normal alveolar distension. It is important to emphasize that oxygen supplementation is necessary during the thoracotomy procedure, and strongly recommended in the postoperative period. Nitrogen wash‐out is, however, an important pathophysiologic contributing factor to atelectasis in the thoracotomy patient. Prolonged lateral recumbency may lead to pressure atelectasis, which results in significant low ventilation–perfusion (V–Q) mismatching and intrapulmonary shunting. Hypoxemic pulmonary vasoconstriction is the natural defense mechanism to improve V–Q mismatching under such conditions. This emphasizes the importance of body positioning and frequent rotation of recumbent patients in the immediate postoperative period. Hypoxemia and Hypoventilation Postthoracotomy hypoxemia can result from many etiologies, including hypoventilation, V–Q mismatching, and diffusion impairment. Delivery of oxygen (DO2) is the product of arterial oxygen content (CaO2) and cardiac output. In relation to hypoxemia, arterial oxygen content is dependent upon hemoglobin concentration (g/dL) and saturation as well as the partial pressure of dissolved oxygen (PaO2). Hypoventilation is defined as a PaCO2 greater than 45 mmHg in the dog or 40 mmHg in the cat. Common causes of hypoventilation in postthoracotomy patients include pain, residual anesthetics, respiratory fatigue, loss of chest wall elasticity, decreased total pulmonary compliance, increased airway resistance, and increased dead space ventilation. There is an approximate 1:1 ratio to explain hypoxemia due to hypoventilation. That is, for every 1. mmHg elevation in PaCO2, there is a corresponding 1.0 mmHg decrease in PaO2. Calculation of the alveolar to arterial oxygen gradient can be performed to rule out pulmonary pathology as the cause for hypoxemia.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Decreased transpulmonary pressure, pain, and recumbency impede proper ventilation via atelectasis and decreased tidal volume. As previously mentioned, gravitational forces may cause significant atelectasis and decreased alveolar ventilation within the dependent lung, but perfusion is maintained (or even increased) within these pulmonary segments. The overall net result in this situation is moderate to low V–Q mismatching and hypoxemia. Finally, hypoxemia following thoracotomy may also result from a combination of diffusion impairment and low V–Q mismatch secondary to reexpansion pulmonary edema or increased pulmonary vascular hydrostatic pressure. The pulmonary vasculature possesses significant permeability to protein compared to the extrapulmonary blood vessels. Since there is less intravascular oncotic pressure within this vascular circuit, elevated intrapulmonary hydrostatic pressure will favor the movement of fluid into the alveolar and interstitial space relatively quickly. Thus, pulmonary hydrostatic pressure is the main determinant for the development of pulmonary interstitial fluid accumulation and subsequent edema. Net filtration in the pleural cavity (pleural effusion) is dependent upon the capillary hydrostatic pressure of the visceral and parietal pleura, intrapleural hydrostatic pressure, plasma oncotic pressure, and intrapleural oncotic pressure. Unlike the low pulmonary interstitial to vascular space oncotic gradient, a substantial gradient is generated within the pleural space. The normal colloid osmotic pressure (COP) of the pleural space is 3.2 cmH2O (compared to 24.5–27.0 peripherally) with the average pleural pressure being –5 cmH2O.

erated idioventricular rhythm, bundle branch blocks, and ventricular tachycardia. Hypotension Postoperative hypotension is a common challenge for clinicians caring for patients following thoracotomy. Similar to the transpulmonary pressure gradient, subatmospheric pleural pressure is also important to maintain vascular distension within the thoracic cavity. During a normal breath, the elastic recoil of the lungs upon inhalation allows distension of the large intrathoracic vessels (caudal and cranial vena cava), promoting venous return and thus increasing preload. Positive intrathoracic pressure exerted during exhalation causes mild collapse of these vessels, resulting in slight changes in pulse pressure between inspiration and expiration. During mechanical ventilation, the opposite effect occurs. With positive pressure ventilation, the majority of venous return occurs during expiration, not inspiration. Anesthesia along with underlying pulmonary pathology often necessitates the use of positive end‐expiratory pressure (PEEP). Baseline PEEP is generally set at 5.0 cm H2O, but decreased pulmonary compliance and atelectasis often require higher values of PEEP. Constant, low‐grade, positive intrathoracic pressure, as PEEP increases, further impedes venous return, resulting in significant reductions in cardiac preload. Decreased preload leads to compromised stroke volume, cardiac output, and subsequent reduced delivery of oxygen. Positive pressure ventilation also has beneficial effects as it decreases afterload during inspiration, although this can be offset by dramatic decreases in preload.

Hypothermia

Pain

Hypothermia is a common occurrence during and immediately following thoracotomy. With either median sternotomy or lateral thoracotomy, the pleural (visceral and parietal) body surface area is exposed to relatively  cool ambient temperatures. This results in loss of body heat via evaporation and to some extent convection. Unimpeded hypothermia can have serious ­detrimental cardiovascular, respiratory, acid–base, and coagulopathic effects.

Postoperative pain control is of critical importance following median sternotomy or lateral thoracotomy. Both of these procedures induce pain. In addition to surgical trauma, all postthoracotomy patients have an indwelling thoracostomy tube(s) that is a frequent source of pain. Hypoventilation and hypoxemia may be a direct result of pleurodynia, incisional pain, or thoracostomy tube‐ induced discomfort. Some patients are unwilling or unable to maintain an adequate tidal volume due to painful thoracic expansion. Hypoventilation secondary to pain may also contribute to significant atelectasis resulting in hypoxemia secondary to V–Q mismatching.

Arrhythmias Cardiac arrhythmias may be observed during and subsequent to thoracotomy. Intraoperative cardiac arrhythmias may result from manual cardiac manipulation and  are typically ventricular in origin. Postoperative arrhythmias, however, are classically secondary to underlying pathophysiology including pain, hypoxemia, or physical irritation from thoracostomy tube(s). Common arrhythmias include sinus tachycardia, accel-

Continual Effusion and Pneumothorax The goal of thoracic surgery is generally to palliate or resolve the underlying disease process to improve cardiopulmonary, vascular, or esophageal function. Often, the underlying disease process may not be immediately terminated post thoracotomy, such as chylothorax, ­

33  Surgical Approaches to Thoracic Disease

­ yothorax, or pneumothorax. In such cases, thoracosp tomy tubes remain in place for several days following surgery for continual thoracic evacuation and fluid sampling. Occasionally, continual effusion or pneumothorax occurs postoperatively, exceeding expected volumes. Predictable pleural effusion secondary to an indwelling thoracostomy tube should approximate 2.0 mL/kg/ day. Fluid production above this may represent continual disease. Fluid analysis, including cell counts and cytology, can help the clinician assess if the fluid is reflective of ongoing disease. If cell counts and cytology do not support ongoing inflammation, infection or neoplasia, then it is possible that the ongoing fluid production is related to a systemic vasculitis, low oncotic pressure or elevated hydrostatic pressure. The pleural space volume is approximately 100–140 mL/kg in small animals. As such, when volumes of air or fluid exceed this value, it can be classified as a continuous process (e.g., continuous pneumothorax).

­Thoracostomy Tubes Thoracostomy tubes are a mainstay in the management of postthoracotomy patients. Depending on patient stability, a thoracostomy tube may be placed preoperatively or intraoperatively. Patients with viscous thoracic effusions or large‐volume pneumothoraces may require bilateral thoracostomy tube placement. Conventional thoracostomy tubes consist of large‐bore plastic tubing made from silicone or polyvinyl chloride. Soft, small‐bore thoracostomy tubes (e.g., Mila®, Mila International) that can be placed  using a percutaneous catheter and guidewire are preferred by many emergency and critical care clinicians. The small bore tubes can be placed with ease and minimal risk to the patient either pre or intra-operatively. The smaller diameter low profile thoracostomy tubes are effective for all disease processes, including viscous effusions such as pyothorax. A cadaver study evaluating traditional large bore chest tubes to small bore tubes in efficiency to remove air, low viscosity fluid and high viscosity fluid found that small bore tubes were as effective as large bore tubes. It is the authors’ opinion that the more traditional larger diameter tubes should be very rarely utilized, and that these tubes contribute to patient discomfort without providing any significant advantage with regards to evacuating the pleural space. 

­Postoperative Monitoring Intensive postoperative monitoring is imperative following thoracotomy. Ideal monitoring would include electrocardiography, direct arterial blood pressure

measurement, pulse oximetry, and arterial blood gas analysis, as well as end‐tidal CO2 and tidal volume in patients requiring continued mechanical ventilation. Electrocardiography Electrocardiographic monitoring provides early detection and continuous monitoring of cardiac arrhythmias. Postoperative arrhythmias may be observed primarily as a result of surgical trauma, prior manual cardiac manipulation, or direct pericardial or epicardial irritation from the indwelling thoracostomy tube. Arrhythmias may also occur secondary to pain, hypoxemia, hypovolemia, or electrolyte imbalances. Observation of hemodynamically significant postoperative atrial or ventricular arrhythmias necessitates therapeutic intervention including supplemental oxygen therapy, aggressive analgesia, correction of hypovolemia and electrolyte imbalance, and possibly antiarrhythmic medications. Diagnostic evaluation of postoperative arrhythmias may include electrolyte measurement, arterial blood gas analysis, and echocardiography. Arterial Blood Pressure As mentioned, changes in transpulmonary and pleural pressures can have significant hemodynamic effects. Direct arterial blood pressure measurement is the gold standard for monitoring patient stability and diagnosing changes in intrathoracic pressure. A sudden reduction in blood pressure may indicate an acute increase in pleural pressure, such as pneumothorax, warranting immediate investigation. Central venous pressure (CVP) monitoring can guide fluid therapy for postthoracotomy patients, but CVP is known to be affected by changes in intrathoracic pressure due to respiration, PEEP, and abdominal hypertension as each of these is known to decrease right ventricular compliance and thus artificially augment CVP. For these reasons, routine measurement of CVP in the postoperative patient with significant pleural space disease is not recommended. Pulse Oximetry and End‐Tidal CO2 Pulse oximetry can provide valuable information regarding arterial hemoglobin saturation but it can also be affected by multiple patient factors. End‐tidal CO2 (ETCO2) and tidal volume are helpful monitoring tools in the postthoracotomy patient requiring extended intubation and mechanical ventilation. Changes in ETCO2 may be the first indicator of acute deterioration such as a large pulmonary thromboembolism or cardiopulmonary arrest in these critical patients, emphasizing the importance of capnography.

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Arterial Blood Gas Analysis Arterial blood gas analysis is paramount in patients with pulmonary dysfunction, as it will provide rapid identification of hypoxemia, hypoventilation, and respiratory failure. When arterial access is unobtainable, central venous (ScvO2) or mixed venous (SvO2) oxygen saturation as well as venous CO2 (PvCO2) can be extrapolated to evaluate respiratory function. Normally, ScvO2 is 10–15% higher than SvO2 but in states of circulatory shock or failure, these two measurements closely parallel one another. The normal arterial‐venous gradient for PCO2 is 4–6 mmHg, but it may approach 10 mmHg in some patients. This allows for serial ventilatory monitoring via venous blood sampling.

­Postoperative Management Intense postoperative management following thoracic surgery comprises adequate analgesia, appropriate thoracostomy tube care, intravenous fluid therapy, and intensive nursing care. Supplemental oxygen therapy is often implemented in the immediate postoperative period and occasionally continued mechanical ventilation is required. Analgesia Optimal analgesic protocols are critical for the postthoracotomy patient to prevent secondary hypoxemia, hypoventilation, and cardiac arrhythmias. Multimodal analgesia is preferred to avoid noxious effects of single‐ agent protocols. Combination therapy of systemic analgesics allows for synergistic analgesia and lower dosing of each chosen drug. Local analgesia including intercostal nerve blocks, incisional diffusion catheters, and intrapleural infusion of local anesthetics via the thoracostomy tube provides effective adjunctive analgesia. Intercostal nerve blocks can provide very effective analgesia when utilized for lateral thoracotomy. It is recommended to perform the procedure at the intercostal space of the incision, one space cranial and one space caudal to the incision. As discussed later, much of the postoperative pain seems to be associated with large‐ bore thoracostomy tubes. Thoracostomy Tube Management and Pleural Drainage Thoracostomy tubes generally remain in place for  12–24 hours post thoracotomy in patients without  ongoing pleural space disease, but certain ­

c­ircumstances may necessitate maintenance beyond this ­ timeframe due to continual effusion, residual pneumothorax, and for therapeutic lavage or delivery of chemotherapeutics. Pleural drainage may be passive or active, but active drainage is more commonly implemented in veterinary patients. The normal transpulmonary pressure is 4–8 mmHg, which is equivocal to 5–10 mL of suction in a syringe. Intrapleural infusion, via the thoracostomy tube, of lidocaine and bupivacaine (1.0 mg/kg each) provides local analgesia following intermittent suction and may be performed every 4–6 hours. The rationale behind using both lidocaine and bupivacaine is that the lidocaine provides immediate relief and allows for a more comfortable infusion of the longer‐acting bupivacaine. This method of analgesia delivery has been investigated and deemed safe in patients following pericardectomy. Intermittent active suction is applied often postoperatively, but continuous active suction may be necessary for larger volumes of air or effusion which compromise ventilation. The pleural space in small animals can accommodate approximately 100–140 mL/kg of fluid or air. Beyond this volume, continuous pneumothorax or hydrothorax (usually hemothorax) should be suspected. Obstruction of the thoracostomy tube may rarely occur, depending on tube diameter and viscosity of the thoracic effusion. The Mac technique consists of turning the thoracostomy tube 180° in each direction and observing whether the tube returns to the prior position. This method can be used to evaluate kinking of the tube when persistent negative pressure is observed despite evidence of significant pleural effusion or pneumothorax. If the tube consistently spins back into the initial position, tube kinking should be suspected. The thoracostomy tube itself will induce approximately 2.0 mL/kg/day of pleural effusion. Once the volume of pleural effusion drops below this value, removal of the thoracostomy tube may be considered. Ideally, consistent negative pressure should be observed for 24 hours following pneumothorax prior to thoracostomy tube removal. Fluid Therapy Fluid therapy is a mainstay of postthoracotomy therapy. Most postoperative patients will require, at minimum, maintenance fluid therapy, as they may be unwilling to drink following surgery. Of further importance, continual fluid losses via the thoracostomy tube should be accounted for, and at least a percentage of this loss should be replaced with IV fluids to avoid hypovolemia due to third spacing. Postthoracotomy patients may experience significant protein loss with pleural effusion. The ideal

33  Surgical Approaches to Thoracic Disease

therapy for protein replacement in critically ill patients is to provide enteral nutrition in order for the patient to synthesize albumin. Oxygen Therapy Oxygen therapy may be provided intermittently via facemask or continuously by nasopharyngeal insufflation, nasal prongs, oxygen cage, or intubation and mechanical ventilation. Provision of intermittent supplemental oxygen at an FiO2 of 50–60% can be achieved via facemask with oxygen flow rates of 8.0–12.0 L/min. Nasopharyngeal insufflation or nasal prongs may be utilized with oxygen flow rates of 50–150 mL/kg/min to achieve an FiO2 of 30–70%. More commonly, postoperative thoracotomy patients are maintained in an oxygen cage with flow rates of 0.5–1.0 L/min to maintain a FiO2 of 40–60%. Patients requiring an FiO2 above 60% to maintain normoxemia will likely require intubation and mechanical ventilation. Caution must be exercised to avoid oxygen toxicity when supplemental oxygen levels are maintained above 60% for more than 24 hours. When mechanical ventilation is utilized, PEEP ventilation can be beneficial to prevent atelectasis and augment work of breathing. In contrast to the spontaneously breathing patient, permissive hypercapnia is tolerated in mechanically ventilated postthoracotomy patients. Arterial PCO2 may rise to 50 mmHg before intervention is necessary, as long as normoxemia is maintained. Proper humidification is an important component of oxygen therapy for preservation of proper mucociliary function and clearance of respiratory secretions. Dyspnea Dyspnea is the perceived feeling of breathlessness. The pathophysiology of dyspnea is multifactorial and is described elsewhere. Multiple therapeutic targets have been investigated to decrease the perception of dyspnea in human patients. In human studies, nebulized or aerosolized furosemide has been found to prevent bronchoconstriction and inhibit the cough reflex in addition to reducing pulmonary vascular hydrostatic pressure and edema when absorbed systemically. Furosemide has been shown experimentally and clinically in people to reduce bronchial slow‐acting receptors (SARs), thereby reducing the sensation of dyspnea. Furosemide exerts a diuretic effect when administered by the enteral or parenteral route via inhibition of the Na:K:2Cl co‐transporter within the thick ascending loop of Henle. A similar mechanism is postulated for the antidyspneic effect of nebulized furosemide causing a local increase in

sodium ion concentration within the airways, increasing pulmonary stretch receptor activity. Hypothermia Hypothermia is an inevitable occurrence during and immediately following thoracic surgery. Warming IV fluids prior to administration can help diminish large  reductions in intraoperative body temperature. Postoperative temperature monitoring, either intermittent or continuous, is recommended to ensure euthermia and active warming using warming blankets or heat lamps should be performed until the patient’s temperature has reached 99.5 °F. Arrhythmias For hemodynamically significant, multifocal, sustained, or R‐on‐T phenomena ventricular arrhythmias, lidocaine remains the mainstay therapy. Lidocaine may also be useful to slow the heart rate, allowing differentiation of ventricular tachycardia from sinus tachycardia with bundle branch block. The waveform of the latter often will not change following lidocaine bolus (2.0 mg/kg IV), but P‐waves may be appreciable once the heart rate has decelerated. For lidocaine‐responsive arrhythmias, a constant rate infusion of 60–80 μg/ kg/min may be implemented postoperatively. A beta‐ blocker may also be ­considered for patients that do not have known underlying cardiac disease with compromised contractility. Anecdotally, sotalol 2.0 mg/kg PO BID has favorable postoperative antiarrhythmic ­properties for patients with supraventricular and ventricular tachyarrhythmias. Nursing Care Nursing care is a critical yet often underemphasized aspect of postthoracotomy care. Patients undergoing median sternotomy often prefer lateral recumbency in the immediate postoperative period due to incisional pain. Turning or rotating the patient every 4–6 hours prevents atelectasis and V–Q mismatching secondary to gravitational forces in lateral recumbency. Turning the patient also promotes patient comfort and hygiene. Optimal analgesia aids in patient well‐being and may promote tolerance of sternal recumbency and therefore improved oxygenation and ventilation. Complications Complications post thoracotomy may include edema formation, hypoproteinemia, continual pleural effusion,

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secondary infection, hemorrhage, residual or continuous pneumothorax, seroma formation, sternal osteomyelitis, pyrexia, iatrogenic chylothorax, fibrosing pleuritis, surgical dehiscence, and ipsilateral thoracic limb lameness (lateral thoracotomy). Crystalloid fluid therapy, anorexia, vasculitis, and postoperative inflammation can contribute to peripheral edema formation. The most common complication involving the thoracostomy tube is improper placement. Insufficient subcutaneous tunnel formation with large‐bore thoracostomy tubes can lead to continuous air leakage and pneumothorax. The thoracostomy tube percutaneous insertion site should be inspected carefully when continuous pneumothorax is observed. Continual pleural effusion may be primary or secondary in origin, depending on the etiology requiring thoracotomy. Pyothorax, chylothorax, and neoplastic effusion represent sources of continual pleural effusion during the postoperative period. Pleural drainage is important to maintain proper ventilatory function as well as oxygenation. However, excessive pleural effusion can lead to significant systemic protein and fluid loss with subsequent intravascular volume depletion. Vasculitis and surgical inflammation are contributors to this protein‐losing pleuropathy and third spacing, which may encourage perpetual effusion. Resolution of the underlying disease process will likely terminate the cause of effusion. Infection is an undesirable complication in any postoperative patient. Preoperative infection may be present in some patients, such as pyothoraces, in which septic effusion necessitated surgical intervention. Continual monitoring of thoracic fluid cytology and cell counts aids in determining resolution of pyothorax. Responsible antimicrobial administration is of paramount importance in the care of critically ill patients. A full discussion on this topic can be found elsewhere. Generally speaking, clinicians should be familiar with antibiotic sensitivity patterns of common bacterial isolates from their hospitals and deescalate coverage to single‐agent therapy as soon as possible. Infection may be introduced intrathoracically via the thoracostomy tube or surgical incision. This may be due to bacterial contamination associated with prolonged surgical exposure, improper handling of the thoracostomy tube, or bacterial invasion of the thoracostomy tube insertion site and/or surgical incision. Appropriate samples should be obtained for bacterial culture and antimicrobial susceptibility testing when ­ infection is suspected. ­ Proper intraoperative hemostasis ensures minimal postoperative hemorrhage, therefore significant hemorrhagic pleural effusion is not expected in the ­

­ ostthoracotomy patient. Serosanguinous pleural effup sion (PCV 90 mmHg for optimal organ and tissue perfusion. Hypotension that is nonresponsive to fluid therapy may indicate the presence of an underlying acid–base or electrolyte disorder or the need to begin therapy with a vasoactive agent. Central venous pressure and lactate monitoring can provide additional information on the need for vasopressor therapy. While less commonly encountered clinically, monitoring for and treatment of hypertension are useful to prevent complications including retinal detachment, neurologic impairment, and end‐organ injury. Cardiac Function Heart function should be evaluated not only by heart rate and rhythm but also ability to effectively pump blood forward through evaluation of peripheral limb temperatures and capillary refill time. Electrocardiographically confirmed arrhythmias should be treated if perfusion is compromised. Echocardiography is helpful in identifying underlying structural heart disease as well as visualizing cardiac filling and contractility. Additional treatments directed at optimizing cardiac function, including IV ­fluids, inotropic support or afterload reduction, should be introduced as indicated. Albumin Albumin is a blood protein produced by the liver that plays an essential role in a multitude of physiologic processes, including fluid balance, drug transport, tissue healing, and coagulation. Synthetic colloids can be used to maintain fluid in the intravascular space but they do not substitute for albumin’s other roles in the body and may be associated with acute kidney injury. Albumin levels 70% is not maintained despite supplemental oxygen and normalization of intravascular volume and systemic blood pressure. Renal Function and Urine Output Kidney function should be frequently evaluated in critically ill patients as hypotension, thrombosis, and nephrotoxic medications can cause temporary or permanent injury resulting in renal insufficiency. Urine output should be at least 1 mL/kg/h but oliguria should be evaluated in light of fluid input for an individual patient. An indwelling urinary catheter, and recording fluid in and fluid out, assists in evaluating a patient’s overall fluid status. In addition to urine output, insensible losses, losses from surgical drains, chest tubes or wounds should also be accounted for. Commonly accepted insensible loss rates are 10–20 mL/kg/day. Fluid input includes IV fluids (crystalloids and colloids) as well as nutritional support in the form of parenteral or enteral nutrition. In patients with possible renal impairment, close monitoring of fluid ins/outs may assist in maintaining fluid balance. Daily monitoring of serum BUN and creatinine is recommended to detect potential renal injury early. Especially in patients with normal renal values at the time of presentation, even mild changes in BUN or creatinine (even within the reference range) can be a harbinger of renal injury or multiorgan failure. Early detection and aggressive treatment can have a dramatic impact on patient outcome. Analyzing urine for casts, glucose, and protein is also used to identify renal injury and dysfunction. Immune Status and Antibiotic Coverage Critically ill patients are often battling infections and may have overextended or overwhelmed immune systems. Empirical antibiotic therapy is recommended for patients with suspected infections and antibiotic selection should be based on presumed site of infection, likely organism involved, and hospital surveillance profiles. When empirical antibiotics are used, initial broad‐spectrum coverage is recommended with deescalation based on results of culture and sensitivity panels. In addition to bacterial infections, clinicians should be aware of the concern for rickettsial, protozoal, fungal, and parasitic infections that may require additional antimicrobial therapies. In all critical patients, care should be taken to practice aseptic techniques when examining and treating them. Clinicians and nurses should practice good hygiene with frequent hand washing. Exam gloves should be worn at all times when handling any patient and the use

of isolation gowns should be considered if the patient is severely immunocompromised. Gastrointestinal (GI) Motility and Integrity Many underlying conditions can result in gastric and intestinal ileus and compromise of the GI mucosal barrier. Decreased intestinal motility results in stasis of intestinal contents and overgrowth of normal bacterial flora. An altered mucosal barrier results in an increased risk for bacterial translocation and subsequent sepsis. Gastric and small intestinal stasis may result in vomiting, while large intestinal derangements may lead to diarrhea, which can create and/or exacerbate hypovolemia. Medical intervention (promotility medications, physical activity) to increase GI motility and protect the mucosal barrier is recommended. If possible, enteral nutrition is recommended to maintain enterocyte health, decrease villous atrophy, increase GI motility, and improve mucosal integrity. Drug Metabolism and Dosage Medication doses and routes should be reviewed daily for accuracy, safety (side‐effects, metabolism, and drug interactions) and continued necessity. In the face of altered renal or hepatic function, it may be prudent to reduce the dose or increase the interval of medications to decrease the risk of toxicity and side‐effects. Highly protein‐bound drugs should be used with caution in hypoalbuminemic patients due to the potential for higher than expected plasma levels. Nutrition All attempts should be made to provide appropriate nutritional support to critical patients in order to avoid a negative energy balance contributing to organ dysfunction, delayed healing, immunocompromise, and GI ­dysfunction. Enteral nutrition is ideal and can be accomplished by voluntary patient eating or feeding tubes (e.g., nasogastric, esophageal, gastrostomy or jejunostomy tubes). Total or partial parenteral nutrition is an option for providing nutritional support to patients with a contraindication to enteral nutrition (severe ileus, intractable vomiting, malabsorptive conditions). Analgesia Appropriate pain management is necessary to improve patient comfort and decrease the secondary effects of pain. Common signs of pain include increased heart rate, agitation, and discomfort upon palpation of wounds. Multiple pain scoring systems have been validated for

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use in veterinary medicine and provide a systemic and repeatable approach to evaluating pain and a patient’s response to pain medications. Nursing Care and Patient Mobility While focusing on the medical care of a critically ill patient, it is easy to overlook patient comfort and husbandry. In patients that require extensive nursing ­ care, these measures may be skipped due to the time and staffing they require, especially in larger patients that may require multiple people to move and support them. Many of these patients are nonambulatory and physical therapy should be provided several times a day. Passive range of motion of limbs keeps muscles and joints flexible and can improve circulation. For patients with peripheral limb edema, tissue massage from distal to proximal can assist in resolution. Deep bedding and frequent repositioning are necessary to prevent pressure sores, lung atelectasis, and edema formation. Standing sessions are useful and allow patients to attempt supported ambulation. If possible, underwater treadmill therapy is an option for assisted mobility to increase flexibility, strength, and circulation. Efforts should be made to frequently take nonambulatory patients out of their cages and, if feasible, outside for a “visual vacation.” For patients with lung consolidation from pneumonia or other conditions, positional coupage based on afflicted region can be useful in resolving consolidation. In patients with surgical incisions or traumatic tissue injury, cold laser therapy can be considered to improve tissue blood supply and increase healing. Patients should be diligently cleaned to prevent skin irritation and burns from urine and feces soiling.

peripheral edema can rapidly develop and cause wraps or bandages to become tight and constricting. Soiled or wet bandages should be immediately changed as they provide an excellent source of infection. Wounds and incisions should be closely monitored for healing and any changes that occur, including bruising, odor, swelling, and discharge, should be immediately investigated. Tender Loving Care Amongst all the medications, procedures, and treatments, time should be taken to simply sit with a patient to provide comfort and affection. Once again, this is an area that is easy to omit when a patient has intense medical treatments and support staff is spread thin. Patients that require many treatments are often mislabeled as being “difficult cases” rather than appreciated for the multiple levels of complex care they require to support their severe disease states. Many times patients will only eat when being petted. Owner visits should be encouraged and efforts made to allow visits to occur outside the hospitalization area to allow patients stress‐free periods during which they may eat or rest. In 24‐hour care facilities, efforts should be made to allow the patients to have treatment‐free periods during which uninterrupted rest can occur. Clinicians should attempt to arrange treatments in groupings to establish greater intervals of time when the patient is not being handled or interacted with. Especially during evening hours, further emphasis should be placed on treatment‐free periods. During these evening times, lights should be dimmed and attempts made to decrease ambient noise. There may be benefit to playing relaxing music such as soft classical or nature sounds.

Bandage and Wound Care All IV catheters should be routinely changed and catheter insertion sites evaluated for infection and inflammation. Each facility should establish a protocol for the placement and maintenance of peripheral IV catheters and advanced catheters such as central IV catheters and arterial catheters. At a minimum, daily bandage changes should be required but other considerations include acceptable indwelling duration, flushing/locking procedures, level of sterility during placement and handling, and a protocol for investigation and treatment of suspected catheter site infections. Indwelling urinary ­ catheters also require set protocols for placement and maintenance including prepuce/vulva flushing, collection system handling/changing, acceptable indwelling duration, and a protocol for investigation of suspected infections due to catheterization. Bandages placed on wounds require frequent changing to evaluate for changes in underlying wounds. During critical illness,

­Euthanasia For many critical patients, compassionate euthanasia may be the most humane treatment available. As a case progresses, lines of communication regarding euthanasia should remain open between owners and the medical staff. Despite being common in veterinary medicine, some owners may not know it is an option for their pets. Broaching the subject of euthanasia can be difficult and needs to be handled with special attention. Even in cases with a grave or guarded prognosis, owners may not ­consider euthanasia due to religious or ethical beliefs.

­Conclusion Each critical patient will require a unique and customized approach to every element of their care. While the

35  Approach to the Patient in the Critical Care Setting

care for these cases is more complicated, with it comes a greater feeling of success when a patient is discharged to its family. Many critically ill patients will not have a positive outcome and often the deciding factor in outcome is determined by the attention provided to the patient and owner. It is easy to influence a client to terminate care for a patient with a guarded prognosis, but experience and

clinical knowledge should be used to assist clients in making the best decision for them and their pet. The care of critical patients can be exhausting for the entire medical team. Clinicians should take the time to frequently discuss the details of these cases with support staff to ensure that they understand the importance and motivation behind medical treatments and owners’ decisions.

­Further Reading Downing R. The role of physical medicine and rehabilitation for patients in palliative and hospice care. Vet Clin North Am Small Anim Pract 2011; 41(3): 591–608. Hackett T. Physical examination. In: Silverstein D, Hopper K, eds. Small Animal Critical Care Medicine. St Louis, MO: Saunders Elsevier, 2008, pp. 2–5. Hopper K, Powell L. Basics of mechanical ventilation for dogs and cats. Vet Clin North Am Small Anim Pract 2013; 43(4): 955–69.

Pachtinger G. Monitoring of the emergent small animal patient. Vet Clin North Am Small Anim Pract 2013; 43(4):705–20. Prittie J. Optimal endpoints of resuscitation and early goal directed therapy. J Vet Emerg Crit Care 2006; 16(4): 329–39. Rule of 20. http://files.dvm360.com/alfresco_images/ DVM360//2013/11/11/b67d4ee5‐e8cd‐4002‐8d45‐ 39e6ce9bd68f/article‐804765.pdf (accessed May 22, 2019).

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36 Fluid Therapy Teresa Rieser, DVM, DACVECC Veterinary Specialty Care, Mt. Pleasant, SC, USA

Intravenous fluids are a commonly used treatment in small animal medicine. A full understanding of the indications and potential fluid choices allows clinicians to reap the maximum benefit from fluid therapy while minimizing possible complications.

­Vascular Access Catheter Types Fluid therapy can be administered via a number of different routes, including intravenous, subcutaneous, and intraosseous. In order to administer fluids intravenously, venous access must first be established. Most commonly, this involves the placement of a catheter within a peripheral vein or within the jugular vein. There are a variety of catheter types currently available, including winged needle (butterfly catheters), over the needle, through the needle and catheters that are placed using a guide wire. When placing an intravenous catheter, a number of questions should be considered to ensure the most appropriate catheter is selected. Butterfly‐type catheters are rigid needles with a plastic “butterfly” at the needle hub to make handling easier. The needles are then attached to a length of tubing that can be connected to a syringe for bolus drug delivery or blood sample collection. This type of catheter is useful for instances where the catheter does not need to be maintained within the vein. An example would be an IV bolus injection of medication or a single episode of venipuncture to collect blood samples. If a catheter needs to be maintained within a peripheral vein for any duration of time then an over‐the‐needle type catheter is most commonly used. Theses catheters are relatively easy to place and can be maintained within a vessel for a number of days with diligent nursing care. If placed in the critically ill, emergent or surgical patient, the short length of the catheter coupled with a reasonable

diameter allows for rapid bolusing of fluids. This type of catheter is usually secured in place with medical tape. Through‐the‐needle catheters are also widely available. This type of catheter tends to be greater in length and is used when central venous access is desired for repeated blood sampling or the administration of a hyperosmolar solution such as total parenteral nutrition. Catheters placed using a guide wire can be single lumen or multilumen and are placed into a central vein. This type of ­catheter allows for blood sampling, fluid administration, including the concurrent administration of noncompatible fluids through separate lumens, as well as the administration of hyperosmolar solutions that would cause significant phlebitis if administered peripherally. Placement of a central venous catheter also allows for monitoring of central venous pressure in patients to guide fluid therapy. Intravenous catheters may be made of a number of different materials, including polyurethane, Teflon®, and silicon elastomer (silastic). Polyurethane and Teflon catheters are most commonly used, as they are cost‐ effective and relatively easy to insert. Silicon elastomer (Silastic®) catheters are very flexible and less likely to result in catheter‐related thrombophlebitis but they are more expensive and technically challenging to place. Silastic catheters are used for long‐term catheterization, while Teflon or polyurethane catheters are reasonable choices for shorter term catheterization. The placement of intravenous catheters is common in small animal veterinary medicine. Peripheral catheters are most commonly placed in the cephalic, medial or lateral saphenous or femoral vein. In some animals, the veins of the ear can also be used for catheterization. Central catheters are placed within the jugular vein or threaded from a peripheral vein into the vena cava. In order to preserve the integrity of the catheter and minimize discomfort and inflammation, catheters should not cross a joint. Common sense would also dictate that the

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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patient’s disease process might make one catheterization site preferred over another. An example is the parvoviral enteritis patient with voluminous diarrhea; a hind leg catheter should not be the first choice, as it would probably become soiled in short order. The area selected for placement should be clipped and aseptically prepared, taking care not to abrade the skin, as this can increase the rate of catheter‐related complications. In addition to preparing the patient, staff members should also wash their hands and wear gloves. Once the catheter is seated within the vein, it is flushed with sterile saline. There has been no proven benefit to using heparinized saline in lieu of sterile saline as a catheter flush. Peripheral catheters are secured to the limb with medical tape. It is common to suture central lines in place and then to protect them further with a bandage. Catheter sites should be kept clean and dry and inspected at least daily for evidence of inflammation. Catheter‐Related Complications Catheter‐related complications are common but can be minimized with relative ease. Any material used for an intravenous catheter will cause some inflammation within a vessel so it is important to diligently assess catheter sites for indication of not only infection but also thrombophlebitis. Catheter sites should be inspected daily for swelling, pain, or induration of the vessel. An indurated vessel will feel firm and ropey when palpated. It is also important to monitor patient response when infusing medications. As catheter integrity degrades, the animal may object when drugs are infused. The possibility of infection or thrombophlebitis should be suspected in any patient who has an intravenous catheter and develops signs of infection or exhibits a reaction to injections that were previously well tolerated. A patient who develops a low‐grade fever in hospital without another potential nidus of infection or inflammation should have all catheter sites promptly inspected. While in the past it has been recommended that catheters be removed after 72 hours to prevent infection, multiple studies have shown that the rate of infection does not increase if catheters are allowed to remain in place for longer periods of time and that it is more important to keep the area clean and the catheter dressing unsoiled. Infiltration occurs when a catheter moves out of a vein into the surrounding tissue. Any fluids or drugs administered through the catheter are therefore infiltrated into that area instead of being delivered to the vein. Most commonly, infiltration is identified by swelling and ­tenderness at the catheter site. If vesicant medications are administered perivascularly, the term used is extravasation. The sequelae of extravasation can be more severe and may include heat, pain, redness, swelling, and

i­nduration. Sloughing of the skin and the perivascular tissue can then follow. If the catheter is a central line, it may not be immediately evident that the position of the catheter has changed. Infiltration of fluid in this instance can result in mediastinal or pleural fluid accumulation that can be severe enough to cause dyspnea. The key to preventing catheter‐related complications is vigilance. Place catheters away from joints and avoid using rigid materials, like a butterfly catheter, for prolonged infusion. Inspect the catheter site frequently. Before infusing drugs, aspirate the catheter for blood (a “flash”) and then flush with saline to ensure proper placement and patency of the catheter. Catheter foreign bodies occur when a portion of the catheter becomes free within a vessel and lodges (most commonly) in the heart or a pulmonary artery. Catheter foreign bodies may occur when catheters are accidently cut during removal or when a through‐the‐needle catheter is cut off by the introducer needle. It can also occur when an over‐the‐needle catheter is partially advanced and then replaced on the needle while seated within a vein. Infrequently, the shaft of a catheter can become disconnected from the catheter hub. These types of ­ ­mishaps can be associated with serious complications, including cardiac perforation, endocarditis, pulmonary embolism, sepsis, and arrhythmias. Options for management of catheter foreign bodies include surgical retrieval, endovascular retrieval, or leaving the catheter in situ. In human medicine, attempted retrieval of the foreign body is preferred. Successful retrieval of intravascular foreign bodies using endovascular techniques has been reported in five dogs, a goat, and a horse. Air emboli occur when air is introduced into the vascular system through a catheter. This can be seen when a central line is advanced into the thoracic cavity and the change in pressure draws air into the system. More commonly, air can be introduced through incorrectly prepared IV lines or when air is injected into rigid fluid containers to speed an infusion. Small emboli are usually clinically silent. Large air emboli can result in a dramatic increase in pulmonary vascular resistance followed by pulmonary edema formation and dyspnea. If a large enough bolus of air is delivered, the right ventricular outflow tract can be obstructed. If a central line is in place when an air emboli occurs, aspirating the air out of the heart is the best choice for treatment.

­Body Fluid Compartments Body Water Distribution Approximately 60% of body weight is water. This ­percentage can change based on age, gender or body

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to dilution. Finally, as the hydrostatic pressure increases in the interstitium, lymphatic drainage is increased. If increased capillary hydrostatic pressure persists, however, these safety factors can be overwhelmed and the meshwork within the interstitium can be disrupted. When this occurs, the resistance to further fluid accumulation becomes minimal and manifests clinically as pitting edema, chemosis, and, in rare circumstances, pulmonary edema. Fluid dynamics in pulmonary tissue have unique characteristics. The pulmonary capillary pressure is usually lower than in other tissues and the interstitial pressure is more negative (some tissues actually have a negative interstitial pressure) while the COP is higher than other tissues. The fact that the pulmonary capillaries are relatively permeable to protein contributes to this higher COP. In addition, the alveolar epithelial cells actively transport sodium into the interstitial space, promoting movement of fluid out of the alveolar space and into the interstitium. The net result of these characteristics is that fluid movement from the alveoli into the interstitium and then to the lymphatics is favored. This tendency is overcome when the pulmonary capillary pressure exceeds 25 mmHg, at which point fluid will accumulate within the alveoli. If the animal also has an increase in pulmonary capillary permeability such as occurs in sepsis or systemic inflammatory response syndrome (SIRS), or if the animal is hypoalbuminemic, pulmonary edema will occur at lower hydrostatic pressures.

­Intravenous Fluid Types In order to make logical choices with fluid therapy, it is important to understand the components and behavior of the different classes of intravenous fluids available in clinical practice. Crystalloid Fluids Crystalloid fluids are the most common type of fluids used in veterinary medicine. Crystalloids are defined as fluids containing electrolyte and nonelectrolyte solutes that are capable of entering all the body fluid compartments. They can further be described as “balanced” or “unbalanced.” A balanced crystalloid has a composition similar to that of the ECF. An example would be lactated Ringer’s solution (LRS) or Normosol‐R®. An unbalanced crystalloid’s composition does not mimic that of the ECF. An example would be 0.9% NaCl. Crystalloids can also be described by their osmolarity. An isotonic crystalloid has an osmolarity similar to normal plasma osmolarity (300–310 mOsm/L) while the osmolarity of a hypotonic solution is less than that of

normal plasma and a hypertonic solution is greater than normal plasma. An example of this can be seen with the different sodium chloride‐based crystalloids: 0.9%NaCl is an isotonic crystalloid solution (308 mOsm/L), 0.45% NaCl is a hypotonic crystalloid (154 mOsm/L), and 7.2% NaCl is a hypertonic crystalloid (2464 mOsm/L). Finally, crystalloids can be described as replacement fluids or maintenance fluids. The composition of these two categories is quite different and warrants a brief explanation. When a crystalloid fluid is administered intravenously, the majority of the volume given will move out of the vascular space and into the interstitium. When a 1 L bolus of an isotonic crystalloid is given, it can be anticipated that after one hour only 25% of the delivered volume remains within the vascular space. The majority of the delivered volume has moved into the interstitium and for this reason, a replacement fluid is designed to approximate the fluid composition of the ECF. By contrast, a maintenance fluid is designed to meet the ongoing insensible losses of the patient and has a lower sodium concentration and usually a higher potassium concentration. A unique crystalloid fluid that does not fall within the replacement versus maintenance classification scheme is 5% dextrose in water (D5W) that is composed entirely of dextrose and water. When given, D5W is effectively equivalent to administering pure water, as the dextrose is rapidly metabolized to CO2 and water. Specific clinical presentations may respond most effectively to specific crystalloid choices but in the majority of instances, a balanced, isotonic crystalloid is appropriate for initial fluid therapy. Some of the clinical presentations where one fluid may be preferred over another include hypochloridemic metabolic alkalosis and Addisonian crisis (0.9% NaCl) and urethral obstruction (LRS or Normosol‐R), among others. Additives such as 50% dextrose, potassium chloride, magnesium sulfate, and calcium gluconate can be incorporated into crystalloid fluids to meet specific patient needs; however, not all fluid additives are compatible with all isotonic crystalloids. Investigation into compatibility is required before fluid supplementation is undertaken. Certain crystalloids such as 0.9% NaCl and D5W are often used as carriers for a variety of drugs, including anesthetics, inotropes, vasopressors, and insulin, but compatibility should always be verified before administration. Hypertonic saline is available in a variety of concentrations ranging from 3.0% to 7.5% and is a potent expander of the vascular volume. With an osmolarity of 2567 mOsm/L, 7.5% NaCl will effectively increase the extracellular fluid volume by five times the administered volume due to the movement of water from the intracellular and interstitial spaces into the intravascular space

36  Fluid Therapy

in response to the profound osmotic gradient created. However, like other crystalloid solutions, the intravascular volume expansion will not remain for long if administered alone. For this reason, hypertonic saline is often combined with a synthetic colloid to extend the duration of effect. Because the action of hypertonic saline is dependent on movement of volume from the interstitial space to the intravascular space, it is relatively contraindicated in dehydrated patients. Hypertonic saline is a popular choice for small‐volume resuscitation and is the preferred choice for the resuscitation of the hypotensive head trauma patient. In addition, it may also be used in patients with intracranial hypertension with, or in lieu of, mannitol. Synthetic Colloids Colloids are molecules of high molecular weight, which exert osmotic activity and thus promote the movement of fluid from the interstitium into the vascular space. Colloids can first be described as natural or synthetic. An example of a natural colloid is the albumin molecule. Synthetic colloids can further be divided into three classes: gelatins, dextrans, and hydroxyethyl starches (HES). The most commonly used class of synthetic colloid is the hydroxyethyl starches, which are plant‐derived proteins that have undergone modification to prevent rapid degradation. When describing hydroxyethyl starches, a number of parameters are used. First is the molecular weight, which affects the degree of “oncotic pull” exerted by the solution, with lower molecular weight solutions generally exerting a greater effect. Molecules in HES formulations tend to be polydisperse and can range in size from a few thousand to a few million kilodaltons. The distribution within a solution follows a bell‐shaped curve. By convention, the molecular weight used to describe the solution is the average molecular weight. The second parameter used in description of HES solutions is the degree of substitution, determined by measuring the number of substituted glucose molecules and dividing by the total number of glucose molecules. HES solutions with a higher rate of substitution resist hydrolysis more effectively than molecules with lower substitution rates and thus persist in the body for a longer period of time, extending duration of action. The rate of substitution is used to describe the fluid and while it is common in veterinary medicine to refer to all hydroxyethyl starches as “hetastarch,” this is a misnomer; in fact, a substitution rate of 0.6 is termed a hetastarch, 0.5 is a pentastarch, and 0.4 is a tetrastarch. The final parameter that can be considered when evaluating HES solutions is the carrier solution. Classically, the majority of HES solutions are provided in 0.9% NaCl.

Some solutions are also available in balanced crystalloid solutions such as LRS. The use of synthetic colloids is somewhat controversial. Studies in both the human and veterinary literature have shown impaired coagulation and decreased platelet clump strength as determined by thromboelastography. The higher the rate of substitution, the greater effect on coagulation. For this reason, HES solutions with a lower rate of substitution such as the tetrastarches are considered theoretically safer as their effect on coagulation is less. Overzealous administration of any of the HES s­ olutions can still result in a dilutional coagulopathy. In addition to effect on coagulation, synthetic colloids can impact serum total protein and urine specific gravity (USG) measurements. The administration of HES (670/0.75) has been previously shown to elevate the urine specific gravity. In one study, the largest increase in USG occurred 150 minutes following administration with a mean USG of 1.070 +/− 0.021. Both hydroxyethyl starch and dextran 70 have been shown to affect total solids measured by refractometry. When evaluated using a refractometer, HES will yield a reading of 4.5 g/dL, therefore when administered in vivo, the refractometric measurement of total solids will trend to 4.5 g/dL. For this reason, total solids should be interpreted with caution in patients receiving synthetic ­colloids. In addition, serum amylase may be elevated 200–250% due to binding with hydroxyethyl starch ­molecules, leading to a decrease in clearance. Ideally, the efficacy of synthetic colloids is monitored through serial measurement of the COP using a membrane osmometer. In practice, this technology is not widely available and the response to therapy is used instead to guide treatment. Finally, in human medicine, in particular in the subset of patients with refractory sepsis, concerns have been raised about HES administration and renal injury based on an increase in the need for renal replacement therapy in patients receiving hydroxyethyl starches. At this time, this concern appears to be related to the cumulative dose of HES administered. There have been some studies examining the issue in small animal medicine but the topic remains controversial. Dextrans are neutral glucopolysaccharides based on glucose monomers. Dextrans are described based on their average mean molecular weight as well as the tonicity of the fluid. Examples of dextrans include products such as dextran 40, dextran 60, and dextran 70. In humans, dextran administration has been associated with anaphylactoid reactions as well as being implicated in the development of renal failure in critically ill patients. This has not been documented in veterinary patients.

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The last class of synthetic colloid is the gelatins  – polydisperse molecules derived from bovine collagen. These solutions have a mean molecular weight of 35 000 daltons and are rapidly excreted upon infusion. As a result, they are far less effective than the hydroxyethyl starches or dextrans for sustained volume expansion. In addition, the gelatins are associated with the highest rate of anaphylactoid reactions among the synthetic colloids. As a class, gelatins are not available in the United States. Albumin Physiologically speaking, albumin is an extremely important protein. It contributes roughly 80% of the COP within the vascular space. In addition to its osmotic activity, albumin has many other roles in the body and functions as a carrier for a variety of substances, including bilirubin, as well as many different drugs. Albumin also plays a role in platelet function and in the scavenging of free radicals. The ability of albumin to create colloid osmotic pressure is enhanced by its negative ­ charge as well as the Gibbs–Donnan effect that describes how charged molecules may not be equally dispersed across a semipermeable membrane due to the relative ­permeability of the membrane.

­Indications for Fluid Therapy Rapid Volume Expansion The rapid administration of IV fluids is indicated for the treatment of hypoperfusion. Rapid volume expansion is not appropriate if hypotension is the result of forward cardiac failure for any reason other than cardiac tamponade. The goal of rapid volume expansion is to restore the effective circulating volume (ECV) to levels that permit adequate perfusion and thus tissue oxygenation. Serial monitoring of perfusion parameters is used to guide therapy during rapid volume expansion. Parameters that are commonly monitored include physical examination parameters such as heart rate, pulse rate, pulse quality, mucous membrane color, and capillary refill time (CRT). In addition, monitoring may also include toe‐web to core body temperature gradient, urine output, central venous pressure, and blood lactate measurements. A full discussion of end‐points of resuscitation can be found in Chapter 37. The dosing of IV fluids for the treatment of shock varies with the type of fluid chosen for resuscitation as well as the patient’s response to therapy. The most common fluid types used for rapid volume expansion are isotonic crystalloids. Classically, a full “shock” bolus is considered to be 90 mL/kg in the dog and 45–60 mL/kg in the cat. These amounts approximate one blood volume. Boluses

can be administered over a specific amount of time (for example, a 250 mL bolus of 0.9%NaCl over 10 minutes), or in profoundly compromised patients as rapidly as possible. Frequently, one‐quarter of the calculated shock bolus is administered as a starting point for resuscitation. After the bolus is complete, the animal is reassessed. If their perfusion parameters have not responded adequately, additional fluid therapy is indicated. It should be remembered that when using an isotonic crystalloid for rapid volume expansion, that crystalloid should be devoid of fluid additives such as supplemental dextrose, potassium chloride or magnesium sulfate. Synthetic colloids such as Hespan® or Vetstarch® can also be used for rapid volume expansion. Because synthetic colloids are osmotically active, they promote the movement of water from the interstitium into the vascular space. This movement of fluid between body fluid compartments as well as the volume of colloid administered collectively results in the expansion of the ECV. An initial shock bolus of synthetic colloid would be 5 mL/kg administered over 10–15 minutes.The total shock dose is equal to the recommended total daily dose of 20 mL/kg in the dog. In the cat, the total daily dose is somewhat lower at 10 mL/kg. Rapid administration of hydroxyethyl starch has been associated with nausea and vomiting in the cat. Again, reassessment of the patient following completion of the bolus will guide further fluid therapy. If the response to therapy has not been adequate, additional fluids should be administered until the animal has been stabilized. If the animal is unstable but is deemed to be volume replete, other therapies such as inotropes or vasopressors may be needed. It should be remembered that synthetic colloids are potent osmotic agents so overzealous administration can result in volume overload as well as dilutional coagulopathy. Hypertonic saline can also be used for rapid volume expansion. Hypertonic saline is commonly supplied as a 7–7.5% NaCl solution. The initial shock bolus is 4–6 mL/ kg over 10–15 minutes. Hypertonic saline should not be administered at a rate exceeding 1 mL/kg/min as vagally mediated bradycardia, hypotension, and bronchoconstriction may be seen with too rapid administration. If needed, this initial bolus can be repeated once; further administration of hypertonic saline can result in serious complications such as hypernatremia and hyperchloridemia. Hypertonic saline administered alone will not expand the vascular space for much time, and is usually combined with a synthetic colloid to prolong its volume‐ expanding effects. Hypotonic crystalloids such as 0.45% NaCl are not appropriate for rapid volume expansion. In certain situations, fluid resuscitation with whole blood or blood component therapy may be indicated. This fluid choice not only provides volume expansion but, in the case of whole blood or packed red blood cells,

36  Fluid Therapy

it also provides oxygen‐carrying capacity that may be crucial in the anemic, hypoperfused patient. One concern that has been visited many times over the years is the risk of aggressive volume resuscitation in patients that are actively bleeding. Concern centers on the disruption of immature clots and the worsening of bleeding. The classic example of this in veterinary medicine is the patient that presents with a hemoabdomen due to splenic trauma. The animal is aggressively resuscitated which results in a short‐term improvement in hemodynamic stability (i.e., blood pressure) followed by a decline as the clots forming on the traumatized spleen are disrupted and active bleeding worsens. “Delayed resuscitation” refers to the practice of not administering fluid support to correct hemodynamic instability until definitive control of hemorrhage has been achieved. “Hypotensive resuscitation” is the practice of administering rapid volume expansion to the hemodynamically unstable patient but only resuscitating to a mean arterial pressure of around 60 mmHg. Here, the goal is to strike a compromise between the need to perfuse vital organs and the desire to avoid supraphysiologic resuscitation that may worsen injuries such as pulmonary contusions and abdominal bleeding. Any of the fluids appropriate for rapid volume expansion can be used for hypotensive resuscitation. When considering the use of either hypotensive or delayed resuscitation, it is important to keep in mind that these techniques were designed for human trauma patients with penetrating injuries, where the time from presentation to definitive treatment rarely exceeds one hour. There has been limited research in veterinary medicine to examine their efficacy in small animal patients. If definitive treatment is not possible or will not be pursued then delayed or hypotensive resuscitation should not be attempted. “Small‐volume resuscitation,” which is also known as limited‐volume resuscitation, uses resuscitative fluids in small volumes to achieve moderate increases in hemodynamic stability. In this type of resuscitation, synthetic colloids and hypertonic saline take center stage as they allow for expansion of the effective circulating volume in excess of the administered fluid volume. Again, the goal here is to minimize the deleterious effects seen with overaggressive crystalloid resuscitations. Endpoints with this style of resuscitation include a mean arterial pressure of 70 mmHg, a systolic pressure of 90 mmHg, and clinical improvement of perfusion parameters. Dehydration The assessment and correction of dehydration remains one of the more challenging parts of fluid therapy. The challenge lies in the poor sensitivity of physical and labo-

ratory parameters to quantify dehydration. It is important to grasp just how limited these parameters really are. It has been shown that an animal can range from 5% to 16% dehydrated based on body weight and yet have a normal skin turgor. This flies in the face of commonly accepted guidelines where the percentage of dehydration is estimated based on the presence or absence of certain physical parameters. In reality, all of these parameters are flawed to a greater or lesser degree. As skin turgor essentially assesses the interstitium, it can be altered not only by the water balance within the interstitium but also by alterations in the other components of the interstitium. For example, in the geriatric or cachectic patient, changes in collagen content may alter the skin turgor so that the animal appears more profoundly dehydrated than it truly is. By contrast, the obese patient may have dehydration but a normal skin turgor. Skin turgor is also affected by the location at which it is assessed, the posture of the animal (standing versus supine), age, and body condition. Although the use of percentage dehydration based on physical parameters is crude, it does still provide a starting point for selecting an initial fluid rate. Usually changes in skin turgor with normal hemodynamic parameters are consistent with mild dehydration of about 5–7% body weight. As dehydration becomes more severe and approaches 10–12% dehydration, hypovolemia may also be evident when perfusion parameters are evaluated. This is evidenced by tachycardia, poor pulse quality, prolonged capillary refill time, and cool extremities. Another indicator of profound dehydration is the presence of sunken eyeballs (enophthalmos). If hypoperfusion is present, rapid volume expansion to stabilize the patient is indicated prior to attempting correction of interstitial or intracellular fluid deficits. When the tissue safety factors of the interstitium are understood, it is clear that rapid volume expansion is not effective for correcting interstitial deficits. This type of deficit must be corrected slowly over at least 12–24 hours. When fluids are administered at an aggressive rate in the hope of shortening the time needed for correction, the results are unrewarding. This is due to the characteristics of the interstitium which oppose rapid expansion. Aggressive fluid administration will result in rapid increases in interstitial hydrostatic pressure as well as decreases in interstitial oncotic pressure. This in turn will increase lymphatic driving pressure and promote lymphatic flow. The end‐result is the clearance of volume via the kidneys instead of a more rapid rehydration of the interstitial space. Commonly assessed laboratory parameters should provide more insight into fluid balance, but they, too, are limited. Alterations in hematocrit and total solids have

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36  Fluid Therapy

Albumin Replacement

Constant Rate Infusions

In critically ill veterinary patients, it may become necessary to address albumin deficiency due to ongoing albumin loss (e.g., protein‐losing nephropathy, septic ­ peritonitis) or decreased albumin production (e.g., acute phase response, liver failure). The majority of total body albumin actually resides within the interstitial space so correction of an albumin deficit can be difficult to achieve using albumin‐containing products such as fresh frozen plasma. Existing interstitial albumin deficits must be replaced before improvement of plasma albumin concentrations will be seen. Albumin concentrate is administered as either a 5% solution or a 25% solution. The 5% solution of human serum albumin (HSA) has a COP of about 20 mmHg and an albumin concentration approximately equal to plasma while the 25% solution has a much higher COP of about 70 mmHg and an albumin concentration five times that of plasma. In veterinary medicine, both 5% and 25% HSA have been administered for albumin replacement in both dogs and cats. Studies have shown both acute and delayed hypersensitivity reactions in dogs. The incidence of these reactions appears lower in critically ill patients but the studies that have investigated the use of these solutions have been limited. In patients with significant albumin deficits, the volume of plasma that would need to be administered to effect change in serum albumin often precludes its use as an effective therapy. For this reason, concentrated albumin products such as 25% HSA or canine lyophilized albumin (a canine albumin product has been introduced to the US market but little has yet been published regarding its use) are more effective in correcting albumin deficits. When correcting albumin, the following equation can be employed:

In addition to their uses in volume expansion and the correction of dehydration, intravenous fluids are often used as carriers for drugs that are best administered by constant rate infusion (CRI). Examples include vasopressors, some inotropes, and some vasodilators. Also, CRIs for the administration of anesthetics and analgesics are commonplace. Before starting a CRI, the compatibility of the drug and carrier fluid must be determined. In most instances, 0.9% NaCl or 5% dextrose in water are acceptable choices. Constant rate infusions are dosed based on both weight and time. For example, a CRI of dopamine might be started at a dose of 5 μg/kg/min. This dose of drug is then administered in a specific volume of fluid. The volume of fluid administered is determined based on the patient’s body weight and fluid status, and the concentration of drug that requires delivery. To calculate a CRI, the following information must be known.

Albumin deficit grams 10 [desired albumin g / dL patient albumin g / dL ] body weight kg 0.3



As an example, to increase the albumin in a 10 kg dog from 1.5 to 2.0 g/dL:

Albumin deficit 10

2.0 1.5

10 0.3 15 grams

Assuming an albumin concentration of 5 g/dL of canine plasma, this patient would require approximately 300 mL of plasma (identical volume to 5% HSA). By contrast, this patient would require 60 mL of 25% HSA. Given the current costs of HSA versus canine plasma, this could result in a significant saving to the client. The recommended dose for the lyophilized canine albumin product has been reported to be 800–884 mg/kg IV.

●● ●● ●●

●● ●●

Patient weight (kg). Drug dose (5 μg/kg/min, for example). Starting rate of fluid administration (5  mL/h, for example). Concentration of the drug administered. Total volume of carrier fluid to be supplemented.

To be precise, when formulating a CRI, an equivalent volume of carrier fluid should be removed and replaced by the drug being delivered. For examples of CRI formulation, the reader is referred to supplemental materials available online.

­Monitoring of Fluid Therapy One of the great challenges of fluid therapy is to meet the patient’s ongoing requirements and replace deficits without overloading the body’s ability to cope with the volume. An understanding of the behavior of fluids ­ within the body as well as body fluid compartment dynamics helps to avoid complications. When attempting to assess the adequacy of fluid therapy, various parameters can be monitored. Perfusion parameters such as heart rate, respiratory rate, capillary refill time, and pulse quality are valuable in determining the need for rapid volume expansion but they do not provide information regarding the correction of dehydration. Perfusion parameters can be normal in the dehydrated animal, although the profoundly dehydrated animal will exhibit signs of hypovolemia. Patients should be weighed at least once daily to track alterations in fluid balance, keeping in mind that 1 L of water is effectively equivalent to 1 kg of weight. The

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importance of body weight monitoring is especially true in the patient in which the potential for third space loss is a concern. Quantification of urine output allows assessment of adequacy of urine output, especially if oligoanuria from renal injury or polyuria from underlying disease processes is of concern. In addition, serial measurements of USG may provide insight into fluid balance. For example, consider the patient on crystalloid fluid therapy who has a decrease in urine output. Once technical complications such as catheter patency and placement have been considered and it has been determined that perfusion is adequate, USG will help to differentiate between physiologic and pathophysiologic oliguria. Recall that the use of hydroxyethyl starches will invalidate the use of ­specific gravity measurement in the urine. The venous side of the circulation is a capacitance system. As such, it has a relatively low pressure until fully replete and distended. Once the venous side of the circulation has no further ability to distend, pressure within the vasculature will rise. This can be correlated to the volumes and pressures within the heart. Based on Starling’s law, it is known that initial expansion of ventricular volume will increase cardiac output and cardiac contractility. Overdistension of the ventricles proves maladaptive and results in decreased cardiac contractility as well as increased intravascular pressures. Central venous pressure is generally reflective of venous volume status and can be a useful invasive monitoring technique to guide fluid therapy. When measuring central venous pressure (CVP), a catheter is placed in the jugular vein in close proximity to but not within the right atrium of the heart. When measured in the absence of pulmonic obstruction or pulmonary hypertension, CVP approximates right atrial pressure. CVP can be measured manually using a water manometer or continuously using a pressure transducer. Normal CVP is considered to be 0–10 cmH20 (1 mmHg is roughly 1.36 cmH2O). CVP should ideally be measured at the end of expiration in the spontaneously breathing or mechanically ventilated

patient but the use of positive end‐expiratory pressure (PEEP) will increase CVP measurements. Although serum lactate is used commonly as an indicator of hypoperfusion, its usefulness as an indicator of dehydration is limited since dehydration must be severe enough to result in decreased oxygen delivery to result in hyperlactatemia. Elevations in serum lactate can also occur due to severe liver dysfunction and spurious elevations in lactate can be seen when difficulty is encountered with sample acquisition.

­Conclusion Fluid therapy is an integral part of small animal practice. Approaching a patient’s fluid needs in a logical fashion where the needs for rapid volume expansion versus correction of dehydration or electrolyte abnormalities are identified and then targeted will maximize the benefits of fluid therapy. Each patient is unique. Each of the commonly available fluid types has properties that can be leveraged to the advantage of our patients. This chapter illustrates how perfusion, hydration, and water and electrolyte balance affect not only the type of fluid therapy that is indicated but also the rate and route of fluid administration. By understanding the physiology of fluid balance in the body, clinicians can tailor their fluid prescription to address interstitial dehydration, correct free water and albumin deficits, and manage electrolyte abnormalities safely. This knowledge also allows fluid choices to be altered during rapid volume expansion to meet the special challenges of each case. The decision to use a specific fluid should be based on the goals of therapy and how effectively that fluid choice will achieve that goal, instead of a knee‐jerk response. Veterinary patients are dynamic and the unique needs of each patient vary. The need to identify our patients’ fluid needs and to address them appropriately, even if their clinical presentation is unusual, cannot be overestimated.

­Further Reading Boldt J. Modern rapidly degradable hydroxyethyl starches: current concepts. Anesth Analg 2009; 108: 1574–82. Craft EM, Powell LL. The use of canine‐specific albumin in dogs with septic peritonitis. J Vet Emerg Crit Care 2012; 22(6): 631–9. Hansen B, DeFrancesco T. Relationship between hydration estimate and body weight change after fluid therapy in

critically ill dogs and cats. J Vet Emerg Crit Care 2002; 12(4): 235–43. Valverde A, Gianotti G, Rioja‐Garcia E, Hathway A. Effects of high‐volume, rapid‐fluid therapy on cardiovascular function and hematological values during isoflurane‐ induced hypotension in healthy dogs. Can J Vet Res 2012; 76: 99–108.

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37 Cardiopulmonary Resuscitation Nathan Peterson, DVM, DACVECC VCA West Los Angeles Animal Hospital, Los Angeles, CA, USA

Approximately 330,000 people die of acute cardiac arrest in the United States annually. The incidence of unexpected cardiopulmonary arrest (CPA) in veterinary patients is currently unknown. What is known is that survival rates for dogs and cats that receive cardiopulmonary resuscitation (CPR) range from 2.6% to 25%. The reported survival rates for people experiencing CPR are equally dismal, with overall survival rates following in‐hospital arrest ranging from 3% to 27%. The reason for the wide range in outcomes is due in part to the variance in the nature or cause of cardiopulmonary arrest. What the literature generally agrees on is that the chance of a patient surviving to discharge after CPA is small. The likelihood of survival increases if CPA was due to an anesthetic event rather than the culmination of a chronic or traumatic disease process. While the survival statistics are dismal, the long‐term neurologic outcomes of patients surviving CPA are generally good to excellent, making CPR a worthy endeavor. In 2010, veterinary‐specific CPR guidelines were developed using an evidence‐based approach modeled on the American Heart Association’s CPR guidelines. The result of this effort was the publication of the Reassessment Campaign on Veterinary Resuscitation (RECOVER) in 2011. The RECOVER initiative (http://onlinelibrary.wiley. com/doi/10.1111/vec.2012.22.issue‐s1/issuetoc) has provided veterinarians for the first time with evidence‐based clinical guidelines for performance of CPR. The progression from severe illness to cardiopulmonary arrest in the veterinary patient is complex. Diseases that can lead to CPA in veterinary medicine include but are not limited to sepsis, pulmonary failure, neoplasia, polytrauma, head trauma, and cardiac failure. Although not all animals progressing toward CPA follow the same clinical course, some physical exam findings that may indicate impending CPA include decreasing level of consciousness, hypothermia, bradycardia, hypotension, and changes in respiratory patterns. Once CPA occurs, all

patients will exhibit loss of consciousness, absence of spontaneous ventilation, absence of heart sounds during auscultation, and absence of palpable pulses. The importance of recognizing CPA quickly is underscored when the hemodynamic impact is considered. Following the onset of CPA, the aortic blood pressure drops precipitously, with intrathoracic aortic pressure dropping from a normal of approximately 120 mmHg to 20 mmHg within 30 seconds. Additionally, carotid blood flow decreases from an average of 190 mL/min to 15 mL/ min within 15 seconds and to 0 mL/min within four minutes. Coronary perfusion pressure likewise drops from 60 mmHg to 15 mmHg within 15 seconds and to 0 mmHg within four minutes. Although the underlying cause of CPA may not be immediately apparent to the clinician, the nature or cause of the arrest can have an impact on the expected outcome and should be ascertained as soon as possible. If a patient is known to have suffered respiratory arrest prior to cardiac arrest, the likelihood of resuscitation may be diminished due to the continued depletion of oxygen from circulating blood resulting in severe hypoxemia at the onset of cardiac failure and institution of CPR. Conversely, if an animal suffers sudden cardiac arrest, the blood oxygen content would be expected to be high at the onset of CPA and subsequent CPR, possibly making successful resuscitation more likely than in the previous instance.

­Principles of CPR Cardiopulmonary resuscitation is intended to provide blood flow to the heart and brain, preserving them until return of spontaneous circulation (ROSC) is achieved. Subsequently, the quality of chest compressions is of the utmost importance, since this is the mechanism by which circulation is generated. Whatever the ­technique,

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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the provision of chest compressions is an inefficient way to move blood out of the heart and into systemic circulation. Even ideal chest compressions are only capable of generating 25–30% of normal cardiac output (CO). However, to maximize the efficiency of chest compressions, the provider should attempt to achieve 30–50% compression of the thoracic diameter. Good– quality chest compressions consist of a 50% duty cycle, meaning 50% of the entire compression cycle is spent with the chest wall being compressed and 50% is spent allowing the chest wall to recoil. Failure to allow complete recoil of the thoracic wall can result in severe impairment to cardiac filling during the noncompression phase. It is often helpful to lift the heel of the hand off the chest wall to ensure that complete recoil has been accomplished. Hand position during CPR in veterinary patients is not standardized and is left to the individual providing compressions to find what works best for them. In small dogs and cats, a one‐handed technique can be employed by placing the sternum in the palm of the hand with the thumb and fingers centered over the heart on opposite sides of the chest. The heart is then compressed between the thumb and fingers in a rhythmic pattern. For larger dogs, it may be necessary to use one or two hands to achieve the desired amount of chest wall compression. In small dogs (15 kg), the compressions should be located at the widest part of the thorax, realizing that this may be at the caudalmost portion of the costal arch. In this case, the clinician is reliant on the intrathoracic pressure generated during chest ­compressions to compress the vascular structures in the thorax forcing blood forward. This is called the thoracic pump theory. An oft‐overlooked facet of chest compressions is patient positioning relative to the compression provider. It is important that the position of the provider is comfortable and will allow for the best‐quality compressions with the least amount of fatigue. Therefore, having the patient at approximately waist level or lower is ideal. If the patient’s position is fixed then the provider should stand on a step stool to achieve the desired amount of leverage. If compressions are provided on a table, the tabletop should be rigid and would ideally be incapable of conducting electricity. Wet tables should be avoided. Larger dogs can often be more comfortably resuscitated on the floor with the compression provider kneeling next to the patient. An adjustable height hydraulic table should be considered if CPR is performed on a regular basis.

­Diagnosing Cardiopulmonary Arrest Prompt recognition of cardiopulmonary arrest is paramount to the success of CPR. Therefore any nonresponsive, apneic patient should be immediately evaluated for CPA and resuscitative measures should be started. The clinician should spend no more than 10 seconds attempting pulse detection and cardiac auscultation before beginning chest compressions. The risk of causing serious injury to a patient by providing unnecessary chest compressions is quite small. In a study of human patients receiving unnecessary chest compressions, the injury rate was less than 2% with no life‐threatening injuries present, leading to the conclusion that the provision of CPR is not harmful, while inaction may prove fatal. If the patient is found to be only in respiratory arrest following the brief pulse evaluation then the clinician should proceed with securing an airway with an endotracheal tube or tracheostomy tube. After the airway is secure, the patient should be manually ventilated with 100% oxygen. If not already obtained, attempts at venous access should be made at this time and the patient should be monitored for progression to cardiac arrest. If, on the other hand, the patient is found to be in cardiac arrest at the time of the pulse check then basic life support (BLS) should be instituted immediately.

­Basic Life Support Chest Compressions While the definitive order in which BLS measures are implemented is not established, it is reasonable to continue using the Airway, Breathing, Circulation approach in which ventilation precedes compressions. Practically, however, instituting compressions does not require any equipment and can often be started earlier than an airway can be secured. Therefore, BLS begins with high‐ quality chest compressions performed at a rate of 100–120 beats per minute, making every effort to avoid interruptions. During the first cycle (two minutes) of chest compressions, the patient should be instrumented with an electrocardiograph (ECG) and capnography if available. Attempts can also be made to achieve intravenous access if this has not already been done. Intubating or securing an airway can be attempted during this first round of compressions while minimizing interruptions. It is often easier to perform the first cycle of compressions while other members of the arrest team obtain proper sized endotracheal or tracheostomy tubes and laryngoscope and mobilize the crash cart to the location of the arrest.

37  Cardiopulmonary Resuscitation

Following the first round of compressions, the first ECG rhythm check will be performed and a definitive attempt will be made to secure an airway. The purpose of the rhythm check is to determine if a cardiac rhythm may be amenable to electrical conversion. If no such rhythm is identified then chest compressions are immediately resumed. Ideally, the chest compression provider is changed at this time. This two‐minute cycle is repeated as needed, alternating the compression provider at each planned rhythm check (i.e., every two minutes) or sooner if fatigue occurs. There are four arrest rhythms, two of which are responsive to electrical defibrillation: pulseless electrical activity (PEA), asystole, ventricular fibrillation, and pulseless ventricular tachycardia. The most common arrest rhythms in veterinary medicine are asystole, which accounts for 72% of arrest rhythms, and pulseless electrical activity, neither of which is responsive to defibrillation. Interruptions in chest compressions must be minimized during CPR. Following each interruption in chest compressions, it takes approximately 60–90 seconds to return CO to the preinterruption level. Eliminating hands‐off time is therefore a major focus in improving both veterinary and human CPR. Minimizing interruptions during CPR must be balanced with the need to ­prevent compression provider fatigue. Many studies in human CPR providers document a tendency to lean during the provision of chest compressions. This tendency increases as the duration of compressions increases. The effect of leaning on a patient during CPR decreases cardiac filling and the overall efficiency of CPR by at least 25%. CPR is hard work and two minutes of high‐quality chest compressions on even a small to medium‐sized dog should be exhausting. Airway Orotracheal intubation of dogs and cats is relatively easy, making rapid intubation possible in most circumstances. In the majority of patients that experience cardiopulmonary arrest, an airway can be secured via routine orotracheal intubation. Occasionally, the glottis is obscured by vomit, fluid or saliva. If suction is available, it can be helpful in clearing the oropharynx of excess fluid. If suction is unavailable a gauze 4 × 4 or paper towel can be used to swab the area free of fluid or debris. Care should be taken to avoid excessive manipulation of the epiglottis as this can induce a vagal response. A laryngoscope should be used to facilitate intubation. If the glottis cannot be visualized, the orotracheal route may still be considered for intubation if the epiglottis can be palpated. A blind intubation is performed by palpating and depressing the tip of the epiglottis and directing the endotracheal tube through the glottis

­orsally. Successful intubation can be confirmed by d direct visualization of the endotracheal tube between the arytenoid cartilages or visualization of normal chest rise ­during positive pressure ventilation. Capnography may be useful but should not be used as the sole means of ­confirming correct placement of an endotracheal tube ­during CPA since end‐tidal CO2 in CPA patients may be 0 due to lack of pulmonary blood flow. Once an endotracheal tube is placed and correct positioning is confirmed, the tube should be secured to the patient with tape, gauze tie or rubber band to prevent tube dislodgment. The cuff on the endotracheal tube should be inflated to ensure an airtight seal. This will allow the patient to be effectively ventilated with positive pressure during chest compressions. In some cases, orotracheal intubation may not be possible due to severe swelling, inability to open the mouth or severe facial trauma necessitating emergency tracheostomy. Ventilation Positive pressure ventilation with 100% oxygen is begun once an airway is secured. Breaths can be provided by a number of means, including rescue bag, anesthesia machine or mechanical ventilator. Hyperventilation is common in the clinical CPR scenario and can be caused by an excessive respiratory rate, tidal volume or inspiratory time. Regardless of the cause, hyperventilation is detrimental to outcome. During positive pressure ventilation, venous return occurs during the expiratory phase and if insufficient expiratory time is allowed (due to either excessive rate or inspiratory time), cardiac filling will be reduced. Hyperventilation can also lead to decreased cerebral carbon dioxide levels and subsequent cerebral vasoconstriction, further impairing blood flow to the brain during the critical time of resuscitation. No clear association has been made regarding the timing of rescue breathing with compressions and it is not recommended to attempt to synchronize breaths with compressions. Current recommendations for ventilation are a rate of 8–10 breaths per minute with a tidal volume of approximately 10 mL/kg and an inspiratory time of one second. Assigning the role of rescue breathing to a single team member and providing them with clear ­ revent instructions on ventilation rates is the best way to p hyperventilation.

­Advanced Life Support Advanced life support (ALS) consists of all resuscitative measures that are not included in the BLS algorithm, including vasopressor therapy, vagolytic therapy, fluid and electrolyte therapy, and electrical defibrillation.

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The rationale for the use of vasopressor therapy is grounded in the fact that cardiac output during CPR is a fraction of that during spontaneous circulation, making it imperative that the limited cardiac output is directed to the organs that need it most. Therefore, the provision of vasopressors is intended to increase peripheral vascular resistance, forcing more blood into the central circulation and subsequently improving the perfusion of vital structures (i.e., brain and heart). Stimulation of the alpha‐1 receptor results in peripheral vasoconstriction whereas stimulation of the beta‐1 receptor increases both inotropy and chronotropy. While epinephrine is a balanced alpha‐ and beta‐adrenergic agonist, its utility during CPR comes primarily from its action at the alpha‐1 receptor. In fact, the effects of beta‐1 stimulation may be detrimental as they can lead to increased myocardial oxygen consumption and predispose to arrhythmias if ROSC is achieved. The best dose for epinephrine use during CPR has been a topic of debate for many years, with mixed evidence. High‐dose epinephrine (0.1 mg/kg IV) has been shown to result in slightly higher rates of ROSC but has not been shown to confer a survival benefit. Based on the lack of documented improved survival, the use of low‐dose epinephrine (0.01 mg/kg) is currently recommended early in CPR efforts, with high‐dose epinephrine being reserved for cases of prolonged CPR. Epinephrine should be administered following the first cycle of BLS and then following every other cycle. Adrenergic receptors have been demonstrated to be less responsive in acidotic conditions, such as those found during CPA, making an alternative drug that is not dependent on catecholamine receptors an appealing option. Vasopressin acts on the V1 receptors of vascular smooth muscle cells independent of the alpha‐1 receptor. In addition, it maintains its efficacy in the face of acidosis. Due to the lack of adrenergic effects, vasopressin does not cause increases in inotropy or chronotropy, minimizing its impact on myocardial oxygen consumption. The human literature regarding the usefulness of vasopressin in CPR is mixed, with promising results in small trials but failure to demonstrate any clear advantage over epinephrine in larger metaanalyses. The successful use of vasopressin in CPR has been reported in the veterinary literature. Administration of vasopressin (0.8 U/kg IV) in place of or in conjunction with epinephrine may be considered, with subsequent doses administered following every other cycle of BLS. The use of atropine in ALS is somewhat unique in that atropine is a parasympatholytic drug intended to ameliorate high vagal tone during cardiac arrest rather than directly stimulating catecholamine receptors. The use of atropine should be considered in animals that have asphyxia‐induced cardiac arrest or other co‐morbid

c­ onditions that would be likely to increase vagal tone. A dose of 0.04 mg/kg IV is recommended as higher doses are more likely to result in severe tachycardia and subsequently, increased myocardial oxygen consumption if ROSC is achieved. Repeated dosing of atropine is unlikely to be of benefit (assuming actions directed at the underlying cause have been taken) and should be considered carefully due to the potential for severe tachycardia if resuscitation is successful. Corticosteroids currently have no place in the CPR algorithm and their use is not recommended. Given the lack of documented improvement in outcome and the known deleterious side‐effects, especially in patients with perfusion impairment, this is unlikely to change. The administration of ALS drugs by intravenous or intraosseous routes (interchangeable in the context of CPR) is always preferred to the intratracheal route. If IV or IO access is not available or not possible then intratracheal administration of drugs can be considered while efforts to attain venous or intraosseous access are continued. If this route is utilized, a long catheter should be advanced through the endotracheal or tracheostomy tube to the level of the carina or beyond. Drugs should be diluted with saline or sterile water. The optimal dosing of drugs for intratracheal administration is not established but doses up to 10 times the normal dose have been recommended. Once established, the IV or IO route should be used for all subsequent drug administrations. The need for rapid volume expansion with IV crystalloid or colloid solutions during CPR is dependent on the underlying cause of CPA. Animals suffering CPA from severe hemorrhage or hypovolemia should be aggressively resuscitated with IV fluids to reestablish normal circulating volume. Since coronary perfusion pressure is determined by the difference between diastolic aortic blood pressure and right atrial pressure, care should be taken to prevent overresuscitation. IV fluids accumulate in the venous compartment, increasing the central venous and right atrial pressure and consequently reducing coronary perfusion pressure. In general, animals that were euvolemic or hypervolemic (congestive heart failure) do not require volume expansion during CPR and routine use of IV fluids during CPR on these patients is not recommended. Electrolyte and acid–base disturbances are common during CPA. Severe acidemia occurs frequently during CPA, especially later in the course of resuscitation, and can result in vasodilation, predisposition to arrhythmias, and impaired neurologic function. The administration of 1 mEq/kg sodium bicarbonate can be considered in prolonged instances of CPA lasting 10–15 minutes. The most common electrolyte disturbances during CPA include hyperkalemia and hypocalcemia. Because of the lack of documented improvement in outcomes and the

37  Cardiopulmonary Resuscitation

potential complications associated with over‐ or undercorrection, the management of electrolyte derangements during CPA should be reserved for animals with confirmed rather than assumed disturbances. Recently, the use of an impedance threshold device (ITD) intended to increase cardiac output during CPR has been described in canine patients and results appear promising. Unfortunately, the use of an ITD is limited to patients greater than 10 kg.

­Defibrillation The purpose of defibrillation is to depolarize as many cardiac myocytes as possible, making them temporarily refractory, allowing the native pace‐making cells to resume organized electrical conduction. Electrical defibrillation should only be attempted on rhythms that are potentially responsive to it. Of the four arrest rhythms, only two are responsive to electrical defibrillation: ventricular fibrillation (most significant in veterinary medicine) and pulseless ventricular tachycardia. Any attempts at converting nonresponsive rhythms will not result in return of spontaneous circulation and will only needlessly expose team members to accidental defibrillation and increase time spent without compressions. There are two types of defibrillators commonly used in veterinary medicine. Monophasic defibrillators function by passing a single electrical current from one electrode through the heart to a receiving electrode. Biphasic defibrillators pass a current from one electrode to the other; the current then reverses direction and is passed back to the original electrode. This technology allows effective defibrillation to be performed using less energy than monophasic defibrillation, resulting in less myocardial injury. If available, biphasic defibrillation is preferred. Being familiar with the type of defibrillator in the ­practice is important due to the difference in energy selection, with monophasic defibrillator doses beginning at 4–6 J/ kg and biphasic defibrillator doses beginning at 2–4 J/kg. If defibrillation is not successful following the first attempt, the dose delivered can be increased by 50% for subsequent defibrillation attempts. Historically, it was recommended to administer three stacked defibrillation attempts (discharges in rapid succession) prior to resuming chest compressions. The advent of biphasic defibrillators and the recognition that delays in resumption of chest compressions can have a negative effect on outcome have made this recommendation obsolete. It should be remembered that successful defibrillation means that fibrillation has been stopped, not that ROSC has been achieved. Successful defibrillation therefore can result in asystole or an electrocardiographically normal rhythm. However, resumption of

normal electrical function does not necessarily correlate with return of normal mechanical function of the heart (i.e., PEA). It is therefore recommended that a single defibrillation attempt be made followed by immediate resumption of chest compressions for a full two‐minute cycle prior to ECG evaluation. This improves coronary blood flow to the myocardium, making successful defibrillation with resumption of mechanical activity more likely. Performing defibrillation begins with selecting appropriately sized paddles. Most defibrillators come equipped with standard‐sized adult paddles, which are suitable for most patients larger than 15 kg. Smaller pediatric paddles are usually available as a clip‐on attachment and should be used for smaller dogs and cats to ensure focused delivery of the shock through the heart. Defibrillator paste or contact gel is generously applied to the paddles. The animal is typically placed in dorsal recumbency and the defibrillator paddles are placed on either side of the thorax at the level of the costochondral junction with the heart centered between the paddles. The patient is firmly grasped between the paddles and is maintained in dorsal recumbency by the person delivering the defibrillation. Once the patient is positioned, depressing the charge button on the paddles or on the base unit charges the defibrillator. To maintain safety, the patient should be positioned prior to charging the paddles to lessen the likelihood of accidental discharge. Once the defibrillator is charged, the provider should ensure that no one is in contact with the patient or metal tabletop. Verbal commands to “CLEAR” the patient are effective. Once all rescue providers are clear of the patient, the shock is delivered by depressing the shock delivery buttons on the paddles or defibrillator base unit. Animals with thick undercoats that inhibit paddle contact should be rapidly clipped at the intended paddle locations prior to defibrillation attempts. The use of alcohol to improve paddle contact should be strictly avoided due to the risk of fire. Alternative defibrillation techniques involve the use of defibrillation pads attached directly to the patient and left in place while connected to the defibrillator base unit. The pads are placed on the chest in the same location as the paddles. If pads are used then clipping of the hair is mandatory to ensure good contact between the pads and the patient. The advantage to the use of pads is that no rescue providers have to be in contact with the patient and patient positioning is minimally interrupted. The disadvantage is the necessity of clipping the hair that results in longer time without chest compressions. The use of a posterior paddle assembly, a flat paddle replacement resembling a spatula, may be the best method for delivering defibrillation in veterinary patients. The paddle assembly is coated with paste or gel, as a normal

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­ addle would be. The assembly is then placed under the p patient in lateral recumbency using the normal paddle positioning recommendations. A standard paddle is then used on the upward‐facing thoracic wall. This technique allows the patient to remain in lateral recumbency while eliminating the need to clip the area as for defibrillator pads. Nonelectrical methods of cardiac defibrillation include both chemical and mechanical means. Historically, chemical defibrillation with magnesium or antiarrhythmic drugs has been considered acceptable. Based on the current understanding of CPA in both human and veterinary patients, the use of antiarrhythmic agents is no longer routinely recommended. Among the possible agents available, only amiodarone has been found to be routinely effective. Magnesium salts have only proven to be useful for the treatment of torsades de pointes and electrical defibrillation should always be considered as the best option for ventricular fibrillation or pulseless ventricular tachycardia. The use of a precordial thump has been described as a method for providing mechanical defibrillation. Given the superiority of electrical ­defibrillation, a precordial thump should only be considered if a defibrillator is not available.

­Monitoring During CPR Monitoring during CPR can be difficult due to lack of normal physiologic responses. The two modalities most  useful during CPR are ECG and capnography. Electrocardiography is monitored to determine if an arrest rhythm is amenable to defibrillation while ETCO2 can be utilized as a measure of the effectiveness of chest compressions, as an adjunct to verify placement of the endotracheal tube, and to determine when ROSC occurs. Traditionally, patients with CPA would be subjected to pulse detection attempts before and during CPR. Unfortunately, pulse detection takes time, is technically difficult in some cases and can lead to increased “hands‐ off ” time, with the patient receiving fewer chest compressions. Capnography has allowed CPR providers a method to monitor not only the effectiveness of chest compressions but also whether or not ROSC has occurred. Following CPA, blood flow through the pulmonary circulation decreases rapidly and ETCO2 levels drop to near 0 since carbon dioxide is not being returned to the alveoli for exhalation. During CPR, pulmonary blood flow is generated by chest compressions. Therefore, any change in ETCO2 levels can be attributed to the ­cardiac output achieved by the chest compressions, provided the minute ventilation is held constant. ETCO2 is also useful for the detection of ROSC, with a rapid and sustained rise in ETCO2 indicating ROSC due to the

much more efficient delivery of blood to the pulmonary circulation by spontaneous cardiac contraction than that achieved during manual chest compressions. Following the initiation of chest compressions and attainment of an airway, the patient should be instrumented with ETCO2 monitoring if available. Patients with higher ETCO2 levels (15 mmHg dogs, 20 mmHg cats) are more likely to have ROSC than those with lower levels. During CPR, if ETCO2 is less than 15 mmHg, efforts should be made to improve the quality of chest compressions. If compression providers have not been rotated and ETCO2 is found to be low or is decreasing, a rotation should be considered. Blood gas monitoring during CPR is challenging but is the only way to identify the severe electrolyte and acid– base derangements that are common during CPA. Ideally, a blood sample (venous or arterial) is obtained shortly after the onset of CPA and is evaluated for any abnormalities. Interventions should be directed at severely abnormal results, understanding that complete resolution is not likely and may not be necessary. If sampling is possible, blood gas analysis should be performed every 5–10 minutes during CPR as long as collection of samples does not interfere with the actual provision of rescue efforts. Information regarding acid–base and electrolyte status during CPA and outcome is lacking and should not be used to predict the likelihood of ROSC.

­Open Cardiac Massage Open chest cardiac massage is capable of generating better cardiac output during CPR than closed chest compressions. However, the procedure is invasive and requires significant resources both during and after resuscitation. Therefore, its use is reserved for patients with a clear indication including significant intrathoracic disease (tension pneumothorax, severe pleural effusion), major chest wall defect (flail chest) or pericardial effusion. To perform open chest CPR, the patient is placed in right lateral recumbency. The fur is rapidly clipped over the 4th–6th intercostal spaces and the skin is quickly wiped with alcohol. The skin is incised at the 5th–6th intercostal space with a scalpel blade. The intercostal muscles are incised with Mayo scissors and the incision is extended from the costochondral junction to the proximal one‐third of the rib. Care should be taken to avoid laceration of the lung tissue during this process. Respiration can be temporarily held during this time to further reduce the risk of iatrogenic trauma to the lung. The ribs are then spread with manual retraction or a Fineccheto retractor. The pericardial sac is incised at the

37  Cardiopulmonary Resuscitation

apex of the heart to avoid the phrenic nerve and the sac is then reflected dorsally to the base of the heart. The heart should be compressed from the apex to the base at a rate of 100–120 compressions per minute. This can be done with one or two hands, depending on patient size. In some patients, the descending aorta can be briefly (10–15 minutes) cross‐clamped to help direct blood flow to the cerebral and coronary circulation. If ROSC is achieved, the patient should be maintained in an anesthetized state while a thoracostomy tube is placed and the thoracotomy site is closed. Volatile gas anesthetics have profound vasodilator effects and may be detrimental to a recently resuscitated patient. All efforts should be made to avoid hypotension and total IV anesthesia may be the best option during this time. Prophylactic antibiotics should be considered for patients that have undergone open chest CPR due to the likelihood of contamination.

­Postresuscitation Care Despite initial reported ROSC rates of 35–45% in veterinary medicine, survival to discharge rates range from 2% to 10%. This highlights the need for continued aggressive critical care following ROSC. Initial postcardiac arrest (PCA) care should focus on preventing rearrest, specifically targeting hemodynamic stability and pulmonary/ respiratory function. Specific goals of hemodynamic stabilization should include normalization of central venous oxygen saturation, central venous pressure, cardiac output, arterial blood pressure, and arterial oxygen content. IV fluid therapy should be judicious and efforts should be made to prevent overresuscitation. Ventilatory function may be altered in PCA patients with resultant hyper‐ or hypoventilation. Ventilation should be assessed and positive pressure ventilation should be provided as ­ needed to maintain normocapnia (PaCO2 35–45 mmHg). Supplemental oxygen or mechanical ventilation is used as needed to maintain normoxemia (PaO2 90–100 mmHg).

As the PCA period increases, focus should be directed at preventing or minimizing secondary organ damage and neuroprotection. The routine use of corticosteroids, hyperosmotic fluids, and anticonvulsants is not recommended. If evidence of increased intracranial pressure is present then treatment with mannitol or hypertonic saline is warranted. Should generalized seizures develop then appropriate treatment with an anticonvulsant should be started. Prophylactic treatment with anticonvulsant medications may be considered in animals demonstrating global cerebral signs (coma, compulsion) in the postarrest period. The use of corticosteroids has not been documented to improve outcome and their use should be restricted to animals with a clear indication due to the potential for severe adverse effects. Induction of mild therapeutic hypothermia is rapidly becoming the standard of care in human PCA management. Its use has been described in the veterinary literature but is not widespread at this time. Due to the risks associated and the technical difficulty of safely inducing and maintaining appropriate hypothermia, it is not currently recommended. If, however, mild hypothermia develops spontaneously, the patient should not be aggressively warmed; rather, the return to normothermia should be gradual (0.25–0.5 °C/h).

­Discontinuing CPR The decision to terminate CPR generally lies with the clinician. When making the decision to withhold or terminate CPR, several factors should be considered including the inciting cause of the arrest, interval from onset of arrest to initiation of CPR, duration of CPR, and long‐term prognosis associated with the underlying disease process. Although no definitive guidelines exist, continuing efforts for 15–20 minutes is reasonable, after which time the likelihood of a good neurologic outcome is exceedingly small and CPR can be discontinued.

­Further Reading Boller M, Kellett‐Gregory L, Shofer FS, et al. The clinical practice of CPCR in small animals: an internet‐based survey. J Vet Emerg Crit Care 2010; 20(6): 558–70. Fletcher DJ, Boller M, Brainard BM, et al. RECOVER evidence and knowledge gap analysis on veterinary CPR. Part 7: Clinical Guidelines. J Vet Emerg Crit Care 2012; 22(S1): S102–S131. Lee SG, Moon HS, Hyun C. The efficacy and safety of external biphasic defibrillation in toy breed dogs. J Vet Emerg Crit Care 2008; 18(4): 362–9.

Plunkett SJ, McMichael M. Cardiopulmonary resuscitation in small animal medicine: an update. J Vet Intern Med 2008; 22: 9–25. Waldrop JE, Rozanski EA, Swanke ED, et al. Causes of cardiopulmonary arrest, resuscitation management, and functional outcome in dogs and cats surviving cardiopulmonary arrest. J Vet Emerg Crit Care 2004; 14(1): 22–9.

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38 Respiratory Monitoring in Critical Care Mathew Mellema, DVM, PhD, DACVECC Applaud Medical, San Francisco, CA, USA

­Purpose The purpose and goals of respiratory monitoring in critical care typically include the following: ●●

●●

●● ●● ●● ●●

identification of patients in respiratory distress and those experiencing dyspnea determination of response to specific or empiric therapies assessment of the adequacy of alveolar ventilation assessment of the adequacy of oxygenation evaluation of gas exchange efficiency evaluation of pulmonary mechanics in order to categorize disease states, explain other related clinical findings, or monitor response to therapy.

I­ dentifying Respiratory Distress and Dyspnea Respiratory distress is a state in which breathing efforts are excessive or inadequate relative to the healthy state or the patient’s current requirements. While many definitions rely on inadequate oxygenation or ventilation, such approaches fail to account for patients in whom adequate gas exchange is obtained only with excessive effort. Tachypnea (increased respiratory rate) and hyperpnea (increased respiratory effort and/or chest excursions) are often considered the hallmarks of respiratory distress; however, a smaller subset of patients may present with hyponea (decreased respiratory effort and/or chest excursions), bradypnea (decreased respiratory rate), or both. This second type of patient is also in respiratory distress. Dyspnea is a term often used synonymously with respiratory distress in veterinary practice, although it has been suggested that this convention is not ideal. In human medicine, dyspnea refers to the unpleasant

s­ ensations of feeling unable to catch one’s breath or being consciously aware of one’s breathing effort. The three subcategories of dyspnea sensations identified in humans are air hunger, increased work/effort, and asthmatic chest tightness. Experimental evidence exists to suggest that vertebrates other than humans are able to experience the first two of these. To date, no experimental approaches have been developed to determine if animals are likely to experience asthmatic chest tightness. Ideally, the term dyspnea would be reserved for consideration of the sensory experience (akin to pain) and respiratory distress would be used to describe what clinicians might observe (akin to nociceptive responses). Serial monitoring of patients for signs of respiratory distress or dyspnea is the cornerstone of respiratory monitoring. Respiratory monitoring may include many other tools, but these are always best evaluated in light of current patient status and clinical context. Clinical Signs of Respiratory Distress Box 38.1 provides an outline of broad categories of clinical signs that may be present in a patient with respiratory dysfunction or distress. Two categories of sounds may herald respiratory distress: those heard with the ear and those heard with the stethoscope. Stertor and stridor are sounds that typically do not require a stethoscope to be audible. Stertor is the presence of snoring‐like noises during respiration. It is the result of abrupt displacement of upper airway soft tissue structures and altered gas flows. It is typically low‐pitched, discontinuous, and nonmusical in character. It may be evident on inspiration, exhalation, or both. Stridor is a high‐pitched, musical respiratory sound generated by altered gas flows in the large airway. It is more typically heard on inspiration. Either stertor or stridor can indicate an abnormality of the proximal upper airways (i.e., cranial to the thoracic inlet).

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Box 38.1  Categories of signs associated with respiratory distress in dogs and cats Abnormal sounds Heard without a stethoscope –– Stertor –– Stridor ⚪⚪ Heard with a stethoscope –– Diminished breath sounds Regional Focal –– Adventitious sounds Crackles Wheezes Rhonchi –– Borborygmus ●● Changes in position/posture/orientation ⚪⚪ Orthopnea ⚪⚪ Trepopnea ⚪⚪ Neck extension ⚪⚪ Upright stance (flexed pelvic limbs, extended ­thoracic limbs) –– Sternal recumbency (Sphinx pose) ⚪⚪ Abducted elbows ●● Reflex compensatory changes due to hypoxemia and hypercapnia ⚪⚪ Tachypnea ⚪⚪ Tachycardia ⚪⚪ Hyperpnea ●● Signs associated with reduced/altered gas flow velocities ⚪⚪ Phase prolongation –– Inspiration –– Exhalation –– Both ⚪⚪ Inability to vocalize ●● Adaptive responses which reduce respiratory system resistance ⚪⚪ Open‐mouth breathing –– Retracted commissures of the lips ⚪⚪ Nares flaring ⚪⚪ Neck extension ●● Signs secondary to marked hypoxemia ⚪⚪ Cyanosis ⚪⚪ Obtundation ●● Abnormal or excessive abdominal motion ⚪⚪ Passive abdominal motion due to hyperpnea ⚪⚪ Active expiratory abdominal muscle contraction ⚪⚪ Paradoxical motion ●● Signs of the sensation of dyspnea ⚪⚪ Obtundation ⚪⚪ “Anxious” expression ⚪⚪ Dilated pupils ⚪⚪ Poorly tolerant of restraint/fractious ⚪⚪ Not eating, drinking, sleeping, and/or grooming ●●

⚪⚪

Altered breath sounds can also herald respiratory distress in small animal patients and may also aid in the determination of the underlying cause. Breath sounds may be altered in their distribution, their volume (i.e., loudness), and their character. Diminished breath sounds in the nondependent lung fields may indicate accumulation of gas in the pleural space, whereas the same finding in the dependent lung regions suggests pleural fluid accumulation or nonventilated lung (or both). Focal absence of breath sounds may indicate that one or more lung lobes are obstructed, consolidated, collapsed, torsed, infiltrated, or compressed by an intrathoracic mass lesion. In addition, adventitious sounds (crackles, wheezes, rhonchi) may indicate lower airway and pulmonary parenchymal pathology. Crackles are explosive, discontinuous, nonmusical sounds generated by small airways abruptly opening. They may be fine or harsh in character. While often heard on inspiration (opening during expansion), they may also be present during exhalation in some settings. Wheezes are higher pitched, musical, and of longer duration than crackles. They are often the result of diffuse lower airway narrowing. Rhonchi are similar to wheezes but substantially lower in pitch. Abnormal sounds that might be ausculted over the thorax also include borborygmus, which may indicate a compromise of the diaphragm with displacement of intestinal viscera into the thoracic cavity. Changes in body position/posture, orientation of the head and limbs, respiratory rate, and the magnitude of inspiratory excursions all may indicate respiratory system compromise. Patients in respiratory distress often assume an upright posture with the pelvic limbs flexed and the thoracic limbs extended. This posture allows the weight of the abdominal contents to rest on the pelvis rather than the diaphragm and reduces the work of breathing. In cats, a more typical posture is to sit propped in sternal recumbency (sphinx pose). Patients that exhibit greater respiratory distress when in recumbency are termed orthopneic and this may manifest as a reluctance to lie down or be restrained. The head is often held in dorsoflexion with the neck extended. This position reduces the work of breathing by straightening the trachea and minimizes narrowing. Moreover, by fixing the head position in this way, accessory respiratory muscles (e.g., serratus ventralis muscle) can be utilized to promote expansion of the cranial thorax. Elbow abduction and fixing the position of the forelimbs allows for the recruitment of other accessory respiratory muscles and reduces the impact of the weight of these limbs on chest expansion. Respiratory compromise that is more severe in one lateral recumbency than the opposite is termed trepopnea and often is due to asymmetric pulmonary parenchymal disease. Increases in respiratory rate are often the earliest sign of respiratory system compromise. Tachypnea at rest may frequently precede more overt signs in patients with

38  Respiratory Monitoring in Critical Care

chronic cardiac or respiratory disease. Tachypnea and hyperpnea with exertion are also considered early signs in this same setting. Both tachypnea and hyperpnea are adaptive responses in most settings, but not all. These responses represent the clinically apparent consequences of attempts to maximize alveolar ventilation. However, they may be triggered by pain, hypotension, and other factors. In patients with upper airway disease, these responses are generally maladaptive as they increase the work of breathing and may promote further airway narrowing via the Venturi effect. Tachypnea must be distinguished from panting which occurs in hyperthermic (not febrile) animals with the mouth open. Febrile animals have a reset thermoregulatory set‐point and thus the elevation in body temperature will only result in occasional panting when body temperature temporarily exceeds this threshold. Tachypnea does not imply open‐mouth breathing whereas this is a hallmark of panting. Panting does not alter ventilatory status whereas tachypnea may result in hyperventilation in some settings but not others. Respiratory monitoring also includes assessing patients for clinical signs relating to reduced respiratory gas flow velocity. Such flow reductions may manifest as prolongations of the inspiratory (upper airway disease) or expiratory (intrathoracic airway disease) phases of respiration. Both phases of respiration may be affected concurrently in some disease states. One example is feline asthma. Bronchoconstriction results in increased resistance to both inspiratory and expiratory flows, although the increase is generally more marked during exhalation. An inability to vocalize indicates a severe reduction in expiratory gas flows and should prompt immediate assessment of upper airway patency. Several clinical signs that manifest in veterinary patients with respiratory distress represent adaptive responses that serve to reduce airway resistance. Neck extension as discussed above is one example. Flaring of the nares represents recruitment of the dilator naris muscle and is stimulated by hypoxemia and hypercapnia and inhibited by pulmonary stretch receptor activation. Breathing through an open mouth also ­ reduces respiratory system resistance while foregoing the benefits of gas conditioning (humidification, particle trapping) by the nasal passage. Animals in respiratory distress often breathe with their mouths open although other conditions (pain, hyperthermia) can also elicit this same response. Animals in respiratory distress typically do not hang the mouth passively open, but rather actively retract the commissures of the lips and extend the tongue outward. Activity of the genioglossus muscle may be synchronized with the respiratory cycle, resulting in cyclic protrusion of the tongue outwards. Cyanosis is a severe but unreliable sign of respiratory compromise. When present, it generally indicates severe

hypoxemia. However, in anemic patients, the amount of deoxygenated hemoglobin present in the capillary beds may be too low to result in a color change detectable to the clinician. Abnormal or excessive abdominal motion during respiration may indicate respiratory distress. Passive abdominal motion and increased abdominal effort are not the same thing, although they may look similar from a distance. More pronounced outward motion of the abdomen on inspiration is passive and caused by increased contraction of the diaphragm. Active contraction of the abdominal muscles (e.g., rectus abdominis) represents recruitment of accessory respiratory muscles whereas passive displacement of the abdominal contents during inspiration does not. The clinician may need to palpate the ventral abdominal muscles during exhalation to distinguish between the two conditions. Tensing of the rectus abdominis during exhalation can usually be clearly felt when increased abdominal effort is truly present. Activation of the abdominal muscles during respiration actively aids exhalation and passively assists inspiration. Increasing intraabdominal pressure (and thus passively increasing intrathoracic pressure) raises the pressure driving expiratory gas flows. This increase in abdominal pressure also serves to place the diaphragm in a more cranial position prior to the next inspiratory effort. This diaphragmatic shift places it in a more favorable orientation for contraction and thus abdominal muscle activity can enhance diaphragmatic performance. True increased abdominal effort generally indicates expiratory flow limitation due to intrathoracic airway narrowing or collapse. Some patients may have atypical abdominal wall motion. Paradoxical motion (abdomen moving inward on inspiration) may be seen with diaphragmatic hernia, fatigue, or paralysis. A more focal inward movement of the abdominal wall adjacent to the costal arch is often seen in patients with pleural effusion. Any significant motion of the soft tissues in the region overlying the thoracic inlet may indicate abnormal or increased ­ ­respiratory effort. Clinical Signs of Dyspnea Small animal patients may also exhibit clinical signs that one might attribute to the unpleasant sensation of dyspnea. Such signs include a blunted response to the environment (obtundation) as the patient focuses solely on the act of breathing. An “anxious” facial expression with dilated pupils bilaterally is common. Such patients are typically not eating, drinking, or sleeping adequately. A patient that has been in respiratory distress and is now devoting energies to these other needs is likely improving. In cats, the absence of grooming behaviors may be associated with sensations of dyspnea. Returning to ­normal hygiene routines often accompanies a marked

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improvement in respiratory status. Patients that are experiencing dyspnea are often excessively fractious and resistant to physical restraint. Respiratory monitoring in the critically ill veterinary patient should include initial and serial evaluation for the presence of all the above physical exam findings. Serial evaluation often allows the clinician to put quantitative data (see later) into their proper context. The response to specific or empiric therapies is best assessed based on a combination of these physical exam findings, imaging, and quantitative laboratory assay results.

­Determination of Response to Specific or Empiric Therapies Respiratory monitoring plans are frequently designed with the goal of assessing response to therapy. Serial monitoring of respiratory rate is perhaps the most widely used approach in most settings (e.g., monitoring response to diuretics in presumed congestive heart failure patients). Other examples might include the assessment of pulse oximetry, respiratory effort, and breathing ­pattern in response to oxygen supplementation. Such minimally invasive approaches can be quite useful but in other cases, more invasive procedures such as intubation, positive pressure ventilation, and arterial blood gas sampling are required both to correct the problem and to monitor progression. Marked improvement in respiratory status following intubation is expected for upper airway obstruction in the absence of other co‐morbidities. Likewise, hypoxemia due solely to hypoventilation should resolve with oxygen supplementation in all cases short of apnea or agonal breathing. Failure of hypoxemia to resolve with oxygen supplementation typically indicates significant venous admixture and physiologic or anatomic right‐to‐left shunting. Hypoxemia that resolves with oxygen supplementation in a patient that simultaneously demonstrates

little or no improvement in clinical signs of respiratory distress can be indicative of pulmonary thromboembolism (PTE). Respiratory monitoring plans should be tailored to the individual patient and the pathology ­ believed to be present.

­Assessment of the Adequacy of Alveolar Ventilation The alveolar gas equation or “PCO2 equation” (PACO2 ~ VdotCO2/VA) provides the rationale for all forms of ventilatory status monitoring. The equation states that the partial pressure of carbon dioxide in alveolar gas varies directly with CO2 production and inversely with alveolar ventilation. The adequacy of alveolar ventilation is defined by the alveolar partial pressure of carbon dioxide. If alveolar ventilation is adequate for the current level of CO2 production then PACO2 will be within the normal range. If alveolar ventilation is inadequate or excessive, the PACO2 will be increased or decreased, respectively. The rate of production of carbon dioxide (VdotCO2) is largely irrelevant. Ventilatory adequacy is defined relative to current carbon dioxide production no matter what that current level might be. In extreme cases such as malignant hyperthermia, carbon dioxide production exceeds the level that any healthy respiratory system could remove from the system; however, those patients are still defined as hypoventilating despite having normal (or even exceptional) respiratory function. Alveolar gas composition is not uniform across all aveoli and is exceedingly difficult to sample. Thus, the partial pressure of carbon dioxide from other sites serves as a surrogate in clinical practice. Clinically useful surrogate samples include arterial blood, venous blood (mixed, central, or peripheral), and end‐tidal exhalate. Typical canine blood values for PCO2 are shown in Table 38.1.

Table 38.1  Blood gas analysis reference intervals

pH

Arterial (cat)

Arterial (dog)

Mixed venous (dog)

Central venous (dog)

Peripheral venous (dog)

7.39 +/−0.08

7.40 +/−0.03

7.36 +/−0.02

7.35 +/−0.02

7.36 +/−0.02

PCO2

31 +/−5

37 +/−3

43 +/−4

42 +/−5

43 +/−3

Base deficit

−2 to +8

−4 to 0

−2 to 0

−4 to 0

−2 to 0

Bicarbonate

18 +/−4

21 +/−2

23 +/−2

22 +/−2

23 +/−1

Total CO2

N.D.

22 +/−2

24 +/−2

23 +/−2

24 +/−2

PO2

105 +/−10

102 +/−7

53 +/−10

55 +/−10

58 +/−9

Source: Adapted from Ilkiw et al. (1991). Values are presented as mean +/− standard deviation. N.D., not determined.

38  Respiratory Monitoring in Critical Care

The adequacy of each of these sample types is dependent on patient factors as will be discussed later. Arterial blood is often an ideal surrogate sample for evaluating ventilatory status. Carbon dioxide is highly soluble and diffusible and alveolar partial pressures typically reach equilibrium with those in the bloodstream as venous blood is arterialized. Diseases that result in impaired diffusion of respiratory gases (e.g., emphysema) often result in fatal hypoxemia before becoming sufficiently severe to cause meaningful limitations in carbon dioxide diffusion, although hypercapnia may develop in these patients by other mechanisms. However, extremes of ventilation–perfusion mismatching can result in a dissociation of arterial (PaCO2) and alveolar (PACO2) carbon dioxide tensions. Even with this limitation, PaCO2 remains the preferred surrogate sample for defining the adequacy of alveolar ventilation. Venous blood PCO2 can be useful in the assessment of ventilatory status in certain settings. As shown in Table  38.1, the carbon dioxide tensions in venous and arterial blood are not the same; however, in a hemodynamically stable patient the difference between the two is suitably constant (~5–7 mmHg) so as to allow reasonably accurate prediction of what current arterial values are likely to be. Venous values also define the maximal value that may be present in arterial samples at that time. For example, if the PvCO2 is 55 mmHg then the PaCO2 is likely to be 49 mmHg and will not be greater than 55 mmHg. Unfortunately, the utility of venous blood samples for defining ventilatory status is strongly dependent on cardiovascular performance. As cardiac output falls, the difference between PvCO2 and PaCO2 becomes progressively larger. In patients in shock or other low‐flow states, venous samples are poor surrogate samples for the assessment of the adequacy of alveolar ventilation. End‐tidal CO2 (ETCO2) is an important means of monitoring ventilation in critically ill animals and encompasses both capnometry (numeric value only) and capnography (waveform representation). Of the various surrogate markers for alveolar CO2, ETCO2, when it is working well, is the closest to actually measuring PACO2. End‐tidal plateau partial pressures of carbon dioxide largely reflect the levels in alveolar gas. End‐tidal CO2 monitoring is the only tool that currently provides continuous, real‐time assessment of the adequacy of alveolar ventilation. In addition, respiratory rate can be determined via this methodology as well. Capnography has the added benefit of allowing waveform inspection, which can be used to monitor for rebreathing, airway obstruction/ bronchospasm, apnea, dyssynchronous alveolar emptying, valve malfunctions, and many other events. Capnography and capnometry can also be useful in determining the status of the cardiovascular system.

Exhaled carbon dioxide originates in the tissues and requires cardiac pumping activity to generate venous return and CO2 delivery. Cardiac arrest is associated with an abrupt decline in ETCO2 levels. Conversely, the adequacy of cardiac compressions can be assessed by capnography or capnometry. Many modern capnography systems (e.g., NICO2®) include a flow disrupter and differential pressure transducers within the same disposable unit that is attached to the patient’s breathing circuit. This allows for monitoring of respiratory mechanics as well as single‐breath capnography (CO2 plotted against cumulative exhaled volume instead of over time). Single‐breath capnography provides additional data relating to airway dead space, gas distribution, and perfusion that cannot be obtained easily (or at all) with standard time‐plotted capnography. The disadvantages of end‐tidal capnography are that it can add considerable apparatus dead space volume in small (24 h) exposure to greater than 60% oxygen can result in pulmonary oxygen toxicity in most mammals although the toxic range varies with species and age. However supplemental oxygen is provided, it is important that the gas be properly conditioned. Conditioning in this sense means appropriately warmed and humidified. The inhalation of improperly conditioned gas is  associated with several adverse effects in humans, including increased nasal airway resistance, damage to the nasal mucosa (both structural and functional, including epithelial metaplasia and keratinization), increased work of breathing, difficulty in subsequent intubation,  patient discomfort, and poor patient compliance. Breathing cold, dry gas from compressed sources can lead to airway dessication (particularly if the nasal passages are bypassed), increases in the tonicity of airway lining fluid, and mast cell degranulation. This airway

A specific prognosis for acute respiratory failure cannot be provided in a meaningful way. Clearly, acute respiratory failure is not good news in any context. However, some causes may be easily resolved and carry a good prognosis (fluid overload) whereas others are challenging to address and carry a quite guarded prognosis (ARDS). The prognosis will naturally vary with the underlying cause. Further, prognosis is impacted by whether a specific, effective therapy is available for that disease process or whether one must rely solely on supportive measures. The issue of prognosis is further complicated by the inherent difficulty in reaching a definitive diagnosis in many cases of acute respiratory failure. Patients are often too unstable to undergo the diagnostics required to definitively identify the underlying cause and thus owners are often required to make difficult decisions based on incomplete information. As mentioned above, retrospective studies of long‐term mechanical ventilation in veterinary patients provide some insights into acute respiratory failure, but represent a highly biased sample of such cases. In one such study, veterinary patients with pulmonary parenchymal disease (i.e., hypoxemic respiratory failure) were able to be successfully weaned 36% of the time, with an overall rate of 22% of cases surviving to hospital discharge. Outcomes were better for patients experiencing hypercapneic respiratory failure, with 50% being successfully weaned and 39% surviving to hospital discharge. Patients with mixed respiratory failure (both hypoxemic and hypercapneic) had the poorest outcomes in this study. Overall, cats fared more poorly than canine patients. Whether this is due to the types of diseases that lead to the need for IPPV in cats or whether cats are at greater risk for complications such as ventilator‐induced lung injury (VILI) remains undetermined. Cats are a smaller species on average than dogs and smaller species develop VILI more readily than do larger species even when identical inspiratory pressures are applied. No direct comparison of VILI incidence in cats versus 3–7 kg dogs has yet been performed.

­Reference 1. Fonfara S, de la Heras Alegret L, et al. Underlying

diseases in dogs referred to a veterinary teaching

hospital because of dyspnea: 229 cases (2003–2007). J Am Vet Med Assoc 2011; 239(9): 1219–24.

39  Acute Respiratory Failure

­Further Reading Fitzpatrick RK, Crowe DT. Nasal oxygen administration in dogs and cats: experimental and clinical investigations. J Am Anim Hosp Assoc 1986; 22: 293–300. Hammer J. Acute respiratory failure in children. Paediatr Respir Rev 2013; 14(2): 64–9. Hopper K, Haskins SC, Kass PH, Rezende ML, Aldrich J. Indications, management, and outcome of long‐term positive‐pressure ventilation in dogs and cats: 148

cases (1990–2001). J Am Vet Med Assoc 2007; 230(1): 64–75. Schneider J, Sweberg T. Acute respiratory failure. Crit Care Clin 2013; 29(2): 167–83. Stefan MS, Shieh MS, Pekow PS, et al. Epidemiology and outcomes of acute respiratory failure in the United States, 2001 to 2009: a national survey. J Hosp Med 2013; 8(2): 76–82.

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40 Mechanical Ventilation Kate Hopper, BVSc, DACVESS, PhD1 and Mathew Mellema, DVM, PhD, DACVECC2 1 2

UC-Davis School of Veterinary Medicine, Davis CA, USA Applaud Medical, San Francisco, CA, USA

A respirator or mechanical ventilator is a therapeutic device that performs some or all of the work of breathing. Ventilators are used in clinical practice to provide short‐ or long‐term support of respiratory function in patients with respiratory disease or failure. Two primary functions of the lung are oxygenation of the arterial blood and the elimination of carbon dioxide from the venous blood in order to provide metabolic substrate, remove metabolic waste, and maintain homeostasis. The ability of the lung to properly oxygenate the pulmonary capillary blood depends largely on several key factors: ●● ●● ●● ●●

adequate replenishment of alveolar gases with inspired air adequate surface area for gas exchange proper matching of ventilation and perfusion maintenance of the structural integrity of the gas exchange barrier.

In contrast, elimination of carbon dioxide is primarily dependent on the repeated movement of fresh gas into the alveoli and removal of portions of alveolar gas, a process known as alveolar ventilation. Respiratory failure can result from diverse disease processes that interfere with one or more of the lung’s primary functions. Patients with respiratory failure can generally be divided into three groups: those with hypoxemic failure, those with hypercapneic failure, and those with mixed respiratory failure, which shares aspects of both of the other two types.

­Indications for Mechanical Ventilation There are three primary indications for mechanical ventilation. The first is persistent severe hypoxemia (PaO2 95%) and a PaCO2 of 35–55 mmHg (35–40 mmHg in patients with intracranial hypertension). In some settings, permissive hypercapnia may be employed. In this approach, PaCO2 is allowed to climb higher to 60–70 mmHg as long as the resultant acidemia does not become too severe (pH 40 breaths per minute

>20 breaths per minute

White blood cells/μL

>19 500 or 3% bands

>16 000 or 3% bands

Source: Adapted from Brady et al. (2000) and Hauptman et al. (1997).

As a consequence, most of the clinical manifestations associated with sepsis and SIRS, with few exceptions, are associated with the response of the innate immune system. It is important to note that the body makes no distinction between these two systems and they function synergistically. For instance, proinflammatory cytokines elicited as a result of an innate immune response help to augment the subsequent or concurrent adaptive response. Innate immunity relies on recognition by the host of some very specific, evolutionarily conserved molecular patterns that are found in association with infection, inflammation, cell death or all of the above. Pattern recognition receptors (PRRs) are a relatively small set of receptors that are able to recognize a wide range of pathogens, including viruses, bacteria, and fungi. Many cell types that participate in immune function, including macrophages, monocytes, dendritic cells, neutrophils, and epithelial cells, express PRRs that allow the rapid recognition of pathogens at the site of infection. Numerous PRRs have been identified (Nod‐like receptors, C‐type lectin receptors, RIG receptors, and RAGE receptors) with remarkable homology across species in all parts of the phylogenetic tree. The fact that these PRRs are found in such diverse species highlights their importance for survival. The most studied and best‐known PRRs are the toll‐ like receptors (TLRs), which are expressed by many cell types including macrophages, dendritic, endothelial, and epithelial cells. TLR activation is generally considered to be proinflammatory and results in the production of antimicrobial peptides, inflammatory cytokines and chemokines, and adhesion molecules, all of which work together to coordinate the innate response. As a result of PRR activation, there is a coordinated response by the immune system to recruit other cells into the fight against infection. This recruitment, the actions of the recruited cells, their associated killing mechanisms, and soluble factors have led some authors to compare the TLRs and other PRRs to keys on a piano. When one key is struck alone, the resultant note does not make a song, but when several keys are played at the

same time and in different combinations, a piece of music emerges. The appropriateness or lack thereof of this process is the central driver in the pathology associated with sepsis and septic shock. Dysregulated Immunity

Once a pathogen has been recognized and the immune response initiated, there are numerous opportunities for dysregulation to occur, adding to morbidity and possibly mortality of the patient. This dysregulated immunity was considered to be exclusively an overexuberant inflammatory response and in fact, the “inflammation only” theory explains most of the clinical signs observed in septic patients. It does not, however, explain all the clinical signs and it has subsequently become clear that hyperinflammation alone does not fully account for the morbidity and mortality experienced by septic patients. A compensatory anti-inflammatory response syndrome (CARS) was proposed as an alternative that may occur in some patients as a result of systemic inflammation. The CARS theory states that in some patients, overcompensation by the anti‐inflammatory arm of the immune system results from a combination of excessive anti-inflammatory cytokine production and, possibly more importantly, from the death and dysfunction of immune cells. This theory helps to explain some of the clinical manifestations of ­sepsis/SIRS. It is convenient to consider these conditions in the extreme as distinct identities but clinically most patients will lie somewhere in the middle. Physiologic Consequences

Patients with sepsis can develop shock that often shares characteristics of hypovolemic, vasodilatory, and cardiogenic etiology due to the actions of inflammatory mediators and their cumulative effect on the circulatory system. In addition to tissue perfusion derangements, patients with sepsis or septic shock can suffer from cytopathic hypoxia or an inability to utilize oxygen at the mitochondrial level. This further complicates understanding the pathophysiology of sepsis and septic shock. Hypovolemia often develops from vomiting, diarrhea or bleeding but can also occur due to accumulation of cavitary effusion or peripheral edema reducing effective circulating volume. Changes in vascular permeability, vascular hydrostatic pressure, decreased colloid osmotic pressure or lymphatic occlusion all occur in septic patients and result in a tendency for fluid to move out of the vascular space. As a result, significant “third spacing” of fluid can occur in the thoracic or abdominal cavities or interstitium of patients with sepsis or SIRS. The net effect of this fluid movement is decreased intravascular volume that contributes to lower cardiac output and impaired oxygen delivery. Besides loss of effective circulating volume, sepsis can induce significant alterations in vascular tone. Because

43  Septic Shock

these patients may have low intravascular volume, compensatory mechanisms are activated in an effort to maintain blood pressure. To minimize the effect of hypovolemia on organ systems in nonseptic patients, systemic vascular resistance generally increases to compensate for the lower cardiac output and maintain organ perfusion. In sepsis, however, numerous factors counteract these compensatory mechanisms to the detriment of the patient. This can result in decreased effectiveness of vasoconstriction or, worse, to pathologic dilation of the vessels. The most well‐understood vasodilatory agent in sepsis is nitric oxide (NO). In sepsis and septic shock, inducible nitric oxide synthase (iNOS) produces supraphysiologic levels of nitric oxide that interfere with myocyte calcium metabolism and impair the ability of vascular smooth muscle cells to contract. The end‐result is vasodilation with a characteristically poor response to vasoconstrictive stimuli, including reduced vascular response to catecholamines. Additionally, the surplus of nitric oxide leads to production of reactive oxygen species, including peroxynitrite, which are directly toxic in the local environment and act to cause further damage to endothelium and smooth muscle myocytes. Nitric oxide is also responsible to some degree for the cardiac depression found in patients with sepsis or septic shock. Endocrine abnormalities occur in sepsis and can also adversely affect the ability of the vascular tree to respond to stimuli. With the additional demand for vasoconstriction, the neurohypophysis (posterior pituitary) depletes its stores of vasopressin and is unable to meet the demand, leading to a well‐documented deficiency in some septic patients. A distinct cardiac failure component is present in some human patients with septic shock but is of unknown clinical relevance in veterinary medicine. It was believed that a circulating myocardial depressant factor was present in the blood of septic patients that decreased cardiac function. This myocardial depressant factor was subsequently identified as tumor necrosis factor (TNF)‐ alpha which, together with interleukin (IL)‐1, has been associated with cardiac myocyte dysfunction. TNF‐alpha and IL‐1, both potent inducers of iNOS, are now known to be the offending molecules that lead to sepsis‐induced cardiac dysfunction. If decreased systolic function is coupled with decreased systemic vascular resistance, as seen in septic shock, the cardiac output often stays the same or even increases although tissue perfusion is severely compromised. However, if vasoconstriction persists, as seen in uncomplicated or severe sepsis, decreased systolic function can lead to a significant reduction in cardiac output. Sepsis‐induced endothelial injury affects the lungs by allowing neutrophil migration into the lung parenchyma

and increased vascular permeability that impairs gas exchange. This disease has been termed acute lung injury (ALI) or acute respiratory distress syndrome (ARDS), depending on severity, with ALI describing a less severe manifestation than ARDS. Both are characterized by acute‐onset bilateral pulmonary infiltrates with no evidence of left‐sided heart failure. Classification is further determined by the degree of hypoxemia. For more information on ALI/ARDS the reader is referred to Chapter 38. Significant alterations in normal gastrointestinal function, motility, and vascular permeability result from inflammatory mediators, altered blood flow, and oxidative stress. This may allow exposure of the bloodstream to the luminal contents of the gastrointestinal (GI) tract and allow bacteria access to the bloodstream or lymph, although the clinical significance and actual incidence of bacterial translocation are unknown. Hepatic dysfunction in sepsis is manifested as hypoalbuminemia, icterus, hypoglycemia, and coagulation abnormalities and decreased detoxifying ability. The underlying cause of hepatic dysfunction is unknown but may result from hypoperfusion of the liver, or may be directly due to the systemic inflammatory process (i.e., inflammatory mediators) as many of the constitutive functions of the liver are downregulated as part of the acute phase response. Coagulation abnormalities can be significant and can lead directly to the death of the patient. Sepsis‐induced coagulopathy is a manifestation of the procoagulant nature of almost all proinflammatory mediators. In general terms, the procoagulant state in sepsis causes generation of microthrombi that result in inflammation within the vascular beds in which they lodge. This systemic activation of the coagulation system finally results in the consumption of coagulation factors and in the later stages ultimately induces a hypocoagulable state. This scenario has been termed disseminated intravascular coagulation (DIC) and continues to be a significant clinical complication in the management of sepsis. Unfortunately, due to clinical limitations in identifying patients in the procoagulant phase of DIC, this complication is often found in the last stages when profound bleeding tendencies predominate and it is extremely difficult to manage or reverse. The discovery of renal dysfunction or acute kidney injury is an easily identified marker of multiple organ dysfunction syndrome. Acute kidney injury results from decreased renal blood flow secondary to systemic hypotension; alterations in renal blood flow from vasoactive substances, microthromboembolism, and poorly understood renal cellular apoptosis are also believed to contribute. Patients may exhibit a progressive increase in creatinine and decreased urine production despite normal intravascular fluid volume. Even small changes in

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serum creatinine (within the reference range) can indicate renal impairment and emerging multiorgan failure. The central nervous system manifestations of sepsis and septic shock may be due to the effects of systemic hypotension and decreased cerebral perfusion or may be related to the direct effects of sepsis‐induced cytokines. Alterations in cerebral blood flow, increased capillary leakage, and disruption of the blood–brain barrier are thought to be mechanisms contributing to brain dysfunction in sepsis. In addition to primary intracranial causes, sepsis‐induced hypoglycemia can cause neurologic dysfunction. The clinical signs seen with sepsis and septic shock are predominantly alterations in mentation (i.e., obtundation, stupor, coma) and seizures.

­Epidemiology The exact incidence of sepsis in veterinary medicine is unknown. However, sepsis is likely a common occurrence in clinical practice and remains among the most difficult to treat conditions. According to the NIH, about 750 000 people become septic in the United States each year with a mortality rate of 28–50%. In fact, the rates of hospitalization for human sepsis have become higher than those of myocardial infarctions. The death rate from sepsis eclipses that of prostate cancer, breast cancer, and AIDS combined. Despite the difficulty in diagnosing sepsis, there are common veterinary diseases that can cause a patient to develop sepsis. Parvoviral enteritis, pyometra, pneumonia, and septic peritonitis are just a few of the diseases responsible for sepsis and septic shock in dogs and cats. Septic peritonitis occurs in both cats and dogs and most commonly results from perforation of the GI tract. Because of the propensity of dogs to ingest foreign objects, septic peritonitis occurs more commonly in that species. Most frequently, perforation of the GI tract occurs from ingestion of foreign material that physically obstructs aboral flow or damages the walls of the GI tract to the point of rupture. Perforation of the stomach and duodenum can occur secondary to nonsteroidal antiinflammatory drug (NSAID) administration. Neoplasia can cause perforation in any part of the bowel. Disease of organ systems other than the GI tract can cause septic peritonitis. This occurs most frequently when a normally sterile organ becomes infected and the infection extends into the abdomen via perforation or through translocation of bacteria to the blood or lymph. Extension of infection from the urogenital tract can occur because of prostatitis, prostatic abscess, pyometra, pyelonephritis or rupture of the urinary bladder containing infected urine or calculi. Septic bile peritonitis results

from perforation of an infected biliary tree, most frequently the gallbladder. Less frequently, infection or abscessation of organs such as the liver or spleen, or necrotic tissue secondary to strangulation or torsion is the source. Spontaneous septic peritonitis can occur in cats with no evidence of gastrointestinal compromise or other identifiable source. Numerous other diseases are capable of causing sepsis in dogs and cats. Penetrating trauma from animal bites is a common occurrence and contamination from bite wounds can result in sepsis. Blunt trauma without penetrating injury (e.g., being struck by a vehicle or falling from a height) frequently results in significant soft tissue injury that can result in sepsis if infection of the damaged tissue occurs.

­Signalment All species, breeds, sexes, and ages can develop sepsis. Consequently, there is no signalment specific to sepsis but there are populations that are susceptible to diseases that can be the underlying cause of sepsis. For example, young, unvaccinated puppies are at risk of contracting Parvoenteritis while intact female dogs and cats may develop pyometra. Dogs that develop septic peritonitis tend to be younger with indiscriminate eating habits that ingest foreign material, resulting in perforation of the GI tract. A report including nearly 75 000 dogs identified some distinct breed‐ and age‐related differences in all‐cause mortality, with younger patients having a significantly higher likelihood of dying of infectious disease or trauma. Geriatric dogs with chronic, progressive diseases necessitating treatment with NSAIDs, corticosteroids, or other immunosuppressive medications are also at higher risk than the general population for developing GI tract perforation. Ruptured neoplasia of the GI tract occurs in both geriatric dogs and cats. Lifestyle of the pet can significantly affect the likelihood of developing sepsis. Pets that are primarily outdoors or that spend a significant amount of time outdoors and unsupervised are more likely to be exposed to injuries that can result in sepsis.

­History and Clinical Signs Patient historical information often relates to the underlying cause of sepsis. For example, animals with septic peritonitis may have historical complaints such as vomiting, abdominal pain or hemorrhagic diarrhea while patients with sepsis secondary to pyometra often

43  Septic Shock

have polyuria, polydipsia, and vulvar discharge as part of their history. Many dogs with septic peritonitis secondary to gastrointestinal perforation will have a history of NSAID or corticosteroid administration. Some of the etiologies of sepsis are subacute to chronic in nature and subsequently the historical complaints may be somewhat vague. Regardless of the underlying cause of sepsis, it is common for owners to report decreased activity levels and loss of appetite for varying amounts of time prior to presentation. Travel history, exposure to any and all medications (including common over‐the‐counter drugs), housing (indoor vs outdoor), and exposure to other animals are all important historical facts to elucidate. Just as sepsis/SIRS occurs on a spectrum, the clinical signs associated with sepsis/SIRS vary depending on the severity of the condition. By definition, all animals with sepsis have clinical signs consistent with SIRS. Additional clinical signs that may be present include lethargy, recumbency, vocalization, or seizures, with some patients even presenting moribund. Once septic shock develops, clinical signs vary but are generally related to deficits in oxygen delivery. Patients with septic shock can have two different presentations: they can be in hyperdynamic (compensated) or hypodynamic (decompensated) shock. Hyperdynamic shock implies that there is greater than normal cardiac output with vasodilation and these patients will be tachycardic, have red mucous membranes, bounding femoral pulses, and warm extremities. Patients in the hypodynamic phase of shock can be tachycardic or bradycardic, have weak pulses, pale to gray mucous membranes, and cold extremities.

­Diagnosis The diagnosis of septic shock is made by fulfilling the SIRS criteria and key hemodynamic parameters in conjunction with identification of an underlying infection. Patients in septic shock will show similar physical exam findings to patients in shock for other reasons, but there will generally be additional abnormalities related to the underlying cause of sepsis. For instance, a patient with septic peritonitis will have traditional signs of shock (tachycardia, weak pulses, and pale mucous membranes) but will also often have abdominal pain and peritoneal effusion as evidenced by abdominal distension with the possibility of a palpable fluid wave. When presented a patient with suspected sepsis or septic shock, it is important to first gather as much data as you can. The minimum database consists of a complete blood count, biochemistry profile, and urinalysis.

Abnormalities in the minimum database generally reflect the underlying disease although specific sepsis‐related changes could be present. An inflammatory response is expected on the hemogram, although individual patients may demonstrate high, normal, or low leukocyte counts with or without toxic change or presence of band neutrophils. Patients may have a regenerative or nonregenerative anemia and may be thrombocytopenic. The biochemistry profile can often be quite abnormal. Septic patients have a significant inflammatory component to their disease that can affect blood protein levels dramatically. Hypoalbuminemia is found frequently in septic patients as albumin loss into exudates occurs concurrently with downregulation of albumin production during the acute phase response. Other abnormalities identified on the biochemistry profile may include hypoglycemia, hyperbilirubinemia, hypocalcemia, and elevated creatinine and blood urea nitrogen. Of these, hypoglycemia is of particular concern since it contributes to morbidity or mortality if not identified and corrected rapidly. Imaging studies are often key to the identification of the source of sepsis. Plain radiographs of the chest and abdomen are useful for identifying possible sources of sepsis and the finding of unstructured interstitial to alveolar pulmonary infiltrates, pleural effusion, obstructive intestinal pattern, free abdominal air, or loss of abdominal detail indicating free fluid should be considered significant and additional confirmatory diagnostic tests should be pursued. Collection of free abdominal fluid, if present, is often the first step in diagnosing septic peritonitis and can be accomplished via blind four‐quadrant paracentesis or ultrasound‐guided needle aspiration. Ultrasound allows the clinician to bring imaging to the bedside and perform a rapid evaluation of the thoracic and abdominal cavities. This can help with the initial identification of disease, and can provide a method to monitor progression. Ultrasound is particularly useful for identifying pockets of fluid within the chest and abdominal cavities that can then be sampled. In clinical practice, ultrasound is more effective at identifying free fluid than is blind four‐quadrant paracentesis, making it an objective and effective tool in the ER and ICU. Once fluid is obtained, cytologic evaluation can ­confirm the diagnosis of septic peritonitis or prompt the  clinician to investigate further. Identification of intracellular bacteria signifies that bacteria present in the peritoneal cavity are being phagocytized by inflammatory cells and indicates infection rather than sample contamination. The finding of even one intracellular bacterium is significant and is an indication for immediate

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surgical exploration. While identification of even one intracellular bacterium confirms the diagnosis of septic peritonitis, the absence of this finding does not rule out sepsis. If the fluid obtained is highly cellular (composed of mixed inflammatory cells) or is turbid, flocculent or malodorous then further investigation may be needed. Comparison of the biochemical values, specifically lactate and glucose, of abdominal fluid with blood can be invaluable for aiding in the diagnosis of septic peritonitis. The comparison of glucose and lactate concentrations of abdominal fluid and blood is both sensitive and specific for differentiating septic peritonitis from other causes of peritoneal effusion. Under normal conditions, the concentration of glucose and lactate in peritoneal fluid is essentially the same as that of blood. As infection progresses within the peritoneal cavity, bacteria and white blood cells utilize glucose for energy and produce lactate as a by‐product. If the lactate concentration of abdominal fluid is 2.0 mmol/L higher than that of the blood or if glucose is 20 mg/dL lower in the effusion than in the blood, a presumptive diagnosis of septic peritonitis can be made and surgical exploration should be recommended. Monitoring coagulation assists with identification of needed therapies rather than identification of sepsis or septic shock and coagulation assays may be normal or abnormal in these patients. A prothrombin time (PT) or partial thromboplastin time (aPTT) that is >50% prolonged indicates coagulopathy. Fibrinogen levels may be normal, low or high, depending on the chronicity of the disease. Patients that have a prolonged disease course tend to have higher fibrinogen levels, but it is not uncommon to find low fibrinogen due to active consumption. D‐dimers are often elevated as a result of the underlying condition or inappropriate coagulation (i.e., DIC). Identification of three of the following in a patient identifies DIC: prolongation of PT or aPTT, hypofibrinogenemia, thrombocytopenia, elevation of fibrin degradation products (FDPs) or D‐dimers, or documented antithrombin deficiency.

­Treatment Treatment of sepsis and septic shock must be aggressive and aimed at eliminating the underlying cause. In broad terms, treatment of sepsis and septic shock can be divided into several distinct components: resuscitation, source control (eliminating the cause of sepsis), antimicrobial therapy, and supportive care. The resuscitation of a patient with severe sepsis or septic shock is different from that of patients with other forms of shock. This is due to

the multifactorial nature of the development of shock in the septic patient, meaning that in any one patient, low systemic vascular resistance, cardiac dysfunction, and hypovolemia may all be coexistent and may all contribute to failure of oxygen delivery (i.e., shock). Resuscitation Complicating septic shock resuscitation is the fact that vascular permeability may be increased, which can decrease the efficacy of crystalloid resuscitation or lead to overresuscitation and subsequent edema formation. The use of goal‐directed therapy, consideration of other fluids options, and use of vasoactive agents as well as fastidious reevaluation can improve the efficacy of resuscitation. The most important initial therapy in the resuscitation of a patient with septic shock is the administration of appropriate doses of IV fluids to restore effective circulatory volume and improve global perfusion. The appropriate amount and character of the fluids that should be administered are a matter of debate. The author prefers to give an initial bolus of 20–30 mL/kg of crystalloid fluids such as lactated Ringer’s solution or 0.9% sodium chloride. This initial dose serves to act as a fluid challenge and will help the clinician determine if continued fluid resuscitation may be beneficial. If a positive response is seen, this dose can be repeated 2–3 more times until one of three endpoints occurs: signs of shock resolve, there is no further improvement in perfusion parameters or volume overload becomes evident. Signs of volume overload include development of peripheral or pulmonary edema or measurement of central venous pressures greater than 10 cmH2O. Colloidal solutions are commonly used as a primary or secondary resuscitative fluid in veterinary patients in shock. Colloids have several theoretical benefits, but one of the most important is that they discourage formation of tissue edema since they do not move out of the vascular compartment as quickly as crystalloid fluids. If colloids are used, total doses of 20 mL/kg are typically broken into aliquots similar to the technique used with crystalloids and are administered in incremental 5–10 mL/kg boluses until endpoints are achieved. Hetastarch (Hespan®, Hextend™) is the most commonly used colloid in veterinary patients and has replaced the dextran solutions that were in use previously. A newer generation of hydroxyethylstarchs, generally classified as tetrastarchs, is also available with a similar dosing schedule. Patients in septic shock, by definition, have hypotension that is not responsive to fluid loading alone, meaning that vascular expansion has been completed without resulting in resolution of shock. Determining when patients are adequately volume expanded is difficult.

43  Septic Shock

One method for determining if volume loading has been adequate is measurement of central venous pressure, with a target of 8–12 cmH2O. Many of these patients will not have a central line in place, in which case lack of improvement in blood pressure following IV fluid boluses can  be  interpreted as evidence that volume expansion is complete. When this happens, vasopressors are needed to increase systemic vascular resistance in an effort to increase blood pressure and maintain vital organ perfusion. There is no consensus on which vasopressor is most effective and selection is largely based on clinician preference. The most commonly used first‐line vasopressors are dopamine (5–20 μg/kg/ min) and norepinephrine (0.05–0.3 μg/kg/min). If normotension is not achieved at the upper end of the dose range for a single vasopressor then the addition of a second agent is needed. Typical secondary vasopressors include vasopressin (0.01–0.04  U/kg/h) or norepinephrine if not already used as a first‐line agent. While vasopressors function to increase systemic vascular resistance and improve perfusion of tissue beds, patients with sepsis or septic shock often have concurrent systolic dysfunction. Positive inotropes are drugs that increase inotropy (contraction strength) and chronotropy (contraction rate), resulting in an increased stroke volume and heart rate, and consequently cardiac output. The most commonly used inotropic medication is dobutamine. If evidence of systolic failure is present then dobutamine should be started (5–15 μg/kg/min) and the dose titrated up every 15 minutes until endpoints are reached or the top of the dose range is reached. If dobutamine fails to result in resolution or improvement in shock then the addition of a vasopressor should be considered. It should be remembered that many adrenergic drugs are positive inotropes in addition to being vasopressors (ex. dopamine and norepinephrine) and the primary effect is often dependent on the dose administered. For more information about resuscitation endpoints, the reader is directed to Chapter 41. Source Control Source control is important for the resolution of sepsis and septic shock and should be performed as soon as possible after the source is identified to improve the likelihood of recovery. Source control almost always requires some degree of surgical intervention, ranging from lancing of an abscess to resection of perforated bowel. Good source control for wounds includes early and aggressive debridement of devitalized or infected tissue. Dogs with necrotizing fasciitis may require radical resection of affected tissue and the use of vacuum bandaging or

application of wet to dry bandages for continued debridement following the initial surgical intervention. If the septic source is an abdominal viscus such as the uterus or bowel then laparotomy should be performed as soon as the patient is stable enough for anesthesia with the goal of eliminating ongoing bacterial contamination with dilution or removal of already affected tissue. Similarly, if the septic source is in the thorax (ex. pyothorax or lung abscess) then thoracotomy should be recommended. Pyothorax can be managed conservatively with placement of thoracostomy tubes acting as source control mechanisms and is reported to be effective 71% of the time. The decision to manage cases with thoracostomy tubes versus surgical intervention depends on geography, as well as clinician preference or training. In areas of the country where inhaled grass awns from foxtails are common, the frequency of surgery is expected to be higher than that of an area in which those plants are not found. Lung abscesses must be managed with lung lobectomy. For cases with severe sepsis or septic shock secondary to pyelonephritis, source control interventions to consider include percutaneous renal pelvis centesis or nephrectomy. Antimicrobial Therapy The early administration of appropriate antibiotics is key for successful treatment of patients with sepsis and septic shock. Every effort should be made to collect culture samples before antibiotics are administered; however, one cannot wait indefinitely to initiate treatment and the goal should be to begin antibiotics within one hour of recognition of septic shock. In patients with sepsis without shock, it is often much more feasible to withhold antibiotic therapy until appropriate cultures have been obtained. Ideally, the antibiotic administered would be effective against the offending organism in question. Unfortunately, in most cases the antibiotic sensitivities or the identity of the organism are unknown prior to initiation of antimicrobial therapy. Even when appropriate culture samples are collected, the results are delayed by 48–72 hours or  longer. Therefore, the initial antimicrobial selection should be effective against bacteria in all four quadrants (gram positive, gram negative, aerobic, and anaerobic). This usually requires the co‐administration of two or more antibiotics. Unless a clear contraindication exists, the dose of all antibiotics should be at the high end of the dose range, they should be given IV, and penetrate the desired tissue bed. Examples of effective antibiotic combinations include combining potentiated penicillins (ex. ampicillin/sulbactam 30–50 mg/kg q8h or ticarcillin/ clavulanate 40–50 mg/kg q8h) with fluorquinolones

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(ex.  enrofloxacin 10–mg/kg q24h) or aminoglycosides (ex. gentamicin 6–8 mg/kg q24h). Once culture results are returned, the antibiotic spectrum can be narrowed based on the microbial sensitivity pattern. The duration of therapy required by septic patients is unknown and difficult to determine. A useful guide is to continue antibiotics therapy until one week after clinical resolution of infection. Supportive Care There are numerous additional components that are integral to the recovery of any critically ill animal. In ­septic patients, nutrition, monitoring, and high‐quality nursing care cannot be emphasized enough. Diligent monitoring is necessary for these animals as their status can change quickly and with little notice. Heart rate, respiratory rate, respiratory effort, blood pressure, hemoglobin saturation, urine production, and blood glucose should be checked at regular intervals. Due to the catabolic state that occurs during sepsis, nutritional support is a very important component of treatment for these patients. In addition to helping prevent hypoglycemia, nutrition allows for some anabolic activity to occur and enables synthesis of components of the immune system. In patients with severe sepsis or septic shock, dextrose supplementation is often necessary to treat hypoglycemia. When possible, it is always preferable to provide these patients with enteral rather than parenteral nutrition. Early enteral nutrition (defined as consistent caloric intake within the first 24 hours) has been shown to decrease length of hospitalization in dogs with septic peritonitis. This can be accomplished through voluntary alimentation by the patient or through the use of feeding tubes. The use of feeding tubes is becoming more common in the management of critically ill patients as more diets specifically formulated to be used in small‐caliber tubes are developed. While most feeding tube options require heavy sedation or general anesthesia for placement, nasoesophagal tubes are small‐caliber tubes that can be placed with little to no sedation and are therefore very useful for providing enteral nutrition to those patients that are unwilling to eat voluntarily. Although patients with sepsis as a rule have a negative energy balance, the goal with nutritional support should be to meet the resting energy requirement. There is little to no evidence that hyperalimentation improves clinical outcomes and it has been associated with development of complications including feeding intolerance and hyperglycemia.

Critical illness‐related corticosteroid deficiency (CIRCI) is a multifactorial deficiency in circulating ­cortisol in critically ill patients. This condition was first  described when it was found that a subset of patients with refractory hypotension improved following administration of physiologic doses of corticosteroids. Initially CIRCI was diagnosed by determining if patients responded appropriately to an adrenocorticotropic hormone (ACTH) stimulation test. It was found, however, that even some patients with normal response to ACTH stimulation benefited from corticosteroid supplementation. This led to the recommendation for supplementation of hydrocortisone in critically ill human patients with unresponsive hypotension regardless of ACTH stimulation results and in fact, ACTH stimulation tests are no longer considered necessary for the diagnosis of CIRCI. Corticosteroid administration should be considered only if fluids and vasopressors do not correct hypotension. If used, the author prefers hydrocortisone 1–4 mg/kg/ day divided q12h or once every 24 hours. Corticosteroids should be discontinued when the patient is no longer vasopressor dependent.

­Prognosis The prognosis for septic shock is tremendously variable in both veterinary and human patients. Severity and reversibility of the underlying disease process as well as the presence of comorbid conditions affect prognosis of sepsis and septic shock. However, it must be recognized that the farther along the septic spectrum (toward development of septic shock) that a patient goes, the worse the prognosis will be. Veterinary survival data are difficult to interpret as the practice of euthanasia complicates the evaluation of outcome, but survival rates of 50–60% are commonly reported. This correlates with studies showing mortality rates of nearly 50% in humans with septic shock. Patients with sepsis or septic shock have significant disease and regardless of the cause, this condition is considered serious. Every effort should be made to set realistic owner expectations with the understanding that the prognosis is guarded to poor (especially in the case of septic shock). The financial and emotional cost of treating a dog with sepsis can be very high. The long‐term prognosis for patients that survive sepsis and septic shock is unknown although good long‐term outcomes appear to be common.

43  Septic Shock

­Further Reading Banta JE, Joshi KP, Beeson L, Nguyen HB. Patient and hospital characteristics associated with inpatient severe sepsis mortality in California, 2005–2010. Crit Care Med 2012; 40(11): 2960–6. Bentley AM, Otto CM, Shofer FS. Comparison of dogs with septic peritonitis: 1988–1993 versus 1999–2003. J Vet Emerg Crit Care 2007; 17(4): 391–8. Bonczynski JJ, Ludwig LL, Barton LJ, Loar A, Peterson ME. Comparison of peritoneal fluid and peripheral blood pH, bicarbonate, glucose, and lactate concentration as a diagnostic tool for septic peritonitis in dogs and cats. Vet Surg 2003; 32: 161–6. Bone RC, Balk R, Cerra F, et al. Definitions for sepsis and organ failure and guidelines for the use of innovative therapies in sepsis. Chest 1992; 101(6): 1644–55. Boothe HM, Howe LM, Boothe DM, et al. Evaluation of outcomes in dogs treated for pyothorax: 46 cases (1983–2001). J Am Vet Med Assoc 2010; 236: 657–63. Brady CA, Otto CM, van Winkle TJ, King LG. Severe sepsis in cats: 29 cases (1986–1998). J Am Vet Med Assoc 2000; 217(4): 531–5. Conti‐Patara A, de Araújo Caldeira J, de Mattos‐Junior E, et al. Changes in tissue perfusion parameters in dogs with severe sepsis/septic shock in response to goal‐directed

hemodynamic optimization at admission to ICU and the relation to outcome. J Vet Emerg Crit Care 2012; 22: 409–18. Dellinger RP, Levy M, Rhodes A, et al. Surviving Sepsis Campaign: international guidelines for management of severe sepsis and septic shock: 2012. Crit Care Med 2013; 41(2): 580–637. Fleming JM, Creevy KE, Promislow DEL. Mortality in North American dogs from 1984 to 2004: an investigation into age‐, size‐, and breed‐related causes of death. J Vet Intern Med 2011; 25: 187–98. Hauptman JG, Walshaw R, Olivier NB. Evaluation of the sensitivity and specificity of diagnostic criteria for sepsis in dogs. Vet Surg 1997; 26(5): 393–7. Lisciandro GR. Abdominal and thoracic focused assessment with sonography for trauma, triage, and monitoring in small animals. J Vet Emerg Crit Care 2011; 21: 104–22. Liu DT, Brown DC, Silverstein DC. Early nutritional support is associated with decreased length of hospitalization in dogs with septic peritonitis: a retrospective study of 45 cases (2000–2009). J Vet Emerg Crit Care 2012; 22(4): 453–9. Takeuchi O, Akira S. Pattern recognition receptors and inflammation. Cell 2010; 140(6): 805–20.

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44 Disorders of Heat and Cold Sarah Allen, DVM, DACVECC Massachusetts Veterinary Referral Hospital, Woburn, MA, USA

­Normal Thermoregulation Body temperature is closely maintained in mammals around an optimal set point at which ideal cellular function can occur. This is a complex process that ultimately results in a balance between heat production and heat loss. The hypothalamus acts as the main regulator of this process with multiple sensors throughout the body in the skin, thoracic and abdominal viscera, and spinal cord. The body can be viewed as two thermal divisions: the core and peripheral compartments. The core compartment is well perfused with a mostly constant temperature. The peripheral compartment is composed of the extremities and peripheral temperature is dependent on heat transmission from the core via blood flow, heat conduction from adjacent tissues, and heat loss to the environment. Heat is generated by muscle activity and metabolism, with the brain, heart, and abdominal organs being the main generators of metabolic heat. Changing blood levels of thyroxine, epinephrine, and norepinephrine can change the basal metabolic rate, and subsequently body heat. Increased muscular activity (i.e., shivering) is also used as a means of generating heat. Heat retention occurs by physiologic responses including piloerection and peripheral vasoconstriction, as well as behaviors such as seeking shelter and warmth and curling up. In order to prevent hyperthermia, methods of heat dissipation are necessary to maintain an ideal body temperature. The four main mechanisms of heat loss from the body are evaporation, conduction, convection, and radiation. Evaporation is loss of heat from moisture on the body surface. In canine and feline patients, the main source of heat loss by the evaporative route is from the respiratory tract, with panting providing a means of increasing heat loss from the body. Conduction is the transfer of heat from body surfaces to objects contacting

the body, such as exam tables or cage floors. Convection occurs when body heat is transferred to the air surrounding the body and radiation is the transfer of heat from the body to distant surfaces without direct bodily contact. In small animals, conduction and convection are the primary means of heat dissipation. Individual patient factors that contribute to increased heat loss may include increased relative surface area in a neonate, decreased fat insulation in a thin patient, inability to prevent excessive heat loss to a conductive surface or decreased heat seeking due to weakness or altered mentation.

­Hypothermia Hypothermia is defined as body temperature of less than 38 °C (100.4 °F) and is classified by severity as mild (32–37 °C/89.6–98.6 °F), moderate (28–32 °C/82.4–89.6 °F), or severe (28 °C/82.4 °F), active external rewarming is required with the main efforts aimed toward the core region rather than the extremities. Once hypothermia becomes severe (42 °C/ 107.6 °F), prolonged contact time between patient and heat source is allowed, or there is a lack of insulation between the heat source and patient. Prognosis Rewarming complications can affect the prognosis of hypothermic patients.

­Hyperthermia Hyperthermia is defined as an increase in body temperature greater than 38 °C (100.4 °F) and can be due to an alteration of the set point of the hypothalamus to a higher temperature with deliberate efforts of the body to raise itself to that temperature (true fever) or can result from loss of the thermoregulatory balance of the body in which heat production is increased or heat loss is inhibited but the set point remains the same. Hyperthermia can be induced by extreme muscular activity. Sustained exercise, especially in an underconditioned or overweight patient, sustained seizure activity or frequent cluster seizures can induce severe hyperthermia if not controlled. An uncommon cause of hyperthermia in dogs and cats is malignant hyperthermia. This is a condition of disturbed calcium metabolism that is initiated by drugs, most commonly volatile anesthetic gases, that results in

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a myopathy with increased metabolic heat production. Clinical signs include elevated body temperature and often muscle rigidity. In patients under anesthesia, an increase in end‐tidal CO2 may be noted due to the increased metabolic rate associated with this condition. Various pathologic conditions, including endocrine disorders (hyperthyroidism, pheochromocytoma), can induce an increase in metabolic rate and heat production, as well as decrease heat loss due to peripheral vasoconstriction. Hypothalamic lesions that result in an increase in the temperature set point will create hyperthermia. Prolonged extreme elevations in body temperature can increase the metabolic rate and demand for oxygen, water, and calories that may exceed the body’s ability to supply them. When this occurs, protein denaturation occurs and resulting cellular dysfunction is manifested as cardiac arrhythmias, liver dysfunction, kidney failure, and damage to the gastrointestinal barrier with secondary bacterial translocation. Acid–base and electrolyte derangements occur due to dysfunction at the cellular and organ level. Muscle damage from high temperatures and hypoxia causes rhabdomyolysis, hyperkalemia, and myoglobinuria, leading to further renal injury. Significant alterations in systemic coagulation manifest as disseminated intravascular coagulation (DIC) with thrombosis adding to organ dysfunction. Consumption of clotting factors results in inappropriate bleeding that may produce clinically significant anemia. Treatment Treatment of hyperthermia should be directed at correcting the underlying cause and returning the body temperature to normal through the use of active cooling. Active cooling techniques include placing the patient in front of a fan to improve convective heat transfer, removing bedding to allow contact with cool cage or floor surfaces to maximize conductive heat loss, and application of alcohol to the foot pads to enhance evaporative heat dissipation. These techniques are generally sufficient for treatment of hyperthermia of less than 41 °C (106 °F) and should be started on animals with hyperthermia greater than 40 °C (104 °F). If these mechanisms are ineffective or if hyperthermia is more severe, then dousing the patient (wet to the skin, not just the hair coat) in cool water and placing them in front of a fan is the best way to transfer heat from the core to the skin and subsequently to the environment. Ice water should be avoided as it will induce peripheral vasoconstriction and inhibit heat transfer from the skin and delay core cooling. Application of ice water may also induce shivering which can result in heat generation. Cooling should be closely monitored and active cooling stopped when the core body temperature is still above 39 °C (102.2 °F)

Hyperthermia secondary to seizure activity requires aggressive medical management of the seizure activity with antiepileptic drugs (diazepam 0.5 mg/kg IV) and active cooling. Treatment of malignant hyperthermia includes stopping the causative medication, active cooling, and administration of dantrolene (2.5–5.0 mg/kg IV).

­Fever Hyperthermia in the form of fever is part of the systemic acute phase response to a pathogen or tissue injury and is aimed at decreasing the ability of infectious agents to replicate and survive. The response may be triggered by a variety of infectious agents including bacteria, fungi, and viruses, as well as tissue injury from trauma or surgery. Neoplastic cells are also capable of inducing fever. A substance that is created by the body and induces fever is an endogenous pyrogen, the most significant of which are the cytokines interleukin (IL)‐1, Il‐6, and tumor necrosis factor (TNF)‐alpha. A substance released by an infectious agent that induces fever is known as an exogenous pyrogen and includes lipopolysaccharide (LPS) of gram‐negative bacterial cell walls. LPS and other exogenous pyrogens bind to immune cells and result in cytokine release and initiate the acute phase response. Infectious agents also create tissue damage that leads to the release of cytokines. Once released, cytokines stimulate the arachidonic acid pathway and result in the production of prostaglandin‐E2 (PGE2). PGE2 is the main mediator of the fever response through its action on the hypothalamus that results in vasoconstriction to prevent heat loss and catecholamine release (among other mechanisms) to increase thermogenesis. The cause of fever may be obvious based on history and physical exam, as in the case of an infected wound or surgical site, or an abscess from a bite wound. If not ­readily apparent, an extensive work‐up may be required to identify the underlying cause. A tiered approach to working up a fever is often proposed, starting with a minimum database including complete blood count, chemistry profile, and urinalysis. Thoracic and abdominal radiographs with ultrasound are also often included in the initial work‐up. If these tests do not provide a diagnosis, further diagnostics including relevant ­ infectious disease titers (based on geographic area), ­ urine culture, blood cultures, echocardiogram, joint taps, and cerebrospinal fluid analysis may be required. Treatment Identifying and eliminating the cause of true fever is the mainstay of treatment. Therapeutic treatment with IV fluids to maintain hydration and electrolyte balance may

44  Disorders of Heat and Cold

be necessary. Broad‐spectrum antibiotics are recommended if a bacterial infection is suspected and when possible, antibiotic therapy should be deescalated based on culture results. Prolonged fever with no response to antimicrobials and a thorough, inconclusive work‐up may indicate an immune‐mediated process that requires immunosuppression, a viral process that requires continued monitoring and supportive care, or a neoplastic process that has yet to be identified. Treatment with antipyretics that inhibit prostaglandin synthesis, such as nonsteroidal antiinflammatory drugs (NSAIDs), is controversial as inhibiting the fever may allow continued propagation of an infectious agent. Possible side‐effects of these medications in each individual patient should also be considered carefully prior to their administration. Although uncommon, true fevers exceeding 41 °C (106 °F) require more aggressive intervention with active cooling to prevent organ damage and other systemic effects.

­Heat Stroke Heat stroke is probably the most common cause of life‐ threatening hyperthermia (>40 °C/104 °F) encountered in veterinary medicine and results from inadequate heat dissipation from the body. Classic heat stroke is caused by exposure to high environmental temperatures, which overwhelm the body’s ability to offload heat by means of convection and radiation. The most common situation causing classic heat stroke occurs when an animal is locked in an enclosed vehicle where the temperature can rapidly increase past the outside ambient temperature due to solar heat and poor interior ventilation. This situation develops more rapidly in larger dogs that produce greater body heat and have less relative surface area for heat dissipation than smaller dogs. Exertional heat stroke is the result of physical activity in a hot and/or humid environment and is often seen in working dogs. Increased environmental humidity decreases respiratory evaporative heat loss while increased work of breathing promotes heat production and further exacerbates this condition. Many cases of exertional heat stroke occur early in the warm weather season due to lack of acclimatization to the change of temperature. Dogs with altered means of respiratory evaporative heat loss, such as brachycephalic dogs and dogs suffering from laryngeal paralysis, may be more vulnerable to exertional and classic heat stroke. Obesity and a thick hair coat can further decrease the ability to dissipate heat from the body in both forms of heat stroke. Some dogs may have a genetic resistance to heat stroke due to increased levels of heat shock proteins (HSP), which protect and repair essential cellular proteins from

heat damage. NSAIDs upregulate HSPs and may confer protection against heat stroke in patients that are on NSAIDs at the time of the heat stroke incident. In fact, in humans, NSAIDs may be indicated for heat stroke prevention during heat waves although there is no published research on this in dogs. Importantly, NSAIDs are contraindicated after a heat stroke event has occurred due to the risk of negative effects on the kidneys, coagulation system, and gastrointestinal tract. The clinical signs of heat stroke depend on the degree and duration of temperature elevation, as well as a significant contribution by individual patient factors. Diagnosing heat stroke is often based on a history of confinement or exertion on a hot, humid day. Most animals will present with an elevated body temperature although it may be normal or low, especially if external cooling has been initiated prior to presentation or the patient is in the advanced stages of shock. On initial physical exam, tachycardia, panting, and injected mucous membranes may be noted. Cardiac arrhythmias may be ausculted or suspected due to the presence of pulse deficits. Neurologic deficits ranging from obtundation to coma and seizure may occur. Skin and mucous membranes should be examined for evidence of petechiae and ecchymosis and venipuncture sites should be monitored for appropriate hemostasis. Initial work‐up includes an electrolyte panel (including blood glucose), packed cell volume (PCV) with total solids (TS), complete blood count, chemistry profile, and coagulation assays. Common abnormalities include hemoconcentration with elevations in PCV and TS, elevations in liver enzymes (ALT, AST, alkaline phosphatase, bilirubin), renal and prerenal azotemia, elevated creatine kinase, prolonged clotting times, and hypoglycemia. Treatment The initial therapy for heat stroke should be aimed at rapidly returning body temperature to normal and correcting hypovolemia and acid–base and electrolyte derangements. Immediately upon recognition of a heat stroke crisis, owners should be instructed to thoroughly wet a patient with tepid water. Support staff should be trained to instruct owners in this crucial step provided it will not unnecessarily delay transport to a veterinary hospital. Upon presentation, the previously described active cooling techniques should be initiated. At least one peripheral IV catheter should be placed and room‐ temperature fluids administered to increase effective circulating volume and improve perfusion, enhancing peripheral heat loss. During fluid resuscitation, heart rate, blood pressure, lactate, PCV/TS, and urine production should be monitored to guide therapy. A description of goal‐directed resuscitation targets is found in Chapter 41. If hypoproteinemia

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is present (albumin 40 breaths/min)

Signs consistent with pneumonia, pleural effusion

Vascular forces

Normal systolic blood pressure (BP) and albumin >18 g/L

Systolic BP 180 mm Hg or albumin 500 μg/L

100

94.7

Peritoneal amylase >1050 U/L

71.4

84.2

Peritoneal total lipase >500 U/L

92.9

94.7

McCord et al. J Vet Intern Med 2012;26:888–96

n = 84 AP = 57 Non‐AP = 27

Combination of clinical and imaging results

Spec‐cPL >400 μg/L

71.7–77.8

80.5–88

Lipase

43.4–53.6

89.3–92.5

Amylase

52.4–56

76.7–80.6

Mansfield et al. J Vet Diag Invest 2012;24:312–18

n = 32 Minimal/no inflammation = 20

Histology

Spec‐cPL >400 μg/L

NA

90

Trivedi et al. J Vet Intern Med 2011;25:1241–7

n = 70 Mild AP = 56 Moderate‐severe AP = 7 Normal pancreas = 7

Histology

Spec‐cPL >400 μg/L

21 (mild) 71 (mod‐severe)

100

TLI

30 (mild) 29 (mod‐severe)

100

Lipase

54 (mild) 71 (mod‐severe)

43

Amylase

7 (mild) 14 (mod‐severe)

100

Mansfield et al. J Vet Diag Invest 2011;23:691–7

n = 61 AP = 41 CP = 3 Pancreatic carcinoma = 5 Nonpancreatitis = 12

Histology

CPE‐1 >17.24 ng/mL

61.4 (all) 66 (AP) 78 (severe AP)

91.7

Neilson‐Carley et al. Am J Vet Res 2011;72:302–7

n = 64 AP = 20 Other disease = 17 Healthy = 27

Histology

Spec‐cPL >400 μg/L

NA

95%

Steiner et al. Vet Ther 2008;9:263–73

n = 22

Gross evidence at postmortem

Spec‐cPL >400 μg/L

63.6

NA

Lipase

31.8

NA

Amylase

40.9

NA

Steiner et al. J Vet Intern Med 2001;15:274

n = 11 (Severe AP)

Clinical findings and histology

Spec‐cPL >400 μg/L

88

NA

Mansfield and Jones Aust Vet J 2000;78:416–22

n = 42 AP = 15 Non‐AP = 27

Clinical findings and histology

Lipase >3 × RI

63.6

54.6

Amylase >3 × RI

22.7

78.1

TLI >100 μ/L

37.5

89.3

Source: Adapted from Mansfield (2013). AP, acute pancreatitis; CP, chronic pancreatitis; CPE‐1, serum canine pancreatic‐elastase‐1; cPL, canine pancreatic lipase (or canine pancreatic lipase immunoreactivity in earlier studies); NA, not assessed; RI, reference interval; TLI, trypsin‐like immunoreactivity.

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(a)

(b)

Figure 55.3  VD radiograph of a dog with acute pancreatitis. The descending duodenum is pushed laterally (arrow) and there is decreased contrast in the cranial abdomen.

(a)

(b)

Figure 55.2  Abdominal radiographs of a dog with free fluid (lateral (a) and VD (b)). The cause of the effusion cannot be determined by radiographs alone. Fluid analysis and further imaging is required.

Cytology and Histology Cytology is not used frequently in evaluating dogs with AP. However, abdominal fluid analysis should form part of the work‐up to ensure that a septic or neoplastic process is not missed. Dogs with AP will generally have a suppurative, non-septic exudate present. A recent study in a small number of dogs with AP determined that measurement of cPL in peritoneal fluid had a sensitivity of 100% when using a cut‐off value >500 μg/L. Caution

Figure 55.4  (a) Ultrasound image of enlarged hypoechoic right pancreatic limb (arrow). (b) In this ultrasound image, the pancreas is larger and more heterogeneous in appearance, with hyperechoic speckling of surrounding fat.

55  Pancreatitis in the Dog

should be applied before relying extensively on this, as the study had only a few dogs with inflammatory/septic exudates as comparison. Aspiration of fluid‐filled collections within the pancreas may have a therapeutic benefit (described later). Fine needle aspirates (using a 25 G needle) can safely be obtained from the pancreas and may be beneficial in some cases of CP. Cytology is generally considered of marginal utility in AP, as there is often such a large degree  of necrosis and inflammation that the samples are  ­nondiagnostic and/or cellular criteria overlap with malignancy. Histologic evaluation of pancreatic biopsy is considered the gold standard for diagnosis of pancreatic pathology. This is seldom obtained in dogs with AP, due to the morbidity associated with this procedure. The most common finding in dogs with AP is widespread necrosis of pancreatic and peripancreatic tissue. In dogs with milder, more intermittent clinical signs, exploratory ­laparotomy is more commonly indicated. The stomach, intestines (distal and proximal small intestine), mesenteric lymph nodes, and liver should all be sampled in addition to the pancreas. A summary of the diagnostic approach recommended when dogs present with severe signs compatible with AP is shown in Figure 55.5.

­Therapy Acute Pancreatitis Treatment of AP is generally supportive and nonspecific. The major “triad” of treatment is IV fluids, analgesia, and nutritional support. Fluid therapy should be tailored to the individual dog, with correction of electrolyte losses and restoration of circulating blood volume and acid– base balance being of the utmost importance. A recent study in humans suggested that using alkalinizing fluids (Hartmann’s solution) produced a better response than acidifying solutions (saline) and so extrapolation would suggest that this (or alternatively, lactated Ringer’s solution) would be a good first‐line fluid choice in dogs. The volume of IV fluids required to correct blood volume may be higher than can be tolerated, and potentially lead to pulmonary edema or other signs of volume overload. In those circumstances, the use of colloid therapy may be of benefit. There is little to no evidence that using plasma will be beneficial in dogs with AP unless they have overt coagulation disorders. Analgesia is essential in dogs that have AP, even if little pain is observed. Abdominal pain can be categorized as mild, moderate or severe (see Table 55.1). The principle behind analgesia in these situations is to start with the

maximal combination/dose considered necessary for the level of pain and then titrate down (rather than starting low and titrating up if ineffective). Ideally, analgesic ­protocols in AP should avoid the use of drugs such as fentanyl that dramatically reduce gastrointestinal ­motility. However, if pain is not controlled by the recommended protocols, then using fentanyl as a constant rate infusion (CRI) should be considered. The addition of methylnaltrexone may mitigate the decreased GI motility in those circumstances. A partial mu‐agonist such as buprenorphine or full mu‐ agonist such as methadone should be considered as the baseline therapy (starting at maximum doses and intervals: 10–40  μg/kg q6–8h and 0.1–0.5  mg/kg q4–6h respectively, then tapering down). If pain is moderate then ketamine (5–20 μg/kg/min) and lidocaine (25–50 μg/kg/ min) CRIs should be started at the higher dosage end. Once pain is well controlled, the ketamine should be reduced first, followed by lidocaine and then tapering of the opioid. With severe pain, epidural morphine (0.1 mg/ kg q12–24h) or a fentanyl CRI (0.2–0.8  μg/kg/min) should be given, along with the lidocaine and ketamine as above. Once  pain is controlled, the epidural/CRI is swapped for intermittent opioid, then all are tapered as previously described. If possible, oral gabapentin (10 mg/kg q12–24h) could also be administered. If there is a sudden relapse of pain, the pancreas should be reexamined via ultrasound for the presence of a fluid collection. In people, aspiration of fluid collections is recommended when they are sterile, in preference to surgical debridement. Even in infected necrosis, control with antibiotics is undertaken before any surgical procedures. As virtually all the fluid collections in dogs will be sterile, and pancreatic surgery is associated with high morbidity and mortality rates, percutaneous drainage of fluid‐filled areas is advocated for dogs as well. This can be performed using mild sedation and fine needle aspiration. Nutritional support is the third major component of management. In people and dogs with critical illness, lack of enteral nutrition has been shown to perpetuate systemic inflammation as well as leading to mucosal atrophy and other changes. In mild‐to‐moderate cases of AP, the current rationale is to start enteral feeding if there has been no/little enteral nutrition for more than five days. In severe cases of AP, the earliest possible enteral nutrition is advocated. Dogs with severe AP can tolerate esophageal tube feeding, and so surgery or prolonged anesthesia to insert jejunostomy tubes is not essential. If the animal is moribund or unable to withstand a general anesthesia, then nasoesophageal tube feeding should be started as soon as possible. Once they are able to have GA safely, then an esophageal (E) tube should be inserted. Dogs can still be offered food by

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55  Pancreatitis in the Dog

mouth, and can be sent home with an E tube in place if necessary. The biggest complications associated with interventional enteral nutrition are aspiration pneumonia and tube site infection. Close attention should be paid to the tube site to avoid the latter. To avoid regurgitation and development of aspiration pneumonia, it is not recommended to feed full resting energy requirements (RER) unless the feeding is extremely well tolerated. Rather, the first 24 hours should be 25% RER and titrate up slowly each day. Similarly, there is no evidence that a specific diet will result in improved outcome or speed of recovery, and as such any balanced convalescent diet for dogs is generally well tolerated. Another important aspect of management of AP is to reduce vomiting and control nausea. Due to the role of substance P in mediating abdominal pain and vomiting, maropitant should be considered a first‐line antiemetic drug. If vomiting is poorly controlled, or the dog appears to have severe nausea, then additional medications such as ondansetron (0.5 mg/kg IV then q12–24h) can be added. There is a theoretical disadvantage in using metoclopramide as it inhibits dopamine in rodent models. Due to the potential prokinetic benefits of metoclopramide a CRI should still be considered if decreased GI motility is thought to be contributing to nausea/ inappetence. There is little to no evidence that administering gastric acid suppressants does anything to reduce morbidity of AP in dogs. However, if there is melena or hematemesis then use of a proton pump inhibitor should be considered. The presence of any systemic complications such as pleural effusion or ventricular tachycardia should be assessed and treated as required. Antibiotics are not necessary unless bacterial translocation is considered likely. The area most likely to undergo investigation in the future is targeted antiinflammatory therapies and analgesic agents. Similarly, the use of corticosteroids ­ may be further evaluated, particularly when hypotension appears to be nonresponsive to fluid therapy. Follow‐Up

If an inciting cause (such as drug or food) is known, then this should be removed. Dogs can be discharged with feeding tubes in place if required, and a low‐fat diet is currently recommended. This should generally be fed for 1–2 weeks and then the dog should be reevaluated and fasting serum triglyceride and cholesterol concentration measured. If these are high, investigations should be undertaken for underlying causes of hyperlipidemia and treatment, including diet, continued as appropriate. If there is no hyperlipidemia, dogs can be transitioned back to their regular diet. Analgesia may be necessary in the

early stages following hospitalization, and medication such as tramadol or gabapentin dispensed. Anecdotally, tramadol may cause inappetence and so gabapentin may be a better alternative. In people, after a bout of severe AP subclinical EPI exists for approximately four weeks following resolution. It is not certain if this occurs in dogs, but it may be therapeutically useful to supplement pancreatic enzymes for a short period after discharge. Animals should not be discharged if they still need antiemetic therapy. Chronic Pancreatitis Unfortunately, treatment options for CP are poorly explored, and the benefit of proposed treatments is unsupported. It does make sense to look for underlying causes of CP that can be corrected, such as hyperlipidemia. If there is hyperlipidemia, then an underlying endocrinopathy should be considered, particularly hypothyroidism. If both triglycerides and cholesterol are increased, then an inherent defect in lipid metabolism is more likely. In the absence of an endocrinopathy, treatment should initially consist of feeding a low‐fat diet, with 80% of cats with pancreatic

disease show this form of pancreatic infiltrate. Remarkably, at least one necropsy‐based study of cat pancreata has suggested that signs of pancreatic inflam­ mation, including lymphocytic‐plasmacytic infiltration and fibrotic change, are common in cats with no history suggestive of pancreatic disease. One problem with the terminology of pancreatitis, particularly the distinction of acute vs chronic pancreati­ tis, is the implication of differing chronologies of the dis­ ease. This is particularly problematic in the feline patient. While it is generally safe to assume that cats with longer histories of recurrent clinical signs such as lethargy, decreased appetite, and recurrent vomiting do indeed have chronic pancreatitis, the converse is not necessarily true. Given the subtle nature of clinical signs and histori­ cal complaints in many cats with pancreatic disease, and the waxing/waning nature of clinical signs in most chronic lympho‐plasmacytic diseases, it is not valid to assume that a cat with a new presentation of relatively acute clinical signs actually has “acute pancreatitis” as defined histologically. As discussed earlier, there is no reliable way to distinguish necrotizing or hemorrhagic pancreatitis from lympho‐plasmacytic disease in feline patients. Therapeutic decision making should be based on assessment of the overall state of health of the patient on presentation, rather than assumptions that a patient may or may not have a particular form of the disease.

­ oninvasive Diagnostics for Feline N Pancreatitis Routine Clinical Chemistries Routine clinical chemistry and complete blood count (CBC) evaluations are a critical part of the approach to any unwell feline patient. In the context of cats with a suspicion of pancreatitis, the most important aspect of the routine chemistry panel and CBC is the assessment of the overall physiology of the patient. There are very few to no findings on routine clinical pathology panels that can be considered highly sensitive or specific for pancreatitis in the cat. One exception to this would be the “pancreas‐specific lipase” assays that are beginning to be promoted more heavily in some routine chemistry panels, and are discussed in more detail later. In the dog and human patients, traditionally we have relied on the measurement of serum amylase and lipase activities to establish a diagnosis of pancreatitis, with the expectation that these activities will be markedly elevated. While these tests are admittedly low sensitivity and speci­ ficity in both dogs and people, marked elevations in patients with compatible clinical signs are at least strongly supportive of the clinical suspicion of pancreatitis.

56  Pancreatitis in the Cat

In the cat, a long‐standing belief in veterinary medicine is that amylase and lipase activities have essentially no diagnostic utility, with unacceptably low sensitivity and specificity in this species. While there is a large element of truth to these assertions, these opinions are based on a limited number of publications that assessed a small num­ ber of cats, and all dated from a period before the veteri­ nary profession came to truly appreciate the very different manifestations of this disease in the cat. The low specific­ ity reported for these tests may owe as much to false‐­ negative diagnoses based on an incorrect understanding of the way in which this disease presents in the cat. Common clinical pathology abnormalities encoun­ tered in cats are related to organ systems that are com­ monly compromised or involved in co‐morbid diseases. Abnormalities in liver enzyme activities (both alanine aminotransferase [ALT] and alkaline phosphatase [ALKP], suggesting both hepatocellular compromise and cholestasis), azotemia, alterations in acid–base status, hypocalcemia, and inflammatory leukograms (typically left‐shift neutrophilia) are all encountered relatively commonly. Some studies have suggested that marked hypocalcemia is a negative prognostic factor, likely due to sequestration of calcium in saponified abdominal fat, a phenomenon that tends to accompany hemorrhagic and necrotizing pancreatic pathology. Specialty Diagnostic Tests Noninvasive diagnostic testing relies on the use of marker compounds, detectable in the serum or some other sample such as urine, that are specific to the organ of interest and are altered in disease states of that organ. Ideally, the marker compound will be altered with dis­ ease of the primary organ, but unaltered by changes in other organ systems or by concurrent medical therapy. In the case of pancreatic disease, these marker com­ pounds are usually digestive enzymes of some type. The use of typical amylase and lipase activity assays has already been discussed. More recent diagnostic tests that have been investigated include serum concentrations of trypsin‐like immunoreactivity (fTLI), feline pancreatic lipase (now offered as the Spec‐fPL™ assay), and activity tests using more “pancreas‐specific” lipase substrates, particularly DGGR‐lipase activity. While trypsinogen/fTLI is pancreas specific, and is released in conditions of pancreatic disease, the peak changes in this enzyme are relatively short and soon after the onset of disease, and some other diseases (specifi­ cally small intestinal disease and severe azotemia) can be associated with elevations in fTLI without histologic evidence of pancreatic disease. This combination of ­ transient changes in serum concentrations and minor alterations with other, nonpancreatic diseases results in

fTLI testing having only moderate sensitivity and speci­ ficity for the diagnosis of pancreatitis in the cat. The greatest utility of fTLI testing lies in the ruling in or out of exocrine pancreatic insufficiency in feline patients (see Complications of Pancreatitis in Cats, later). Serum concentrations of feline pancreatic lipase (Spec‐ fPL) and lipase activity assays using more highly selective assay substrates both show greater utility than routine amylase/lipase activities and the fTLI test; these tests are discussed in greater detail later. Specific Pancreatic Lipase Test

Feline‐specific pancreatic lipase (fPLI) refers to a par­ ticular protein, a lipase enzyme, synthesized and released solely by the pancreas in cats. This particular enzyme’s concentration in serum is measured using the serum Spec‐fPL assay, available either as a “cage‐side” SNAP test or via specialty clinical pathology testing. This assay relies on monoclonal antibodies directed against the feline protein which are highly species specific. The use of canine PLI tests with feline samples can result in a false‐negative rule‐out of pancreatic disease in the cat. As the fPLI protein is specifically and exclusively ­synthesized in the exocrine pancreatic tissue, the con­ centrations in the serum are dependent on the rate of synthesis and the “leakage” of the protein into the inter­ stitial space and then the circulation. Conditions associ­ ated with increased cellular leakage (generally, some form of pancreatitis) are associated with increased serum concentrations of this protein, so the detection of increased circulating fPLI protein (i.e., a high Spec‐fPL test result) is suggestive of pancreatic disease, particu­ larly in cats with compatible clinical signs. The sensitivity and specificity of fPLI testing have been investigated in several studies. Reported sensitivities and specificities for this test are 67–100% and 67–82%, respectively. In one moderately large study (n = 182 cats) using the Spec‐fPL assay, the overall sensitivity for this test was 79%, with a specificity of 82%, for detection of pancreatitis in this group. Overall, the Spec‐fPL has the highest reported sensitivity and specificity (at the time of writing) of any diagnostic modality for the detection of pancreatitis in the cat. The use of Spec‐fPL measurement as a prognostic marker and monitoring parameter for recovery has also received some attention. Marked elevation of Spec‐fPL (values >20 μg/L) has been associated with a poorer prognosis in cats hospitalized with pancreatitis. There are no data in the literature suggesting that greater eleva­ tions in Spec‐fPL are associated with poorer response to therapy in cats with chronic pancreatitis, assuming ade­ quate management of co‐morbidities. A much greater factor influencing response to chronic pancreatitis in cats is adequate management of chronic enteropathy and

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the identification and treatment, if present, of cobalamin deficiency in these cats. Pancreas‐Specific Lipase Assays

The lipase activity assays used in most commercial labo­ ratories and bench‐top analytic systems rely on lipase within the serum sample cleaving a substrate of some sort; this cleavage then results in a change in the test solution (such as changing pH or color) that can be measured, giving a value for lipase activity. There is a large number of lipase activity substrates that can be used, and the substrates typically used by high‐through­ put commercial laboratories are different from those used in the bench‐top systems. Even within bench‐top systems, there are differences in substrates between dif­ fering manufacturers. For this reason, the comparison of lipase activity assays between two different systems is generally fruitless and does not yield any useful data. Many tissues in the body contain quite high amounts of lipase. Obviously, the exocrine pancreas is very rich in these enzymes but other organs, including the liver, gastric mucosa, and duodenal mucosa, contain lipase ­ activity. These lipases can be released from these other organs in disease states, and contribute to the total circu­ lating lipase activity. The specificity of a lipase assay for detection of pancreatic disease thus depends on the speci­ ficity of the substrate used for pancreatic lipase. This sub­ strate specificity varies markedly between s­ ubstrate types. One substrate with greater specificity for pancreatic lipase that has undergone some investigation in cats is 1‐2‐o‐Dilauryl‐rac‐glycero‐3‐glutaric acid‐(6’‐methyl­ resorufin) ester (DGGR). A DGGR‐based clinical chem­ istry assay has been validated for use in feline serum samples, and the sensitivity and specificity of the test compared with the use of both Spec‐fPL testing and abdominal ultrasonography. The DGGR‐based assay had similar, if slightly lower, sensitivity and specificity to the Spec‐fPL assay in the assessment of cats with clinical signs compatible with pancreatitis. In a similar follow‐up study, the DGGR‐based assay was found to show the best agreement with both Spec‐fPL and abdominal ultra­ sonography in cats with hypo‐ or mixed‐echoic pancre­ atic parenchyma, findings that are generally considered consistent with acute pancreatic disease. Overall, DGGR‐based lipase activity assays likely have clinical utility in feline patients, and an elevated activity value in an unwell feline patient should prompt further assessment for the presence of pancreatic disease. However, this approach is reliant on the use of this spe­ cific substrate in whichever analytic system is being employed. Information regarding the actual substrates used by the various reference laboratories and in‐house chemistry systems commonly found in veterinary prac­ tice is not readily available at this time.

Pancreatic Biopsy and Histology Pancreatic biopsy and histologic examination is gener­ ally considered the “gold standard” for the detection of pancreatic disease, including pancreatitis. Unfortunately, pancreatic biopsy is still a relatively uncommon part of the approach to feline patients, for a variety of reasons. Chief among these is likely cost to the client and the invasive nature of celiotomy to obtain biopsies. Recently, however, the use of laparotomy and laparoscopy‐assisted biopsy techniques has provided a lower cost, less inva­ sive method for obtaining pancreatic biopsies, usually in concert with liver and intestinal biopsy procedures. Surgical biopsy of the feline pancreas has been shown to have a low risk for complications such as postsurgical pancreatitis, assuming that the patient is adequately hydrated and hemodynamically stable at the time of sur­ gery. Side‐effects and postsurgical morbidity are uncom­ mon in cats undergoing surgical biopsy of the pancreas using punch biopsy instruments or crush‐fracture tech­ niques using hemostats. Pancreatic biopsy, particularly in cats undergoing assessment for gastrointestinal dis­ ease, is justified in many cases and should be considered part of the routine samples collected during exploratory celiotomy or laparoscopic exploratory procedures. While biopsy samples are rightly considered the most effective way of assessing pancreatic disease, a number of shortfalls of this technique must be recognized. Even with laparoscopic procedures, the collection and assess­ ment of histologic samples are still more expensive and have higher risk to the patient than the use of noninva­ sive testing such as clinical chemistries. In addition, pathology within the pancreas is often heterogeneous in distribution, and differing areas may show differing cel­ lular infiltrates or levels of activity. This may lead to incorrect diagnosis of the overall state of the pancreas when only a single sample is taken, as is often the case with laparoscopic biopsies of the pancreas in the cat. Finally, there is a well‐recognized variability between individual pathologists with respect to assessment and description of pancreatic samples. Histologic scoring systems for pancreatic disease have been proposed and published, including for the cat, but the use of these sys­ tems is generally limited to research publications and retrospective case series descriptions. It is uncommon for histopathologists to report pancreatic biopsy samples using these scoring systems in veterinary medicine. Interpretation of Specialized Tests Interpretation of the specialized tests used for the diag­ nosis of pancreatic disease in cats is sometimes compli­ cated, particularly if the test is part of a broad‐ranging panel used in the initial assessment of a patient if there is  no clear initial diagnosis being assessed. Part of the

56  Pancreatitis in the Cat

­ ifficulty of assessing these tests comes from the obser­ d vation that we do not truly know the prevalence of pan­ creatic disease in the feline population. While at least one study suggests that pancreatic pathology is quite common in the feline population (approximately 60% of cases necropsied, where the patient was not suspected to have pancreatic disease), one caveat about this study is that these were still, largely speaking, unwell cats that were necropsied. If the true prevalence of pancreatic dis­ ease in the feline population is as high as 60%, then an unexpected finding of a high Spec‐fPL concentration or DGGR‐lipase activity on sentinel chemistry panels may be a true indication of pancreatic disease. While the pan­ creatic disease present may not be an important clinical finding, this does not mean that this is an incorrect diagnostic finding. This is particularly true when the clinical chemistry panel is being run on an apparently healthy patient as part of a health monitoring or wellness program. An additional important factor in the interpretation of these tests is the very variable, and often subtle, clinical signs and histories of this disease, as previously dis­ cussed. When clinical chemistry panels and Spec‐fPL tests are being run as part of a broad‐ranging assessment in a cat that is known to be unwell, but with no clear ini­ tial indication of why, abnormalities in these tests are a strong indication of likely clinically significant disease. It is inappropriate to discount these findings in these patients, and important to consider that pancreatic ­disease is, to all indications, common in the sick cat pop­ ulation and also a common co‐morbid condition with other diseases. Diagnostic Imaging Along with the specialized noninvasive testing methods discussed above, diagnostic imaging is commonly used in the assessment of pancreatic disease in the cat. Of the various methodologies available, high‐resolution abdominal ultrasonography typically has the greatest diagnostic yield. It also allows further assessment of other abdominal organs that may be involved in co‐mor­ bid diseases, such as the hepatic parenchyma, biliary tree, and intestinal walls. Plain abdominal radiography, while commonly used in cats with suspected abdominal disease, has a very low diagnostic yield in cats with pancreatitis. The pan­ creas is not visible on plain abdominal radiographs, and changes commonly seen in dogs with pancreatitis (“ground glass” appearance, loss of local contrast, wid­ ening pyloric angle) are infrequent in cats due to the differing underlying pathology in most cases. While abdominal radiography has some possible advantages in terms of lower cost and greater availability, the very

low diagnostic yield is a contraindication for its use in most cases. The increasing availability of high‐resolution abdomi­ nal ultrasonography in companion animal practice has dramatically improved the use of diagnostic imaging to characterize pancreatic disease in the cat. Observation of pancreatic enlargement, hyperechoic mesentery and abdominal fat, altered parenchymal echogenicity (most commonly hypoechogenicity or a mixed pattern), abdominal effusions, pancreatic cysts or pseudocysts, corrugation of the duodenum, and dilation of the pan­ creatic duct are all considered consistent with pancreati­ tis in cats with consistent clinical signs. These findings are specific indicators of the presence of pancreatic pathology, but their sensitivity for accurate diagnosis in cats with histologically confirmed pancreatitis is quite variable, with reported sensitivities ranging from ~11% to 80%. Sensitivity of abdominal ultrasound for the detection of feline pancreatic disease is highly dependent on operator skill and equipment resolution. Given the variable and often low sensitivity of abdominal ultra­ sound examination in the diagnosis of pancreatitis in cats, this modality cannot be reliably used as a “rule‐out” test for this disease. However, significant additional information useful for management of these cases is often obtained, particularly when screening for co‐ morbid conditions such as cholangitis/cholangiohepati­ tis or the presence of extrapancreatic disease that could  explain the clinical signs. Therefore, abdominal ultrasonography, carried out by a well‐trained and experi­ enced ­operator, is recommended in all cats where there is a clinical suspicion of pancreatitis. In humans, the use of abdominal computed tomogra­ phy (CT) examination is considered highly useful in the assessment of pancreatic disease. This modality allows the assessment of pancreatic architecture, presence and severity of fibrotic change and detection of both com­ mon inciting causes for pancreatitis in humans (such as gallstones) and other co‐morbid diseases. The utility of abdominal CT for the assessment of feline pancreatic disease is much lower than in humans. This is due to the much smaller mass of pancreatic tissue in the cat, and less common presence of severe architectural dis­ tortions due to fibrosis. Additionally, abdominal CT examination of cats typically requires general anesthe­ sia, which may be contraindicated in severely unwell cats and also leads to significant additional cost. CT examination has been reported in cats in a small num­ ber of publications; the overall sensitivity and specific­ ity were not higher than those seen with less invasive testing such as Spec‐fPL or abdominal ultrasonography, and thus abdominal CT cannot be recommended as a part of the diagnostic approach to pancreatic disease in the cat at this time.

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­Approaches to Management of Pancreatitis in Cats Care of Cats Hospitalized for Pancreatitis As previously discussed, it is not possible to distinguish so‐called “acute necrotizing” pancreatitis from chronic lymphocytic‐plasmacytic pancreatitis in the cat, regard­ less of presenting clinical signs, history or chemical find­ ings. The approach to the management of cats with this disease is determined by the overall degree of health of the patient as assessed by initial examination and screen­ ing chemistries. Cats presenting with a suspicion of pan­ creatitis that are showing other signs of systemic illness, marked abdominal pain, systemic inflammatory response syndrome (pyrexia, tachycardia, tachypnea), hypother­ mia or hypotension should be considered to have severe disease, and immediate hospitalization and aggressive stabilization therapy are recommended. If multiple abnormalities are detected on routine chemistry panels, particularly if hypoalbuminemia and/or hypocalcemia are present, this is a strong indication of severe disease associated with significant risk of in‐hospital mortality. Cats with severe disease require careful monitoring, appropriate fluid therapy, and effective analgesia. In humans and dogs, severe pancreatic disease is inarguably painful, and while cats may not show as many overt signs of pain, such as vocalization or abdominal guarding, there is no reason to suspect that this disease is not pain­ ful in cats as well. Many of the clinical signs at presenta­ tion of severe cases, such as tachypnea and tachycardia, resolve rapidly with effective pain control interventions. Initial screening chemistry panels are important to assess for other co‐morbid or complicating diseases, par­ ticularly extrahepatic biliary duct obstruction, hepatic lipidosis, and diabetes mellitus (which may proceed to diabetic ketoacidosis). Samples should also be drawn for special diagnostic testing (i.e. Spec‐fPL assay or DGGR‐ lipase). If available, patient‐side SNAP tests for Spec‐fPL can give initial support to a clinical suspicion of pancrea­ titis, but these tests should not be relied on to rule out the disease, and measurement of a quantitated value through a reference laboratory is recommended as soon as possible. Pending results of specialized diagnostics, the management of these cases follows typical manage­ ment protocols for acute abdomen/septic shock patients. The initial aims of therapy are to replace circulating fluid volumes, restore end‐organ perfusion, and main­ tain organ function. Obtaining effective perfusion of the pancreas is particularly important, as pancreatic ischemia is a significant inciting cause for the develop­ ment of necrotizing pancreatitis. With the combination of aggressive fluid therapy and increased endothelial “leakiness,” it is critical to monitor total protein and, if

necessary, restore and maintain plasma colloid oncotic pressure. Colloid fluids, such as synthetic hydroxyethyl starches, are often highly beneficial in the initial resusci­ tation of these cases. Fresh‐frozen feline plasma can also be considered, and likely provides oncotic support while replenishing coagulation cascade proteins; however, there is little information in the veterinary literature regarding use of plasma in severe feline pancreatitis cases. Substantial electrolyte abnormalities, particularly hypokalemia and hypocalcemia, should be anticipated in these cats. Supplemental potassium is administered in combination with crystalloid fluids following routine guidelines for concentrations based on serial determina­ tion of serum potassium concentrations, typically every eight hours during the initial resuscitation phase. Drug Therapy in Feline Pancreatitis Effective analgesia and control of vomiting are important aspects of management of severe pancreatitis in all spe­ cies. A selection of medications that are often useful in the management of pancreatitis in the cat is summarized in Table 56.2. Narcotic pain control is typically indicated in cats with sufficiently severe pancreatitis to warrant hospitaliza­ tion. Transdermal fentanyl patches (25 μg/h) can be very effective for longer term (up to 72 h) analgesia without the need for frequent handling and injection in these patients, but initial therapy with an injectable or sublin­ gual agent (commonly buprenorphine) is necessary as it can take up to 12 hours for therapeutic fentanyl concen­ trations to be reached. Maropitant, a neurokinin‐1 receptor antagonist, is both an effective antiemetic and may have antinociceptive effects in the viscera. The com­ bination of maropitant with a 5‐HT3 receptor antagonist, such as ondansetron or dolasetron, often provides effec­ tive control of vomiting and nausea in these patients with minimal need for repeated handling during the day. Together, obtaining adequate analgesia and control of nausea/vomiting are important for the overall well‐being of the patient, and increase the likelihood of an early return to voluntary eating. Early Nutritional Management Long‐standing dogma for the management of severe pancreatitis in humans and canine patients has been that these patients should be maintained nil per os for some period, with introduction of food occurring only after cessation of vomiting for a minimum of 12 hours, and often 24 hours or more. This attitude is being sup­ planted by early enteral nutrition strategies in both humans and, more recently, dogs. Preliminary evidence suggests that early enteral nutrition in these species is

56  Pancreatitis in the Cat

Table 56.2  Medications commonly used in the management of severe pancreatitis in the feline patient Class

Drug

Analgesic

Antiemetic Antinausea

Antacid

Mechanism

Main indication

Dose and route

Buprenorphine

Opioid

Acute, severe

0.01–0.02 mg/kg SC, IM, IV, TM

Fentanyl

Opioid

Acute, severe

25 μg/h patch, 5 μg/kg IV bolus, CRI 2–4 μg/kg/h

Butorphanol

Opioid

Acute, severe

0.1–0.5 mg/kg SC, IM, IV

Maropitant

Neurokinin‐1 receptor antagonist

Acute, severe chronic

1 mg/kg SC q24h, 2 mg/kg PO q24h

Dolasetron or ondansetron

5‐HT3 receptor antagonist

Acute, severe

0.8–1.0 mg/kg IV q24h

Metoclopramide

Dopamine D2 receptor antagonist

Acute, severe

0.2–0.5 mg/kg PO, SC, IM q6–8h, 1–2 mg/kg/24h CRI

Omeprazole

Proton pump inhibitor

Acute, severe

1.0–1.3 mg/kg PO q12h

Pantoprazole

Proton pump inhibitor

Acute, severe

0.7–1.0 mg/kg IV q12h

Ranitidine

Histamine H2 receptor antagonist

Acute, severe

0.5 mg/kg PO q12h

CRI, constant rate infusion; IM, intramuscular; IV, intravenous; PO, by mouth (per os); SC, subcutaneous.

associated with shorter ICU and hospital stays, lower total costs of care, and fewer complications during ini­ tial management. With cats, the potential for develop­ ment of hepatic lipidosis as a complication of severe pancreatitis and inadequate caloric intake must also be considered. While currently there is limited informa­ tion in the veterinary literature regarding the use of early enteral nutrition in cats with severe pancreatitis, the author uses this modality regularly, and quite aggressively. In many cats, assisted feeding devices such as esophagostomy tubes are placed during the first 24–36 hours of hospitalization to achieve early return to feeding. For both hepatic lipidosis and diabetic ketoacidosis, a return to caloric intake is critical to their successful management, and the presence of pancreati­ tis (either as an inciting disease or a complication) does not alter the importance of a return to positive caloric balance in these diseases. Outpatient Management of Cats with  Chronic Pancreatitis Cats presenting with less severe clinical signs, perhaps with histories of vomiting, weight loss or other gastroin­ testinal signs but without significant abnormalities on screening biochemical panels, can typically be treated on an outpatient basis. In these patients, collection of sam­ ples for specialized testing (Spec‐fPL) is recommended, but there is less utility to the patient‐side testing, as these patients will be treated on an outpatient basis in the same manner regardless of the initial patient‐side test result. Cats that had originally presented with a suspicion of severe pancreatitis should also transition to this more

chronic management approach once they are eating vol­ untarily and are able to be discharged. Cats with a clinical suspicion of chronic pancreatitis are treated in essentially the same manner as cats with chronic enteropathies or diagnoses of idiopathic inflam­ matory bowel disease. There is no meaningful way to distinguish between chronic pancreatitis as a solitary disease entity and the presence of multiorgan inflamma­ tory disease (so‐called feline inflammatory disease or “triaditis”). Initially, dietary modification, typically by use of a novel protein source or hypoallergenic diet, is suggested. In contrast to dogs, where fat restriction is a cornerstone of management for most cases of chronic pancreatitis, fat restriction is not recommended in the cat due to their high constitutive requirement for both fat and arachidonic acid intake. Many cats will respond to dietary manipulation; in those who fail to respond to dietary manipulation, it is rational to consider antiin­ flammatory or immune modulatory therapies, assuming that no other co‐morbidities are present that would con­ traindicate the use of these medications. A common additional finding in cats with chronic pancreatitis, particularly in those with significant gastro­ intestinal disease accompanying their pancreatic disease, is the presence of hypocobalaminemia. Cats with hypoc­ obalaminemia (low vitamin B12) are known to be less responsive to treatment for other diseases, such as chronic inflammatory bowel disease, if this hypocobala­ minemia is not addressed via supplementation. Supplementation of cats with cobalamin is straight­ forward, and can be carried out using injectable or oral supplementation regimes. In some cats, particularly those with diffuse small intestinal disease, serum folate

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c­oncentrations are also low. Supplementation of cats with low folate has not been investigated as thoroughly as cobalamin supplementation, but is also readily achieved with oral supplement products. Often, initial supplementation by injection followed by oral supple­ mentation in the home environment provides a good balance between costs, convenience, and ease of owner compliance.

­Complications of Pancreatitis in Cats Given the apparent frequency of chronic pancreatic dis­ ease in the feline population, the occurrence of severe complications is actually remarkably low. While objec­ tively severe disease in cats presenting with hemody­ namic shock, systemic inflammatory response syndrome and abdominal pain is life‐threatening and associated with a substantial risk for mortality if not addressed appropriately, most cats with chronic pancreatitis require life‐long management for their disease, but other than waxing and waning clinical signs, most do not develop additional complicating diagnoses. The two most common and important complications of pancreatic disease in the cat are exocrine pancreatic insufficiency (EPI) and diabetes mellitus. Chronic pancreatitis is most likely the most common cause of EPI in cats. Loss of exocrine tissue due to ongo­ ing inflammation and fibrosis can eventually lead to the development of clinical signs related to insufficient digestive enzyme synthesis. In comparison to dogs, and a recurring theme in feline medicine, the way in which EPI manifests in cats is different in terms of both history and clinical signs reported. While in the dog the most com­ mon cause of EPI is an early‐onset pancreatic acinar atrophy (PAA), in the cat the disease is an end‐result of a chronic, typically years‐long inflammatory disease. Dogs with EPI secondary to PAA are usually young and usually are otherwise healthy, with good to ravenous appetites. In the cat, there is typically a long history of poor appe­ tite and possible weight loss resulting from the underly­ ing pancreatitis. By the time cats reach an end‐stage of

chronic pancreatitis, they are usually chronically unwell, and they do not develop compensatory polyphagia. Cats also are less likely to produce voluminous stools and frank steatorrhea; this is probably due to the very pro­ longed course of the disease in this species, which allows other compensatory mechanisms (such as intestinal brush border enzyme upregulation) to be more effective. Exocrine pancreatic insufficiency should be suspected in any cat with a history of diagnosed chronic pancreatitis that then goes on to show worsening clinical signs, particu­ larly weight loss. In some cases weight loss will be the only additional clinical sign that suggests the onset of EPI. Diagnosis of EPI in the cat is based on the measure­ ment of serum concentrations of TLI, not Spec‐fPL. While Spec‐fPL values are almost invariably very low in cats with EPI, this is also seen in some normal cats, resulting in an unacceptably low specificity of the Spec‐ fPL assay for the diagnosis of EPI in cats. DGGR‐lipase activity assays have no diagnostic utility for EPI in cats, as most cats with EPI show DGGR‐lipase activities that are indistinguishable from normal cat values. Diabetes mellitus can develop in cats with chronic pan­ creatitis due to progressive loss of islet tissue with inflam­ mation and fibrotic change. These cats may spend extended periods as “prediabetic” patients, with reduced glucose tol­ erance. The actual frequency of chronic pancreatitis as a cause of diabetes mellitus in the cat is unknown, and likely underestimated, due to the low frequency of pancreatic biopsy in these cats. The occurrence of diabetes mellitus in feline patients can significantly complicate their manage­ ment for chronic pancreatitis, particularly if the cat is being managed with glucocorticoid medications to mitigate inflammation. Equally, the presence of chronic pancreatitis in a diabetic cat is a common cause of difficulty in achieving adequate glycemic regulation, and chronic pancreatitis is one of the most important diseases to screen for in cats with diabetes that is hard to regulate. The difficulty of managing diabetic cats with chronic pancreatitis, and the confound­ ing influence of glucocorticoid therapy on glycemic regula­ tion, makes one of the strongest arguments for early intervention using predominantly dietary changes in the management of chronic pancreatitis in the cat.

­Further Reading Bazelle J, Watson P. Pancreatitis in cats: is it acute, is it chronic, is it significant? J Feline Med Surg 2014; 16(5): 395–406. Caney SMA. Pancreatitis and diabetes in cats. Vet Clin North Am Small Anim Pract 2013; 43(2): 303–17. Oppliger S, Hartnack S, Reusch CE, Kook PH. Agreement of serum feline pancreas‐specific lipase and colorimetric lipase assays with pancreatic ultrasonographic findings

in cats with suspicion of pancreatitis: 161 cases (2008– 2012). J Am Vet Med Assoc 2014; 244(9): 1060–5. Pratschke KM, Ryan J, McAlinden A, McLauchlan G. Pancreatic surgical biopsy in 24 dogs and 19 cats: postoperative complications and clinical relevance of histological findings. J Small Anim Pract 2015; 56: 60–6 Simpson KW. Pancreatitis and triaditis in cats: causes and treatment. J Small Anim Pract 2015; 56(1): 40–9.

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57 Rectoanal Diseases – Medical and Surgical Management Craig Ruaux, BVSc (Hons), PhD, MACVSc, DACVIM (SAIM)1 and Milan Milovancev, DVM, DACVS‐SA2 1

 School of Veterinary Science, Massey University, Palmerston North, New Zealand  School of Veterinary Medicine, Oregon State University, Corvallis, Oregon, USA

2

Rectoanal diseases are frequent reasons for presentation of companion animals for veterinary attention. Clinical signs and symptoms, such as straining, “scooting,” fecal incontinence, and paradoxical diarrhea, are very distressing to owners. An effective approach to recognition and management of these diseases relies on a thorough history and clinical examination, including rectal examination wherever possible. In smaller patients, and those that resent digital rectal palpation, effective sedation is needed to allow this examination to proceed. Diseases involving the distal rectum and perineum are often well characterized by local examination and palpation, but the intrapelvic rectum can be difficult to fully assess, as many important structures (pelvic urethra, cranial ­prostate in larger dogs, intrapelvic lymph nodes) are difficult to reach. Diagnostic imaging of this area is also challenging in many cases. Contrast‐enhanced CT examination is the imaging method of choice for many of these cases. In this section, a variety of rectoanal diseases is briefly discussed. For some diseases, medical management is appropriate, while in others surgical management is the primary management modality.

­Anal Sac Adenocarcinoma Clinical Presentation Incidental finding or visible perianal mass/swelling, tenesmus, polyuria/polydipsia, anorexia or inappetence, lethargy, weight loss, constipation, or posterior weakness. Pathophysiology Anal sac adenocarcinoma arises from malignant transformation of the secretory epithelium of the anal sacs.

Diagnosis and Medical Management In up to a third of dogs, anal sac adenocarcinoma is ­discovered as an incidental finding on routine examination. When clinical signs are present, they may be a result of the local mass effect (straining to defecate, constipation due to enlargement of retroperitoneal and intrapelvic lymph nodes with metastatic disease). In some cases the primary presenting complaint is polyuria and polydipsia resulting from loss of concentrating capacity due to hypercalcemia. Routine clinical chemistry panels will often reveal both total and ionized hypercalcemia, with or without concurrent hypophosphatemia. A finding of hypercalcemia in a dog undergoing assessment for polyuria and polydipsia should prompt thorough anal sac and rectal digital examination, using sedation if necessary. With advanced disease and metastasis to the abdominal lymph nodes dorsal displacement of the colon may be visible on plain abdominal radiographs. Abdominal ultrasonography and, if available, computed tomography will usually have a much greater diagnostic yield than plain abdominal radiography. On diagnosis, complete pretreatment staging calls for either three‐view thoracic radiographs and/or computed tomography. Conclusive diagnosis can usually be obtained by examination of fine needle aspiration cytology of the mass. In some cases, the cytology may be interpreted as “hepatoid” but a benign cytologic appearance does not rule out anal sac adenocarcinoma. Surgical Management Preoperative patient staging for sublumbar lymph node and/or pulmonary metastasis is mandatory as this information influences surgical approach (e.g., determines whether concurrent sublumbar lymph node extirpation is indicated). Surgical removal of the affected anal sac is performed via closed anal sacculectomy. The size and

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adherence of the mass to the surrounding tissues ­influence the technical difficulty of the surgery. By the time of diagnosis, most tumors have caused enough distortion and adhesion to the perineal tissues (i.e., external anal sphincter, rectal wall, caudal rectal nerve) to be ­significantly more challenging than anal sacculectomy for ­nonneoplastic causes. Removal of involved external anal sphincter muscle is tolerated with loss of up to two‐ thirds of the circumference of the anus, as well as the ipsilateral caudal rectal nerve, with most dogs regaining fecal continence after a variable period of transient incontinence. Some dogs may have bilateral anal sac adenocarcinoma, which requires meticulous surgical ­ technique to preserve fecal continence. The most common complication after surgical excision of anal sac adenocarcinoma is tumor recurrence, with rates up to 42–45% reported. Other less common complications include surgical site infection, hypocalcemia, tenesmus, perianal fistula, and death. Prognosis Postoperative adjuvant therapy in the form of chemotherapy and/or external beam radiation therapy is quickly becoming the standard of care as several reports have documented prolonged survival in patients receiving multimodal treatment. With surgery alone, reported median survival times range from 7.9 to 16.4 months. In dogs receiving postoperative adjuvant melphalan, median survival time was 20 and 29.3 months in dogs with and without sublumbar lymph node metastasis, respectively. Another study combing postoperative radiotherapy with mitoxantrone reported medial overall survival of 956 days with one‐ and two‐year survival rates of 87% and 66%, respectively. A large (over 800 dogs) multicenter retrospective study in preparation at time of writing found prolonged median survival time (approximately five years) for dogs with sublumbar lymph node metastasis that were treated with surgery and chemotherapy, which was similar to the overall ­survival time for dogs without metastasis treated with surgery alone (J Liptak, personal communication).

­Anal Sac Impaction Clinical Presentation Anal sac impaction is one of the most common rectoanal diseases in companion animal practice, and a very ­common cause for presentation of dogs in particular. The clinical signs are of tenesmus, “scooting,” excessive licking of the perineum region, and pain on palpation of the

anal sacs (which are distended). Overall, smaller breed dogs appear to be predisposed to anal sac impaction. Pathophysiology Impaction of the anal sacs can occur secondary to a variety of primary causes or inciting disorders. Inadequate emptying of the anal sacs may occur in obese dogs, dogs consuming “low‐residue” diets with reduced fecal bulk, and in dogs with seborrhea (which results in increased secretion of the sebaceous glands within the anal sacs). Complications of anal sac impaction can occur, such as anal gland abscessation. Diagnosis and Medical Management In most cases, diagnosis and management of anal sac impaction is fairly straightforward. Diagnosis is based on palpation of the enlarged anal sac on routine physical examination. In many cases, diagnosis and management are concurrent, as the impacted sacs may be able to be gently manually expressed. Heavily impacted and painful glands may require sedation of the patient and irrigation of the sacs with saline. Uncomplicated cases are then  managed by addressing diet, providing regular opportunities for the dog to defecate, and client counseling. Sacs that have become abscessed should be cultured and appropriate antibiotic treatment administered. Uninfected impacted anal sacs do not warrant antibiotic therapy. Surgical Management The decision to proceed with surgical intervention for anal sac impaction or abscessation is based on failure to respond to appropriate medical management, including lancing, flushing, culture and sensitivity testing, use of appropriate antimicrobials, and prevention of self‐ trauma (i.e., use of an Elizabethan collar). Surgical intervention generally consists of anal sacculectomy and may be performed unilaterally or bilaterally, depending on the patient’s condition. When planning bilateral anal sacculectomy, it may be prudent to separate the two ­procedures by 2–4 weeks to allow any transient neuropraxia‐related fecal incontinence to resolve before performing surgery on the contralateral side. Anal sacculectomy for nonneoplastic disease can be performed using either an open or closed technique. With either method, the following principles apply: remove the entire gland such that no secretory epithelium remains in the patient; minimize trauma to the external anal sphincter muscle and avoid damage to the  caudal rectal nerve (branch of the pudendal n.) to

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prevent fecal incontinence; and ligate the anal sac duct near its opening, removing it en bloc with the gland itself. These goals are most readily achieved with the patient positioned in sternal recumbency in a perineal stand with the tail in an elevated position and an anal purse‐ string suture placed cranial to the opening of the anal sac ducts. If a closed technique is chosen, the anal sac itself can be distended with various materials (e.g., yarn, silicone sealant, Foley balloon catheter, plaster of Paris, agar base gel, dental molds, etc.) to facilitate palpation of the  gland itself during dissection. Closure is typically performed in two layers; 3‐0 or 4‐0 monofilament absorbable suture for the subcutaneous tissues and either similarly sized nylon skin sutures or an intradermal skin closure can be used.

imperforate anus), type III (as with type II but the rectum ends further cranially), and type IV (terminal rectum and anus may be normal but there is a discontinuity in the rectum with a blind pouch within the pelvic canal). These animals appear normal until 2–4 weeks of age at which time they become unthrify, anorexic, and restless with abdominal distension, possibly with perineal bulging. Rectoanal strictures appear to be more common in cats than dogs, but can occur in either species, most typically after trauma (iatrogenic or spontaneous). Clinical signs may vary depending on the underlying cause, but include tenesmus, dyschezia, hematochezia, and passing of ribbon‐like feces. Megacolon is a possible sequel, especially in cats with chronic partial rectal narrowing (e.g., due to malunion after pelvic fractures).

Prognosis The prognosis for patients with anal sac impaction is generally excellent, regardless of whether they require surgical intervention or respond to medical management.

­Atresia Ani and Rectoanal Strictures Clinical Presentation Atresia ani is the most common congential rectal/anal anomaly in dogs (Figure 57.1). Four types are recognized: type I (stenosis), type II (persistent anal membrane with  the rectum ending immediately cranial to the

Pathophysiology The functional diameter of the distal colon, rectum or anus is reduced, due to either congenital (atresia ani) or acquired anatomic abnormalities. Diagnosis and Medical Management Atresia ani may be diagnosed at the time of birth, or in the neonatal period, with failure of the infant to defecate and possibly the development of abdominal distension. Later, adult‐onset disorders are usually presented for constipation/obstipation, and can have a history of prior trauma or injury to the pelvis or perianal region. Surgical Management

Figure 57.1  Type II atresia ani in an 8‐week‐old mixed‐breed puppy. Note the imperforate anus and absence of rectal lumen. This particular puppy presented later than is typical for this disorder due to the concurrent presence of a rectovaginal fistula that prevented colonic and rectal obstruction. Note also the abnormalities in vulval anatomy.

Atresia ani type I is treated with gentle bougienage. Type II and III atresia ani are treated surgically via a vertical incision at the anal dimple with caudal advancement of the rectum. The rectum is sutured to the skin at the level of the anus in two layers using 4‐0 or 5‐0 monofilament absorbable simple interrupted sutures. Dissection must be particularly gentle as the tissues are friable and every effort must be made to preserve the anal sphincter as well as opening to the anal sacs, both of which are typically normal and present. Animals with type IV ­ ­atresia ani may require an abdominal approach with or without a pelvic osteotomy to access and anastomose the rectum. Treatment chosen for anorectal strictures is dictated by the severity and location of the stricture. Mild cases may respond to bougienage, but more severe conditions may require surgical resection and anastomosis. However, recurrence of stricture after surgical resection and ­anastomosis, especially in cats, is relatively common.

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Prognosis Surgical repair of type I or II atresia ani often results in long‐term survival and fecal continence. Patients with type III or IV atresia ani have poorer outcomes and the prognosis depends on the possibility of reconstructing the terminal rectum. Prognosis following treatment for anorectal stricture largely depends on the underlying cause. Benign causes may carry a good prognosis but strictures associated with rectal adenocarcinoma are associated with a more guarded to poor prognosis. Surgical complications following rectal stricture resection and anastomosis ­ may include dehiscence with sepsis and/or recurrence of stricture.

­ onstipation, Obstipation, and C Megacolon Clinical Presentation Constipation and obstipation are both manifestations of difficult defecation, and fall within a spectrum of severities. Constipation is defined as difficult or effortful defecation, but feces is passed after some effort. Obstipation is a state of more severe constipation, and may result in intestinal obstruction. Both constipation and obstipation are the result of disorders in large bowel motility that slow fecal transit, but an implication of both diagnoses is that there is some degree of large bowel and rectal motility remaining. Obstipation should be treated aggressively, and the early use of promotility agents is recommended in many cases, as failure to restore functional motility in these patients can result in progression to megacolon, a state in which all large bowel and normal rectal motility is lost. Once established, megacolon is not  effectively treated with laxative or prokinetic medications, as both of these families of medications require some degree of intrinsic motility in order to be effective. Pathophysiology Constipation and subsequent obstipation can result from any disorder or abnormality that slows fecal transit through the large intestine, decreases large intestinal and rectal motility, or mechanically obstructs passage of the fecal material to the rectum and anus. Slower passage of fecal material results in greater dehydration of the large intestinal content, as water absorption by the large intestinal mucosa continues unabated. Progressive overloading of the large intestine will eventually lead to large intestinal atony and flaccid paralysis,

as seen with ­chronically obstructed urinary bladders. Similarly to the urinary bladder, loss of large bowel intrinsic motility is almost universally irreversible, resulting in megacolon. Diagnosis and Medical Management A history of unproductive straining or tenusmus, in concert with a readily palpable, heavily feces‐laden large intestinal loop in the mid to distal abdomen, establishes a diagnosis of constipation readily. Differentiation of constipation/obstipation from megacolon can be more challenging, however. On plain abdominal radiographs, measurement of the maximal colon diameter:L5 vertebral body length ratio has been suggested as a method to help differentiate constipation from megacolon. A colon diameter:L5 length ratio of >1.48 has a sensitivity of approximately 77% with a specificity of 85% for establishing a diagnosis of megacolon. Similarly, a ratio of 12 has been shown to be associated with a much worse clinical prognosis. In these cases, an abbreviated clinical work‐up with endoscopy being scheduled earlier, as well as early aggressive treatment, may be indicated. The diagnostic assessment of chronic gastrointestinal inflammation involves exclusion of other potential causes of the gastrointestinal signs, and thus a full diagnostic work‐up needs to be done to rule out all known causes of extragastrointestinal inflammation first. Commonly, this involves complete blood cell count, serum biochemical analysis, urinalysis, and fecal analysis for helminth and protozoal parasites (such as Giardia; in addition Tritrichomonas in cats should be considered).

Further blood tests are indicated if none of the following tests shows any abnormalities: trypsin‐like immunoreactivity to exclude exocrine pancreatic insufficiency, pancreatic lipase immunoreactivity (Spec‐cPL, Spec‐fPL for dogs and cats respectively) to assess the possibility of pancreatic disease, ACTH test or basal cortisol concentration to exclude hypoadrenocorticism and cobalamin concentrations to assess the absorptive function of the distal small intestine. Total T4 and FeLV/FIV should also be assessed in cats. Abdominal ultrasound will be most helpful to determine whether the small and/or large intestine is affected and if there are any mass lesions that need surgical intervention rather than endoscopic evaluation. In the case of PLE, specific findings on ultrasound, such as speckles in the mucosa, can also be useful additional information. If the results of these tests do not point to an obvious cause for the clinical signs and the patient is stable (i.e., has a normal appetite, good attitude, not lethargic, no to minimal weight loss, normal serum protein concentration with no intestinal thickening on diagnostic imaging) then a well‐conducted therapeutic trial with an elimination diet or hydrolyzed diet for at least two weeks can be performed. If there is no response to a trial within two weeks after starting the diet, it is unlikely that the  patient is suffering from food‐responsive disease (food allergy or food intolerance). Intestinal biopsies for histopathology are collected from those patients that fail to respond to empirical therapy or that are showing worsening of their clinical signs. Most patients with chronic enteropathies can be diagnosed by obtaining endoscopic biopsies, as long as at least 12–15 biopsies from the duodenum, ileum, and/or colon are taken. It is important to realize that good‐quality biopsies are critical in order for histopathology to be useful, so more than 10 biopsies per site are usually recommended to make a diagnosis. In some rare cases, a diagnosis of lymphoma can be missed if no full‐thickness biopsies are obtained, especially in cats and if the ileum has not been sampled.

­Serum and Fecal Markers of Disease Serum Albumin Concentrations in Dogs Decreased serum albumin concentrations have been identified as a negative prognostic indicator in both retrospective and prospective studies of canine IBD. PLE accounts for the loss of albumin through the gut mucosa in severely affected dogs with IBD. PLE in dogs can be associated with severe lympho‐plasmacytic IBD, intestinal lymphoma, or, rarely, primary lymphangiectasia. One study described 12/80 (16%) dogs with hypoalbuminemia and 4/80 (5%) dogs with panhypoproteinemia.

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Box 59.2  Canine Chronic Enteropathy Clinical Activity Index (CCECAI) Attitude/activity 0 = normal 1 = slightly decreased 2 = moderately decreased 3 = severely decreased Appetite 0 = normal 1 = slightly decreased 2 = moderately decreased 3 = severely decreased Vomiting 0 = normal 1 = mild (1× per week) 2 = moderate (2–3× per week) 3 = severe (>3× per week) Stool consistency 0 = normal 1 = slightly soft feces or fecal blood, mucus or both 2 = very soft feces 3 = watery diarrhea Stool frequency 0 = normal 1 = slightly increased (2–3× per day) 2 = moderately increased (4–5× per day) 3 = severely increased (>5× per day) Weight loss 0 = none 1 = mild (10%) Albumin levels 0 = albumin >20 g/L 1 = albumin 15–19.9 g/L 2 = albumin 12–14.9 g/L 3 = albumin 2.0 g/dL) were already associated with an increased risk of refractoriness to treatment. At this level, most patients will not yet show any clinical signs of hypoalbuminemia, such as ascites, peripheral edema, or pleural effusion, but are already at risk of euthanasia because of intractable disease. Another study found that severely hypoalbuminemic dogs that had failed to improve on immunosuppressive doses of steroids were successfully treated with cyclosporin. This suggests that early aggressive treatment in hypoalbuminemic dogs may potentially decrease mortality rates in severely ill animals. Serum albumin concentrations can also be a good measure of monitoring such patients, as improvement of serum albumin concentrations above 2.0 g/dL usually indicates treatment success, even if clinical improvement has been seen ­earlier in some of these cases. It is therefore recommended to serially evaluate serum albumin concentrations every 2–3 weeks in order to  assess when treatment can be tapered off or discontinued. Serum Albumin Concentrations in Cats There is not much published information available regarding serum albumin concentrations in cats with chronic intestinal disease. PLE as a clinical syndrome likely does not exist in cats, as clinical signs such as ascites and peripheral edema do not usually occur in cats with hypoalbuminemia due to intestinal disease. In addition, the hypoalbuminemia seen in such cases is usually mild. However, there is accumulating evidence that cats with chronic intestinal disease and decreased albumin serum concentrations may have concurrent pancreatic disease. In one recent retrospective study, cats diagnosed with IBD and serum feline pancreatic lipase (Spec‐fPL) concentrations ≥12.0 μg/L had a lower median serum albumin concentration than cats with IBD and a normal Spec‐fPL concentration. Further analysis in this study did not identify hypoalbuminemia as a predictor of negative outcome on survival. Therefore, hypoalbuminemia in cats with chronic intestinal disease should prompt the clinician to measure

Spec‐fPL concentrations and/or to perform abdominal ultrasound examination in order to evaluate any concurrent pancreatic disorders. Depending on the severity of the hypoalbuminaemia, the clinician’s approach to ­treatment might be altered. Serum Cobalamin Concentrations in Dogs and Cats Serum cobalamin concentrations should be measured in any small animal patient with chronic intestinal disease. As cobalamin is absorbed in the ileum, decreased serum cobalamin concentrations are most commonly seen when this part of the small intestine is affected. However, absorption of cobalamin also involves intrinsic factor, which in dogs and cats is mainly produced in the pancreas. This is the reason why most small animals with exocrine pancreatic insufficiency will have low serum cobalamin concentrations. Furthermore, serum cobalamin concentrations have been shown to be a negative prognostic factors in dogs with CE. If cobalamin serum concentration is below the reference range, the risk for later euthanasia increases to a factor of 10. It is therefore important to supplement dogs with hypocobalaminemia while they undergo treatment for IBD, as this risk of euthanasia can be reversed by supplementation. Serum cobalamin concentration has long been known to be an important negative prognostic factor in cats with chronic enteropathies. The prevalence of decreased serum cobalamin concentrations in cats with chronic gastrointestinal signs has been reported to be 16–60%. In cats, it has also been reported that cobalamin supplementation can improve clinical signs regardless of the underlying diagnosis and even if given as the sole treatment for their disease. It is therefore recommended that cats with chronic intestinal disease be supplemented with cobalamin regardless of whether a specific cause for the disease can be identified. Recent studies have also shown preliminary data indicating that cobalamin can be given orally to supplement deficiencies. However, large amounts have to be given by this route of administration and concentrations decreased much more rapidly after discontinuation of oral treatment, compared to injectable forms. Supplementation Recommendations for Cobalamin Supplementation of cobalamin should be given parenterally (SC) as a weekly injection for at least six weeks. Exact dosages are not reported, as it is a water‐soluble vitamin and cannot be overdosed. For tested recommendations, please visit the website of the Texas GI lab (http://vetmed.tamu.edu/gilab).

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Canine Pancreatic Lipase Canine pancreatic lipase (now measured with the Spec‐ cPL assay) has recently become available as a commercial test and is useful in the assessment of possible pancreatitis in dogs. However, it can also be elevated in dogs with a primary diagnosis of CE. In a retrospective study of 50 dogs with IBD, clinical signs, age, serum lipase and amylase activities, albumin and cobalamin concentrations, abdominal ultrasound examination, histopathologic review of intestinal biopsies, management of IBD, and follow‐up in dogs with CE were evaluated. Sixteen dogs with increased cPL and 32 dogs with normal cPL values were compared. No significant differences were found in clinical activity score, serum amylase activity, serum lipase activity, serum cobalamin concentration, serum albumin concentration, abdominal ultrasound scores, and histopathology scores for IBD. There was also no difference in the frequency of steroid treatment between the groups. Interestingly, dogs with elevated Spec‐cPL had a higher risk of a poor follow‐up score and were significantly more likely to be euthanized at follow‐up. These data suggest that elevated Spec‐cPL in canine IBD identifies a subset of these patients that could also have chronic subclinical pancreatitis. In patients which have been diagnosed with CE and also have elevated Spec‐cPL without overt imaging evidence of acute pancreatitis, it is recommended to discuss treatment options for IBD that will also treat possible autoimmune pancreatitis. The author has had anecdotal success with cyclosporin at 5 mg/kg SID for eight weeks in such cases.

levels may also be hampered by increases of CRP related to diseases other than IBD. Fecal Alpha‐1‐Proteinase Inhibitor Fecal alpha‐1‐proteinase inhibitor (alpha‐1‐PI) can be used as a test for dogs in which the clinician suspects PLE but clinical signs are not overt. Alpha‐1‐proteinase inhibitor is a plasma protein of similar size to albumin. If the intestinal mucosal barrier is compromised and loss of protein into the intestinal lumen occurs, alpha‐1‐PI is lost at approximately the same rate as albumin. Unlike albumin, however, its proteinase inhibitor properties protect alpha‐1‐PI from being degraded by intestinal proteases. Therefore, it is able to persist throughout the intestinal transit and is passed undamaged in the feces, in which it can then be measured. Prompt diagnosis of PLE in a patient with IBD is important, as hypoalbuminemia is a risk factor for a negative outcome and the cause should be treated aggressively in order to improve survival. The alpha‐1‐PI assay is especially valuable in a patient with intestinal disease that also suffers from concurrent renal or hepatic disease. In these patients, measurement of fecal alpha‐1‐PI can help assess which portion, if any, of the protein loss can be attributed to the intestine. This test is so far only available at the Texas GI lab and ideally, three consecutive freshly collected, frozen fecal samples should be submitted in order to improve the accuracy of the test. This means that fecal alpha‐1‐PI is to date not a useful test for practitioners outside North America.

C‐Reactive Protein

Polymerase Chain Reaction for Antigen Receptor Rearrangement from Intestinal Biopsies

C‐reactive protein (CRP) is a serum acute phase protein that can be elevated in many different diseases. In people with IBD, several indices of clinical disease activity incorporate measurements of CRP. In dogs, a similar correlation between the Canine IBD Activity Index (CIBDAI) and CRP serum concentrations has been found in one large study incorporating 58 dogs. CRP was elevated when compared to normal in 28 of the 58 dogs in this study, which had CIBDAI scores above 5 (which represents mild to moderate disease activity). In these dogs CRP decreased significantly after successful treatment. In another study, CRP did not seem to be very helpful, as it was only elevated in 6/21 dogs with CE. Based on available published data, it is possible that CRP is useful as an additional assessment tool in severely ill patients, even though a large percentage of dogs with IBD do not show any elevations in CRP. Interpretation of elevated

Polymerase chain reaction for antigen receptor rearrangement (PARR) amplifies the highly variable T or B cell antigen receptor genes, and is used to detect the presence of a clonally expanded population of lymphocytes. This test has been advocated to be useful when applied on endoscopic biopsies if a diagnosis of intestinal lymphoma is suspected but not confirmed by conventional histopathology. It is, however, important to note that dogs with CE can in up to 20% of cases have biopsies which test positive for clonality. The test therefore needs to be evaluated as an additional measure to help with the differentiation of intestinal lymphoma versus CE, but should not be used as the sole differentiating factor or even the gold standard. Other tests that can help with this question are immunohistochemistry for B and T cells, abdominal ultrasound and possibly fine needle aspirates of enlarged mesenteric lymph nodes, if they are present.

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Perinuclear Antineutrophilic Cytoplasmic Antibodies Perinuclear antineutrophilic cytoplasmic antibodies (pANCA) have been useful in the diagnosis of human IBD for decades. These antibodies are serum autoantibodies similar to antinuclear antibodies (ANA), which seem to be more specific for intestinal disease than ANA in dogs. They are detected by indirect immunofluorescence assays where a typical pattern of perinuclear staining of canine granulocytes can be seen. In the first study which assessed the possible clinical usefulness of pANCA in dogs with IBD, sensitivity for pANCA was 0.51 and specificity ranged between 0.56 and 0.95. pANCA proved to be a highly specific marker for CE in dogs when the group of dogs with chronic diarrhea of other causes were tested against the group of dogs with CE (specificity 0.95). This is in agreement with reports from human medicine that show a specificity of up to 94% for pANCA when distinguishing between IBD and healthy controls as well as patients with non‐IBD diarrhea. Furthermore, when pANCA were tested in a group of dogs with FRD versus SRD, a positive pANCA titer was significantly associated with FRD. The pANCA assay could therefore provide valuable help to the veterinarian presented with the clinical picture of a dog with chronic diarrhea and possible IBD. If the result is positive, a food‐responsive chronic enteropathy is highly likely, but if the result is negative, CE cannot be excluded. pANCA also seem to be associated with the syndrome of familial protein‐losing enteropathy in soft‐coated wheaten terriers (SCWTs). pANCA were detectable in the serum of dogs on average 1–2 years before the onset of clinical disease and were highly correlated with hypoalbuminemia. This test could serve as a useful adjunct for this specific disease in SCWTs as an early screening test. Care must be taken in interpreting a positive pANCA test result if other inflammatory or immune‐mediated diseases are present in the dog. A recent study showed that many dogs with various vector‐borne diseases or immune‐mediated hemolytic anemia (IMHA) were positive for pANCA. Unfortunately, this test is not commercially available yet, but could become one of the standard tests for work‐up of CE in the future. Calprotectin and S100A12 Calprotectin and S100A12 are calcium‐binding proteins that are abundant in the granules of neutrophils and macrophages. In people with IBD, serum and fecal concentrations of these proteins have been found to be increased compared to healthy people. In addition, fecal

concentrations of calprotectin correlate very well with clinical disease activity in children with IBD. An immunoassay for measurement of canine calprotectin in serum and fecal samples is now available, and it was shown that a serum calprotectin concentration of ≥296.0 μg/L as a cut‐off had a sensitivity of 82.4% and specificity of 68.4% for distinguishing dogs with idiopathic IBD from healthy dogs. However, calprotectin concentrations were not significantly correlated with the clinical severity, serum C‐reactive protein concentration, or severity of histopathologic changes. The clinical usefulness of this test when used in serum or fecal samples still needs to be confirmed in prospective studies. Immunohistochemistry for P‐Glycoprotein on Intestinal Biopsies P‐glycoprotein (p‐gp) is a transmembrane protein functioning as a drug‐efflux pump in the intestinal epithelium. Human patients with IBD who fail to respond to treatment with glucocorticosteroids express high levels of p‐gp in lamina propria lymphocytes. So far, two research groups have evaluated p‐gp expression in biopsies of dogs with CE. In one study, duodenal biopsies from 48 dogs were evaluated by immunohistochemistry. The dogs treated with prednisolone showed a significantly higher p‐gp expression in lamina propria lymphocytes after treatment compared with expression before treatment. In contrast, the group treated solely with an elimination diet showed no difference in p‐gp scores before and after treatment. Moreover, a statistically significant association between refractoriness to steroid treatment and high p‐gp expression was found in the steroid treatment group. In another recent study, p‐ gp expression was compared between dogs with CE and healthy controls, and was found to be higher in duodenal epithelial cells of dogs with CE compared to control dogs. These results indicate that epithelial and lamina propria lymphocyte expression of p‐gp is upregulated in dogs with CE, and they are even higher after prednisolone treatment. In addition, high p‐gp expression could indicate possible multidrug resistance and should be taken into account when dealing with dogs who have failed steroid treatment before. Genetic Testing Over the last decade, numerous genes have been found to be associated with an increased risk of development of IBD in human beings, many of them implicated in the innate immune response in the intestine. In dogs, it has always been obvious to clinicians that IBD could have a genetic component. This is particularly evident in breeds like the boxer, which is predisposed to granulomatous

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colitis, and the German shepherd dog (GSD), which is predisposed to CE. Several mutations in TLR4 and TLR5 were found to be significantly associated with an increased risk of development of CE. In a follow‐up study, we were also able to show that peripheral blood cells of dogs carrying the mutation are hyperresponsive to flagellin, which is the natural ligand for TLR5. This proves for the first time that a genetic mutation implicated in the pathogenesis of dogs with IBD has functional consequences on the protein level. Taken together, these findings make it very likely that TLR5 mutations are causally associated with canine IBD. Genetic testing for these polymorphisms could become important for breeders and practitioners in the future. However, it is likely that in multifactorial disease like IBD in dogs, other factors such as other genetic mutations and environmental factors will also play a major role in the pathogenesis of the disease.

­Work‐up and Diagnostic Tests: Abdominal Ultrasound, Endoscopy, and Histopathology Diagnostic Imaging Ultrasonographic study is the preferred imaging tool for the investigation of animals with GI disease. The whole of the intestinal tract can be examined and the wall thickness measured. Although CE may cause focal or segmental changes, it does not always induce changes that can be detected ultrasonographically, so intestinal biopsy is required to confirm the diagnosis. Neoplastic infiltration is more often focal, shows more severe thickening, and causes loss of wall layering when compared to inflammatory disease. In addition, lymph nodes tend to be larger and more involved in neoplastic disease. However, the overlap in the sonographic appearances of inflammatory and neoplastic infiltration makes definitive diagnosis ­difficult, requiring histopathology for differentiation. Histology and World Small Animal Veterinary Association (WSAVA) Scoring of Intestinal Biopsies Sampling of intestinal biopsies is considered to be an essential step to exclude neoplastic causes and confirm the presence of intestinal inflammation. However, the interpretation of intestinal biopsies is difficult and subject to controversy. In several recent studies looking at conventional histologic interpretation of intestinal biopsy samples, no correlation of clinical activity with histologic grading either before or after therapy was found. In addition, total lymphocyte counts as well as the

number of infiltrating CD3 cells in the lamina propria cannot be used as markers for clinical activity of disease, as there is no difference in cell counts before and after treatment. These findings suggest that the type and degree of histologic infiltrates in canine IBD may not be as helpful as in human medicine, where the clinical scores correlate very well with the histologic grading. Therefore, a new grading scheme for the histologic interpretation of endoscopically obtained biopsies from dogs and cats with IBD has recently been published by the WSAVA working group. The findings in this study suggest that microarchitectural changes seem to be much more important than cellular infiltrates when assessing histologic severity of disease. However, so far, there is limited information on how well this new grading system correlates with clinical disease. This notion was recently confirmed in a prospective study including 20 dogs with FRD. A second endoscopy was performed in these dogs after six weeks of dietary treatment, and WSAVA scoring was the same as before treatment. However, when looking at electron microscopic changes, mitochondrial and brush border changes indicated mucosa healing which was not obvious on light microscopy. In one retrospective study, the interpathologist variability was still very high even when using the picture guide from the original publication. In addition, it is of concern that the only parameter that correlated with clinical disease was the presence of lymphangiectasia and hypoalbuminemia. Currently, there is lack of consensus between pathologists when using the WSAVA histopathologic guidelines for intestinal biopsies, and studies have demonstrated that these scores correlate poorly with clinical severity. Therefore, it is likely that additional criteria need to be assessed when scoring intestinal biopsies and further studies are needed to determine other immunologic components that are dysregulated in animals with CE.

­Treatment and Prognosis Inflammatory bowel disease patients that have mild to moderate clinical disease activity and normal serum albumin concentration are first treated sequentially with a dietary and antibiotic trial. If they fail to respond to either of these trials, immunosuppressive therapy is initiated (see following paragraphs). FRD: Diet A positive response to a dietary trial allows the patient’s CE to be classified as FRD, a term which includes both dietary allergy and intolerance. The main options for a dietary trial include switching to a diet which leads to

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antigenic modification (e.g., novel protein source or protein hydrolysate). Whichever type of diet is chosen, it must be palatable and introduced in gradually increasing amounts over 4–7 days. In dogs with FRD, a clinical response is usually observed within 1–2 weeks of changing the diet. One study demonstrated that dogs that respond to diet tended to be younger and had higher serum albumin concentrations and predominantly signs of large bowel diarrhea than dogs that did not respond to diet. FRD is highly prevalent among dogs with IBD (at least 60–70%) and a favorable response to elimination or hydrolyzed diets within two weeks has been shown to be associated with a very good prognosis over one year after diagnosis. It is important to note that in the studies that show these good outcome measures, the dogs were kept on the diet for at least 12 weeks after diagnosis before they were switched back to their original diet. ARD: Antibiotics An antibiotic trial typically involves oral administration of tylosin, 10–15 mg/kg q8h, oxytetracycline 20 mg/kg q8h, or metronidazole 10  mg/kg q12h. A positive response suggests ARD. The dog is typically maintained on antibiotics for 28 days; if signs recur after stopping then long‐term antibiotic therapy with tylosin 5 mg/kg is used orally every 24 hours. In a recent large retrospective study where all dogs were sequentially treated, only 16% of dogs were ARD. All ARD dogs relapsed shortly after discontinuation of the antibiotics was attempted, making long‐term management of these patients difficult. An additional decision‐making factor may be the increasing problems with antibiotic resistance in our dog populations, making justifications of long‐term treatment with antibiotics difficult. There is also accumulating evidence that antibiotic treatment has long‐lasting effects on the intestinal microbiome, which may lead to lasting dysbiosis and in itself could amplify inflammation in the intestine. Many of these patients will therefore need steroids or other immunosuppressives to control their clinical signs long term. Anti-inflammatory and Immunosuppressive Therapy Patients that do not respond to a diet or antibiotic trial mg/kg are usually administered oral prednisolone 2  every 24 hours that is tapered over an eight‐week period. However, as the side‐effects of glucocorticoids are usually more marked in large‐ than small‐breed dogs, azathioprine may be combined with glucocorticoid treatment at a faster taper in dogs weighing more than 30 kg. If there is poor response to immunosuppression or a relapse is seen after tapering, then consider oral

c­ yclosporin at 5 mg/kg every 24 hours for 10 weeks. A response to prednisolone has been shown in up to only 50% of dogs with CE, so if more severe disease is present or severe side‐effects of steroids are anticipated, other immunosuppressives can be an option. Many steroid‐ refractory canine CE cases can be rescued by cyclosporin single therapy. In cats, use chlorambucil 2–6 mg/m2 q24h with prednisolone if there is inadequate response to glucocorticoid treatment alone. Hematologic parameters should be monitored regularly if chlorambucil is used. If the patient responds then the medication can be tapered gradually, starting with the steroid, to an q48h dosing regimen. Budesonide is a glucocorticoid medication that has been preliminarily shown to be successful in the treatment of canine IBD. However, hypothalamic‐pituitary‐ adrenal suppression and development of a steroid hepatopathy have been demonstrated in dogs, so the hepatic fist‐pass effect of this drug in dogs may not be as great as in human beings. An optimal dose has not yet been determined, although anecdotally a dose of 1 mg/ m2 every 24 hours orally has been recommended. The response rate to budesonide was shown to be similar to the use of prednisolone (about up to 60%), and it should therefore be reserved for dogs that are known to respond to steroids but suffer severe side‐effects. Some dogs will, however, still develop side‐effects of steroid administration while on budesonide, so owners should be warned about this. Sulfasalazine (20–50 mg/kg q8h for 3–6 weeks) and related drugs are often used in dogs when IBD is limited to the large intestine. However, as side‐ effects include keratoconjunctivits sicca, tear production should be monitored regularly. Treatment of Patients with Severe Protein‐ Losing Enteropathy Protein‐losing enteropathy is a recognized complication in a subset of CE cases and a low serum albumin concentration has been shown to be a poor prognostic indicator for CE. Patients with albumin concentrations below 1.5 g/dL are at risk of developing ascites, pleural effusion, and subcutaneous edema. Many of these patients will succumb to the disease within the first 1–2 months of starting prednisolone treatment. As some studies have shown better outcome with single‐therapy cyclosporin at 5–10 mg/kg PO SID, this latter regime may be a better option for many of these patients. One recent study has also shown that the combination of prednisolone and chlorambucil was superior to prednisolone and azathioprine for survival. Evaluation of hemostatic function in these patients is recommended to ascertain if hypercoagulability has developed as a consequence of enteric protein loss. Concurrent therapy with ultra‐low-dose

59  Diagnosis and Management of Chronic Enteropathies

aspirin (0.5 mg/kg) or other platelet inhibitors is recommended in these patients in order to prevent thromboembolism. In addition, the use of elemental diets and partial ­parenteral nutrition may be indicated in some dogs with severe PLE. It is worth mentioning that some PLE patients can fare relatively well with dietary treatment alone, and there are some studies that show that Yorkshire terriers may be a subgroup of dogs developing PLE which can be solely diet responsive. In such cases, it is indicated to try a ­low‐ fat diet first, and wait before adding in immunosuppressive treatment for 1–2 weeks. Finally, these patients may also be at risk of complications associated with intestinal biopsy by laparotomy, so plasma transfusion, human albumin infusion or synthetic colloid may be indicated during anesthesia for endoscopy. Adjunctive Therapy The use of probiotics in people with IBD has led to some promising results, although there is still an insufficient number of large, multicenter, randomized, double‐blind, placebo‐controlled trials that investigate the efficacy of

probiotics in human IBD. Similarly, only one randomized, placebo‐controlled trial investigating the use of E. faecium probiotics in canine FRD as an adjunctive treatment has been performed so far. No additional effect could be demonstrated in the group of dogs receiving probiotics. More studies are needed to be able to make recommendations for the use of probiotics in cats and dogs with CE (see also Chapter 58). Prognosis One retrospective study demonstrated that only 26% of canine CE cases progress to complete remission, with intermittent clinical signs remaining in approximately half of cases, and 4% were completely uncontrolled, with 13% being euthanized because of poor response to treatment. This suggests that the prognosis of CE patients can be poor. The main negative prognostic indicator for CE was identified to be hypoalbuminemia in dogs. Clearly, more prospective treatment trials are necessary, especially in the severely affected and hypoproteinemic animals, in order to improve long‐term survival in these cases.

­Further Reading Allenspach K, Rüfenacht S, Sauter S, et al. Pharmacokinetics and clinical efficacy of cyclosporine treatment of dogs with steroid‐refractory inflammatory bowel disease. J Vet Intern Med 2006; 20(2): 239–44. Allenspach K, Wieland B, Gröne A, et al. Chronic enteropathies in dogs: evaluation of risk factors for negative outcome. J Vet Intern Med 2007; 21(4): 700–8. Day MJ, Bilzer T, Mansell J, et al. Histopathological standards for the diagnosis of gastrointestinal inflammation in endoscopic biopsy samples from the dog and cat: a report from the World Small Animal Veterinary Association Gastrointestinal Standardization Group. J Comp Pathol 2008; 138(Suppl 1):S1–43. Janeczko S, Atwater D, Bogel E, et al. The relationship of mucosal bacteria to duodenal histopathology, cytokine mRNA, and clinical disease activity in cats with inflammatory bowel disease. Vet Microbiol 2008; 128(1–2): 178–93.

Minamoto Y, Otoni CC, Steelman SM, et al. Alteration of the fecal microbiota and serum metabolite profiles in dogs with idiopathic inflammatory bowel disease. Gut Microbes 2015; 6(1): 33–47. Simmerson SM, Armstrong PJ, Wünschmann A, et al. Clinical features, intestinal histopathology, and outcome in protein‐losing enteropathy in Yorkshire Terrier dogs. J Vet Intern Med 2014; 28(2): 331–7. Washabau RJ, Day MJ, Willard MD, et al. Endoscopic, biopsy, and histopathologic guidelines for the evaluation of gastrointestinal inflammation in companion animals. J Vet Intern Med 2010; 24(1): 10–26. Worhunsky P, Toulza O, Rishniw M, et al. The relationship of serum cobalamin to methylmalonic acid concentrations and clinical variables in cats. J Vet Intern Med 2013; 27(5): 1056–63.

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60 Approach to the Patient with Liver Disease Emma O’Neill, BSc, BVSc, PhD, DSAM, DECVIM-CA, MRCVS School of Veterinary Medicine, University College Dublin, Dublin, Eire

The liver plays a central homeostatic role within the body, involved in the orchestration of a vast array of ­metabolic pathways. The metabolic functions performed include carbohydrate, protein and lipid metabolism, drug metabolism, vitamin storage and activation, endocrine hormone metabolism, removal of endogenous and exogenous toxins (e.g., ammonia) along with excretory, digestive, and storage functions (e.g., iron, copper, ­glycogen, triglyceride). The liver is also the site of synthesis of albumin and the majority of clotting factors and plays a key role in immune surveillance. The liver’s position, between the gastrointestinal tract and the systemic circulation, aids its roles in digestion, detoxification, and immune surveillance. However, it also leaves the liver vulnerable to injury from drug or toxin exposure and prone to the secondary effects of ­dysfunction at other sites in the body. The normal liver is typically functioning with significant reserve capacity (approximately 70–80%), providing an important buffer should an animal sustain liver injury. In addition, the liver has a huge regenerative capacity, affording the ­animal significant ability to recover from injury. All of these features of liver function have an important impact on the clinical presentation of animals with liver disease. Symptoms of liver disease typically only become apparent once the reserve capacity of the liver has been exceeded, hence diseases may remain subclinical for long periods of time, with animals presenting later in the disease process. The vast array of liver functions is reflected in the wide variety of possible clinical symptoms. The early clinical signs typically observed include intermittent anorexia, polyuria/polydipsia, vomiting, and lethargy, all nonspecific signs that could be referable to other body systems. More specific clinical signs, such as jaundice or ascites, tend to occur later in the disease process. For example, jaundice occurs in only approximately 20%

of dogs and 30–40% of cats with hepatobiliary disease and hence, although more specific, it is an insensitive indicator of hepatobiliary disease. The aim when approaching patients is to maintain a clinical suspicion of liver disease, particularly in patients with vague, waxing and waning signs, from a breed with known ­predispositions to hepatobiliary disease. The important challenge when approaching these patients is to diagnose the disease early in its course in order to provide a chance to intervene before the disease becomes too advanced and irreversible. Hepatobiliary diseases rarely present with clinical features pathognomonic for a particular disease. However, there are distinct groups of clinical signs that can be appreciated to occur, reflecting important pathophysiologic mechanisms involved in liver disease. In view of this, a basic understanding of these mechanisms is helpful in order to understand both the clinical presentation of patients and the choice of tests to used diagnose them.

­ athophysiology of Important Clinical P Presentations of Liver Disease Cholestasis Cholestasis is the term used for impaired bile flow, which can result from a wide variety of intra‐ and extrahepatic causes. Normal bile production starts with canalicular bile formation. The active secretion of bile salts by hepatocytes into the bile canaliculi results in a large osmotic gradient and secondary movement of water, which in turn drives bile flow. The bile flows sequentially along the biliary tree through bile ductules, various ducts and then finally into the common bile duct. Throughout this passage, it is modified by secretion and reabsorption of

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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fluid and inorganic solutes. The bile ducts contribute substantially to this bile formation and flow, with secretion of bicarbonate, chloride, and water. Approximately 50% of bile is released immediately into the gastrointestinal tract, while the remainder is stored and concentrated in the gallbladder. Intrahepatic cholestasis disrupts this bile flow mainly at the level of the hepatocytes, their canaliculi or the bile ductules, in the periportal zone (zone 1) of the liver lobule, whereas extrahepatic cholestasis results from obstruction of the common bile duct. In both of these situations, significant hepatic reserve and compensatory changes have the net result that, typically, overt clinical signs of cholestasis, such as icterus (jaundice), only result from diffuse liver pathology or severe impairment of the common bile duct flow. Intrahepatic cholestasis occurs commonly to varying degrees in most clinical cases, with various proposed mechanisms involved. Damage to tight junctions separating the bile canaliculi from blood sinusoids can result from endotoxemia, sepsis, some Leptospira infections or following an adverse drug reaction. At the cellular level, swollen hepatocytes may directly obstruct bile flow through canaliculi or bile ductules. In addition, necrosis of hepatocytes may disrupt hepatic architecture, allowing communication between canaliculi and sinusoidal lymph or blood, aided by the inherent biliary pressure. Physical obstruction to bile flow from lobules can occur with any processes causing accumulation of tissue (inflammatory, neoplastic or collagen) in portal or periportal regions. This obstruction can occur at any level in the biliary tree with any process disrupting the normal architecture, for example in cirrhosis or with metastatic tumors. Extrahepatic cholestasis is far less common, with important causes including biliary or pancreatic adenocarcinoma, duodenal disease or pancreatitis causing blockage of the duodenal papilla. Cholestasis can be recognized clinically in several ways which can be helpful to consider when approaching a clinical case. Severe cholestasis results in icterus. Less severe cholestasis can be recognized in varying degrees on the biochemistry panel. This is discussed further in the section on biochemical evaluation, but it is important to be aware of the frequency with which some degree of cholestasis occurs in a wide variety of liver disease. Icterus Icterus (or jaundice) is the yellow discoloration of tissues resulting from hyperbilirubinemia and the build‐up of bile pigments (Figure 60.1). Bilirubin is the major bile pigment and is the normal end‐product of the catabolism of hemoproteins, a broad group of molecules including hemoglobin, myoglobin, and many enzymes located within the liver, (e.g.,

Figure 60.1  Canine eye showing jaundiced sclera. Source: Photograph courtesy of Sheila Brennan.

cytochromes). Senescent or damaged erythrocytes are removed from the circulation by cells of the mononuclear phagocyte system located within the spleen, liver, and bone marrow. These cells break down hemoglobin and release unconjugated, hydrophobic bilirubin into the circulation where it is carried, reversibly bound to albumin, to the liver for clearance. The hepatocytes take up the bilirubin and conjugate it, predominantly with glucuronide, to aid solubility for biliary excretion. This conjugated bilirubin is released into the intestine where it is not reabsorbed but is either excreted unchanged or degraded by intestinal bacteria to urobilinogen and then the stercobilins (brown fecal pigments). Hyperbilirubinemia results from disruption of this normal production and/or processing of bilirubin and  can be considered to result from three broad mechanisms. ●●

●●

●●

Prehepatic causes where there is increased production of bilirubin in excess of the hepatic capacity for excretion, for example severe hemolytic anemia. Hepatic causes where there is impaired uptake, conjugation, and excretion of bilirubin as a result of marked cholestasis. As described in the section on cholestasis, extrahepatic causes such as sepsis can also result in intrahepatic cholestasis due to cytokines directly inhibiting bilirubin transport. Posthepatic causes reflect reduced excretion of ­bilirubin due to disruption of bile flow within the extrahepatic bile ducts.

60  Approach to the Patient with Liver Disease

Portal Hypertension Portal hypertension is abnormally high pressure within the portal circulation, the result of increased blood flow or resistance within the portal circulation or a combination of the two. It is an important consequence of a ­variety of clinical conditions, including liver disease, and is associated with significant morbidity and mortality. The development of multiple acquired ­ portosystemic shunts (MAPSS) and ascites with or ­ without hepatic encephalopathy (HE) are important sequelae of the condition and are essential to recognize clinically from a diagnostic, prognostic, and treatment perspective. In health, the portal blood flow to the liver supplies 75–80% of liver blood supply through a low‐pressure system. The portal pressure is kept low and relatively constant, despite variations in portal blood flow, by a large sinusoidal reserve capacity and the ability to adapt the blood flow as necessary to maintain compliance. In disease, changes in this compliance alter the inherent resistance to blood flow and hence the portal pressure. These changes may be static, in the form of structural changes such as fibrosis, or dynamic, such as increased production of inflammatory mediators and vasoconstrictor agents along with reduced production of ­vasodilator agents. In most cases, there will be a combination of these processes occurring progressively with activation of hepatic stellate cells and progressive injury. In addition, splanchnic vasculature tends to undergo progressive vasodilation, as a result of many of the ­vasoactive agents released, further aggravating portal hypertension due to increased portal blood flow. Ascites An important consequence of significant portal hypertension is ascites. Starling’s law describes that the net movement of fluid between compartments is proportional to the balance between the relative hydrostatic and oncotic pressures in each region. Hence, the forces keeping fluid within the vascular space become imbalanced, with portal hypertension tending to favor fluid movement into the interstitium. Ascites develops once this fluid movement exceeds the capacity of the local lymphatic system. This situation is exacerbated by splanchnic vasodilation, which lowers the effective circulating blood volume due to pooling of blood. Over time, compensatory cascades, including the renin‐angiotensin‐ aldosterone system, sympathetic nervous system and antidiuretic hormone release, produce blood volume expansion, further favoring the formation of ascites. Hypoalbuminemia and reduced oncotic pressure also tend to favor net fluid movement and the development

of ascites, although typically this does not occur until albumin concentrations are below about 1.5 g/dL. These levels would be unusual for patients with liver dysfunction, occurring only with severe disease. A situation that happens more frequently in patients with liver disease is that a moderately reduced serum albumin (1.8–2.2 g/dL) occurs in combination with portal hypertension, ­resulting in exacerbation of the ascites. Acquired Portosystemic Shunts As portal hypertension reaches a threshold level, small embryonic vessels open up within the omentum and mesentery, allowing blood flow between the portal ­circulation and the vena cava. These acquired portosystemic shunts (APSS) may initially lower the portal venous pressure, but this effect is quickly mitigated by increased portal blood flow. The clinical importance of these APSS is that they result in a c­ ollateral circuit that effectively short‐circuits the liver. The net result is the delivery of toxins from the gastrointestinal tract directly into the systemic circulation. These toxins can trigger the vomiting center, causing nausea, vomiting, and inappetence. Clinical signs of HE may also develop. Hepatic Encephalopathy Hepatic encephalopathy is a clinical syndrome of brain dysfunction occurring secondary to liver dysfunction. It is a fairly common cause of morbidity and potentially mortality in dogs with liver disease, occurring less frequently in cats. The clinical signs of HE can be quite variable, with early signs being quite nonspecific, for example, apathy or reduced alertness. In more advanced cases, circling, head pressing, altered mentation, salivation, seizures, stupor, and coma may occur (Figure 60.2). The reserve capacity of liver function is such that, ­typically, HE only develops when there is concurrent significant impairment of parenchymal liver function ­ and portosystemic shunting, a situation that can occur in patients with congenital or acquired portosystemic shunts (PSS). More rarely, HE can result from fulminant hepatic failure or inborn errors of ammonia metabolism. In cats, an obligatory requirement for arginine to allow normal function of the urea cycle means that arginine deficiency in this species (e.g., the result of anorexia) can also result in HE. The pathogenesis of HE is complex and likely multifactorial. Normally, the liver removes neurotoxic substances from the portal circulation, preventing their delivery to the brain in the systemic circulation. HE develops with failure of this mechanism, the resultant delivery of toxins  triggering neurotransmitter dysfunction and a

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Coagulopathy

Figure 60.2  Dalmatian showing head pressing as a manifestation of hepatic encephalopathy.

constellation of potential clinical signs ranging in severity from apathy and reduced awareness to coma and unresponsiveness. The most commonly implicated toxin in HE is ammonia, although there are some cases where the plasma ammonia concentration is normal or only slightly raised despite signs of HE. Other potential triggers that have been studied include mercaptans, ­ manganese, systemic inflammation, altered gamma‐ aminobutyric acid (GABA, an inhibitory neurotransmitter), benzodiazepine receptor signaling and altered catecholaminergic signaling due to changes in branched chain aromatic amino acid profiles. Whatever the ­underlying cause, the overall result is a disruption of brain function, typically leading to waxing and waning clinical signs. Gastroduodenal Ulceration Patients with hepatobiliary disease are predisposed to gastric and duodenal ulceration for a number of reasons. Portal venous hypertension is an important risk factor resulting in vascular stasis and venous congestion, which together increase the risk of mucosal ulceration due to reduced mucosal blood flow and secondarily reduced bicarbonate and mucus secretion. This risk is further increased in anorexic animals or those exposed to nonsteroidal antiinflammatory drugs or potentially corticosteroids. Gastrointestinal ulceration and bleeding is an important consideration in any patient with HE that suddenly deteriorates or in any patient with liver disease that becomes more subdued or develops a reduced appetite.

The liver is involved in the manufacture of the majority of the coagulation factors, including fibrinogen (factor I), prothrombin (factor II) and factors V, VII, IX, X, XI, and XIII. The site of factor VIII production is controversial, but it is likely that the liver plays an important role in its production. It is also the site of the vitamin K‐dependent activation of factors II, VII, IX, and X and protein C. In addition to this, the liver is involved in the clearance of activated clotting factors and the production of clotting factor inhibitors (e.g., antithrombin and alpha‐1‐antitrypsin) and fibrinolytic proteins such as plasminogen. Hence, in liver disease there is the potential for disrupted production of coagulation factors and their inhibitors along with impaired clearance of both activated coagulation factors and the products of the fibrinolytic system. The extent of the resulting abnormalities reflect the severity of the underlying liver disease, ranging from individual factor deficiencies to disseminated intravascular coagulation (DIC). A study of 42 dogs with histologically confirmed liver disease demonstrated that one or more coagulation abnormalities was present in 57% of the dogs. These abnormalities ranged from single factor deficiencies in four of the dogs to DIC in three dogs. An older study suggested a figure of 93% of dogs with liver disease having abnormal coagulation parameters, although this study involved more severely affected cases. A study of 45 cats with liver disease showed that 98% of the cats had one or more abnormality detected. What is important to recognize clinically is that, whilst spontaneous hemorrhage may be rare in animals with hepatic disease, hemostatic abnormalities occur commonly. Hence, the clinician needs to exercise caution when considering interventional diagnostic procedures and a coagulation profile should be performed in all such cases.

­Signalment Patient signalment is very important when approaching the patient with hepatobiliary disease. There are a variety of known or suspected breed predispositions to hepatic disease which can be helpful to consider, particularly in view of the occult nature of many cases of early liver disease. Congenital portosystemic shunts (CPSS) have worldwide breed predispositions including Irish wolfhounds, cairn terriers, Labrador retrievers, dachshunds, Yorkshire terriers, Australian cattle dogs, Maltese terriers, and miniature schnauzers. In cats, the condition is seen most frequently in mixed‐breed cats, although Persian and Himalayan breeds are also overrepresented. Additionally, copper‐associated hepatitis has been

60  Approach to the Patient with Liver Disease

described in several breeds, including the Bedlington terrier, Dalmatian, Doberman pinscher, Labrador retriever, West Highland white, and Skye terriers. A recent study documented breeds at risk of developing chronic hepatitis in the UK (see Chapter  64). These included three breeds in agreement with a previous, older study from Sweden: the American and English cocker spaniel, Doberman pinscher and Labrador retriever, and additionally highlighted an increased risk in the cairn terrier, English springer spaniel, Great Dane, and Samoyed. However, it should be recognized that the gene pools in different countries can vary, such that certain breed characteristics and predispositions may be more regional. In addition to the breed, the age at onset of clinical signs can be very helpful in aiding diagnosis of some hepatobiliary conditions. For example, animals with CPSS typically present at a younger age. This has classically been described to be less than 1 year of age, although increasingly older dogs seem to be recognized, presumably as clinicians are looking more widely for the condition. There have been reports of increasing numbers of miniature schnauzers not being diagnosed until they are over 7 years, presenting with only mild signs. Reports looking at the age and gender distribution of dogs presenting with chronic hepatitis have shown the mean age at diagnosis to be approximately 8 years when all breeds were considered together, although some breeds tended to present at a younger age (e.g., English springer spaniel). In general, females are overrepresented, although again there are occasional breeds where this does not appear to be the case.

access to toxins and a history of any recent drug administration are extremely relevant. The clinician should maintain a high index of suspicion whenever a patient develops new clinical signs within four weeks of starting a new drug. There are many known potentially hepatotoxic drugs, common examples being phenobarbital and  itraconazole, but most drugs have the potential for  an idiosyncratic reaction in an individual patient. Consumption of a hepatotoxin is also possible, such as certain mushrooms, xylitol (contained in sugar‐free sweets) or even herbal/dietary supplements that the owner might be administering. Hepatobiliary disease can result in a very broad range of clinical signs reflecting both the sheer variety of functions the liver performs and the central role it plays in many metabolic and detoxifaction pathways. This central role means that hepatic disease can result in the secondary dysfunction of other organs, as in HE. It also places the liver in a position where it can itself become secondarily affected, for example with hyperadrenocorticism or hepatic lipidosis. The net result is that the clinician needs to be alert to this when approaching a case, aiming to first decide if hepatobiliary disease could be present in an animal but then, importantly, whether this represents a primary or secondary phenomenon. These key ­questions sound very straightforward, but they can be  challenging to address in many patients and will be reconsidered as different aspects of the investigation of liver disease are explored.

­Diagnosis ­History and Clinical Signs A thorough history with good attention to detail is key to approaching any clinical case. This is particularly pertinent with potential hepatobiliary cases for a variety of reasons. As outlined in the introduction, the large hepatic functional reserve means that in order to diagnose diseases before there is significant, irreversible loss of healthy liver tissue, the clinician needs to identify cases with subtle and often vague clinical signs. Symptoms such as anorexia, vomiting, lethargy or mild polyuria/polydipsia, which are often waxing and waning, should alert the clinician that liver disease is a potential differential diagnosis. The onset of clinical signs may ­initially appear acute in nature but careful discussion with the owner might elucidate a more chronic background than initially described. Box  60.1 summarizes clinical signs that may be observed in animals with liver disease. The potential for toxin exposure should always be considered so a detailed description of the animal’s potential

Biochemical Evaluation The employment of clinical pathologic testing is central to the approach to animals with hepatobiliary disease. In many cases, recognition of abnormal test results often provides the first indication that liver disease may be present. It is not uncommon to detect high liver enzyme activities as the only abnormality on preanesthetic or health screens, prompting further investigation. In addition, as outlined above, in view of the typical presentation of hepatobiliary cases, it is important to use clinical pathology to aid in the diagnosis is liver disease early on in the disease process. Often at this stage the clinical signs may be vague and nonspecific with clinical pathology aiding greatly in the decision making and further defining of such cases. The laboratory evaluation of hepatobiliary patients has several broad aims: to determine/confirm the presence of hepatobiliary disease and assess hepatic function, to aid in distinguishing primary or secondary hepatic disease and to contribute to diagnostic planning or ­

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Box 60.1  Clinical signs and clinical examination findings that may occur in animals with liver disease Vague signs Vaguely “off form” Lethargy Signs related to gastrointestinal tract Vomiting Diarrhea Anorexia Weight loss GI bleeding ‐ melena Signs related to cholestasis Jaundice Acholic feces ‐ with complete extrahepatic biliary obstruction Signs related to hepatic encephalopathy Vague signs of dullness Alterations in behavior Ptyalism – particularly in cats Head pressing Disorientation Stupor Seizures Vague neurologic signs, seizures or syncope may occur with hypoglycemia Signs relating to CPSS Hepatic encephalopathy signs (see above) Stunting Microhepatica

­ onitoring response to treatment. Occasionally, diagm nostic testing may allow definitive diagnosis of hepatobiliary disease although more typically, it acts as a piece within the overall jigsaw. The biochemical tests directly utilized in the evaluation of liver disease comprise the liver enzymes and liver function tests, which appraise impaired metabolic function or synthetic capacity. However, evaluation of the full hematology and biochemistry panel is important to gain insight into the ­animal’s general health. This aids clarification of the likelihood of another disease process that may either be the cause of a secondary hepatopathy or represent an additional consideration in patient management. Enzyme Markers of Liver Disease

The liver enzymes evaluated can be divided into two main groups dependent on their cellular location and clinical utility. Alanine aminotransferase (ALT) and aspartate aminotransferase (AST) are the two most commonly measured markers of hepatocellular injury and are found primarily within the hepatic cytosol. Alkaline phosphatase (ALP) and gamma‐glutamyl transferase (GGT) are the most commonly measured biliary

Dysuria – relating to ammonium biurate urolithiasis Hematuria – relating to ammonium biurate urolithiasis Copper‐colored iris (some cats) Signs relating to disorders of hemostasis Gastrointestinal bleeding ‐ melena Increased bleeding from invasive procedures Signs relating to the urinary tract Polyuria/polydipsia Discolored urine – jaundice/hematuria Dysuria (see above) May have prolonged recovery from sedation/general anesthesia Clinical findings Poor body condition score Mucous membrane pallor Abdominal discomfort – organomegaly, abdominal distension, inflammation (hepatic/peritonitis/ cholecystitis) Pyrexia – with some inflammatory, infectious or neoplastic causes Ascites ‐ relating to portal hypertension, hypoalbuminemia, biliary peritonitis or neoplastic effusion Hepatomegaly +/‐ irregularity of liver margins – with infiltration, some inflammatory conditions Skin lesions may be seen with hepatocutaneous syndrome (superficial, necrolytic dermatitis)

enzymes. They are located mainly on hepatobiliary membranes and are markers of cholestasis or drug induction (see later). In general, the liver enzymes are sensitive indicators of liver disease or injury but are not specific. There are far more animals in which elevated liver enzyme activities are detected than actually have clinically significant liver disease. This lack of specificity arises from a combination of the susceptibility of the liver to secondary or “reactive” disorders, the ability of certain hormones or drugs, such as corticosteroids, to induce production of particular liver enzymes and the presence of isoenzymes within tissues other than liver. The clinical interpretation of the significance of liver enzyme elevations is challenging and must always be considered in the context of the patent, its history, and the full clinical pathology results. It is important to realize that the liver enzymes do not offer any indication of liver function. In an animal with a primary hepatopathy, the magnitude of hepatocellular enzyme release may broadly relate to the number of hepatocytes affected, particularly in the acute setting. However, in view of the regenerative capacity of the liver, the magnitude offers no prognostic information. It is also

60  Approach to the Patient with Liver Disease

important to appreciate that in severe chronic disease, such as cirrhosis, the liver enzyme release may be minimal due to reduced hepatic synthetic capacity. ­ In  addition, the serial evaluation of serum enzyme ­activities can be more useful than a single time point in determining the importance of liver enzyme elevation and the likely prognosis in a patient. It is generally considered a favorable sign if values continue to fall gradually after a single liver insult whereas ongoing persistent elevation may indicate a chronic hepatopathy and the requirement for further investigation. The serum enzyme activity at any point in time is dependent on the overall hepatic activity of the enzyme or the capacity for de novo synthesis (in response to cholestasis or drug induction), the serum half‐life of the enzyme, and the cellular location of the enzyme (determining the ease of release of the enzyme, e.g., “leakage” of cytosolic enzymes). These factors vary not only between different enzymes but also between species, as will be discussed below. Hence, it is important to ensure species‐specific criteria are being used when interpreting the values in a given patient. When evaluating the liver enzymes, it is important to consider the pattern of enzyme increase (hepatocellular or cholestatic), the magnitude of this change and the temporal pattern of the enzyme changes over time. For example, are they decreasing gradually, fluctuating or progressively increasing? The reference intervals for liver enzymes vary considerably between laboratories due to different assay methods. In view of this, it is more appropriate to consider the magnitude of increase in relation to the upper limit of the reference interval when comparing values between labs. Hepatocellular/Leakage Enzymes

Alanine aminotrans­ferase is a cytosolic enzyme that is released into the serum from hepatocytes with increased hepatocyte membrane permeability or following hepatocellular necrosis. This enzyme is considered to be a sensitive indicator of hepatocellular injury in dogs and cats and is also considered to be the most liver‐specific enzyme in these species. Although ALT is also found in cardiac and skeletal muscle and the kidneys, this is not generally of clinical significance as the isoenzymes found in these other locations either have short half‐lives or are present at low concentrations. However, occasionally severe muscle injury can result in ALT elevation. The half‐life of ALT in the dog is controversial, with reports ranging from about six hours to 2.5 days. What is clear is that the half‐life is considerably shorter in the cat, with the result that smaller elevations are far more clinically significant in this species. The highest increases in ALT activity, in either species, are seen during acute hepatic inflammation or necrosis, the magnitude of

e­ levation, in this setting, being roughly proportional to the number of hepatocytes affected but providing no insight into the severity of injury or its reversibility. Rapid elevations in enzyme activity occur following changes in hepatocyte membrane permeability. Following experimental toxin administration, values rise sharply within 24–48 hours, peaking at values up to 100 times normal within approximately five days and then falling back to normal within 2–3 weeks. In general, with hepatic injury, a reduction in the enzyme activity of approximately 50% every 2–3 days would be viewed as a good prognostic sign. Elevated ALT activity can also occur with cholestasis due to the damaging effect of accumulated bile acids on hepatocyte membranes. The elevations that occur in this setting are typically more gradual in onset and are not as dramatic overall. Certain drugs can also result in ALT enzyme increases, for ­example anticonvulsant drugs or corticosteroids. These elevations tend to be dose dependent but also show considerable inter individual variation. The elevations seen with phenobarbital administration would typically be within the 4–5‐fold range compared to a 10‐fold or higher elevation occurring in animals with hepatotoxicity. However, the results need to be interpreted in light of the overall clinical picture and in combination with liver function evaluation where there is any concern. The interpretation of ALT elevation in the clinical setting frequently poses a diagnostic challenge. Persistent elevations of ALT are characteristic of canine chronic hepatitis and can be the only abnormality noted on a blood panel in animals in the early stages of the disease. Hence, elevations in excess of twice the upper limit of the reference range occurring over 1–2 months are potentially significant and should not be disregarded. However, frequently up to five‐fold elevations in ALT are observed in dogs with primary gastrointestinal diseases such as inflammatory bowel disease or pancreatitis. In addition, hemolytic anemia or cardiovascular disease can cause significant elevations in ALT secondary to hypoxia. Endotoxemia or sepsis can cause secondary hepatic changes, as can many endocrinopathies. These examples demonstrate the limitations of ALT; while it is very sensitive for the detection of hepatocellular insult and hence cases with potential primary hepatopathies, it is equally sensitive for the detection of secondary hepatopathies and “reactive” liver changes occurring in response to other systemic illness or drug exposure. In view of this, the clinician should interpret ALT activity in combination with the remainder of the clinical biochemistry, knowledge of the animal’s breed and history along with maintaining an awareness of the potential interference of extrahepatic disease. Aspartate aminotransferase is also a marker of hepatocellular damage within the group of “leakage” enzymes,

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Section 7  Diseases of the Liver, Gallbladder, and Bile Ducts

having a predominantly cytosolic location within hepatocytes. However, there is in addition a mitochondrial fraction, comprising about 30% of the total hepatic enzyme activity, which is only released during hepatocellular necrosis. The half‐life of AST is about 4–20 hours in the dog and one hour in the cat. In general, in acute liver injury, elevations of AST mirror those of ALT although the overall values tend not to be as high (10–30‐fold increases in the dog and up to 50‐fold in the cat). Following the insult, the AST values should return to normal more quickly than ALT reflecting its shorter half‐life. Hence, it has been suggested that persistent AST elevation in this setting is a poor prognostic sign. In cats, AST has been shown to be more sensitive than ALT for the detection of hepatobiliary diseases, with sensitivities reported to be approximately 83% and 76% sensitivity respectively in one report. Elevations in AST also occur with skeletal muscle disease so any elevation occurring in the absence of ALT elevation should be cross‐referenced with creatinine kinase measurement to exclude this possibility. In general, most biochemical profiles offer only one hepatocellular enzyme measurement, typically ALT, and the measurement of AST in addition to this is unlikely to offer a marked clinical advantage. Biliary Enzymes

Alkaline phosphatase is a membrane‐bound enzyme that has been shown to have a high sensitivity (85%) but low specificity (51%) for the detection of hepatobiliary disease in dogs. The converse is true for cats, with reported values of 48% sensitivity and 93% specificity. These performances reflect differences in the half‐lives and overall enzyme activity in the two species along with differences in the isoenzymes and their potential for induction (discussed further later). In dogs and cats, there are a variety of tissues known to exhibit ALP activity, including intestinal mucosa, renal cortex, placenta, liver, and bone. There are two genes encoding ALP in the dog with post-translation modifications resulting in different isoforms of the enzyme. The tissue nonspecific ALP gene is responsible for the transcription of liver ALP (L‐ALP), bone ALP (B‐ALP), and kidney ALP (K‐ALP), whereas the second gene encodes intestinal ALP (I‐ALP). An additional isoenzyme, glucocorticoid‐induced ALP (G‐ALP), is important in the dog and is thought to represent a post-translational modification of I‐ALP. In the dog, the total serum ALP (T‐ALP) measured on clinical biochemistry profiles reflects the combination of L‐ALP, B‐ALP, and G‐ALP; other isoenzymes fail to contribute to this measurement due to a combination of their short half‐life and/or low overall tissue activities. In the cat, T‐ALP comprises L‐ALP and B‐ALP due to a lack of G‐ALP in this species, although

placental isoenzyme has also been detected late‐term in this species. An appreciation of the contributions of these isoenzymes is important when interpreting biochemistry results. For example, B‐ALP increases with osteoblastic activity and hence its contribution to T‐ALP varies with age. In view of this, in young growing animals T‐ALP is approximately twice the normal adult reference range purely as a result of increased B‐ALP. In addition, elevations may occur in adult dogs with active bone lesions, such as osteomyelitis or osteosarcoma, but these are rarely more than five‐fold increases. Where there is concern in interpretation, it may occasionally be helpful to look at an additional enzyme, such as GGT, with no bone isoenzyme, to clarify the situation. The isoenzyme that causes far more problems for interpretation is G‐ALP, as discussed later. Liver ALP is found predominantly within the periportal zone of the liver, bound to hepatocyte canalicular and sinusoidal membranes. The serum activity of this enzyme is increased in response to cholestasis (intra‐ and extrahepatic) and induction of de novo synthesis. ALP is released into the circulation by enzyme‐induced solubilization from the hepatocyte membranes, a process that is enhanced by bile salts and hence increases with cholestasis. In the latter situation, accumulated bile salts also induce production of L‐ALP, significantly enhancing the resultant increase in serum ALP activity. The half‐life of L‐ALP is approximately 70 hours in the dog and six hours in the cat. This short half‐life in the cat, combined with a lower ALP activity in feline liver, is important practically. Clinically significant elevations of ALP activity in the cat are considerably smaller than those seen in the dog. Cats with hepatic disease may have ALP elevations two‐ or three‐fold higher than normal as opposed to dogs where values are often more than four‐ or five‐fold normal. This situation, in combination with the lack of G‐ALP enzyme in the cat, results in the far lower ­sensitivity of ALP for liver disease in the cat (48%) but enhanced specificity (93%). In view of this, the interpretation of ALP elevation in dogs and cats differs considerably. In dogs, ALP induction occurs in response to endogenous and exogenous corticosteroids and phenobarbital. The response to corticosteroids is rapid (increased ALP mRNA within 24–48 hours) and may occur following oral, topical (including ear/eye preparations) and parenteral exposure, with elevations in serum enzyme activity persisting for up to six weeks after exposure has ceased. In addition, a vacuolar hepatopathy, with glycogen accumulation within hepatocytes, occurs with exposure to excessive corticosteroid levels. This too may contribute to ALP elevation by generating intrahepatic cholestasis. The overall ALP elevations can be dramatic and complicate the interpretation of ALP activity in the dog. G‐ALP,

60  Approach to the Patient with Liver Disease

as outlined earlier, is the product of the I‐ALP gene. As with L‐ALP, the hepatocyte is the site of de novo synthesis of G‐ALP, with membrane accumulation and elution occurring in the same way. Exposure to endogenous or exogenous corticosteroids results in G‐ALP induction and hence increased serum activity. It was hoped that measurement of the distinct isotypes of ALP within a  serum sample would aid the interpretation of serum T‐ALP, but this has not been found to be the case ­clinically. While normal dogs have low G‐ALP (35 μg/mL had the highest correlation with the development of hepatotoxicity. All animals on chronic phenobarbital therapy should have a routine biochemistry panel performed every 6–12 months to monitor for the development of chronic hepatotoxicity. A bile acid tolerance test should be performed to evaluate liver function if ALT levels suddenly increase or if the serum albumin level starts to decrease. Hepatotoxicity is uncommon to occur in cats. Dosage and Monitoring  The appropriate starting dose of

phenobarbital is 2.5 mg/kg orally q12h in dogs and 2.5 mg/kg q24h in cats. An intravenous loading dose can be used to produce a rapid rise in serum blood concentration. This starting dose is the only time a weight‐based dosage is used. All future adjustments should be based on serum drug concentrations in conjunction with clinical assessment. The goal of drug level monitoring is to be proactive rather than reactive in seizure prophylaxis. Serial serum trough phenobarbital concentrations should be evaluated at 14, 60,180, and 360 days after the initiation of treatment, at six‐month intervals thereafter, if the pet has more than two seizure events between these time points, and at two weeks after a dosage change. Although blood level fluctuations may not be dramatic throughout the day in dogs with steady‐state concentrations, blood

samples are best taken in the early morning, prior to dosing, in a fasted dog, to increase consistency in comparison with published information, maintain consistency in interpretation, and remove diurnal or dietary‐induced fluctuations of absorption. Adjustments in AED dosages are undertaken either to enhance the effect or to reduce the adverse effects. The most efficacious and safe trough therapeutic phenobarbital range is 15–30 μg/mL. An optimal starting level is between 20 and 25 μg/mL. Increments of 5 μg/mL are beneficial if seizures are occurring at an equal frequency or worsening after 30 days of therapy. Cats in general can tolerate higher serum levels in the 30–45 range without complications. Adjustments of the trough phenobarbital levels can be calculated with the following formula: Desired concentration / Actual concentration total mg PB per daay Oral daily dose of PB mg Potassium Bromide

Potassium bromide can be used as either monotherapy or as an add‐on AED of choice in the dog. Concomitant potassium bromide and phenobarbital administration decreased seizure number and severity in the majority of dogs in several studies, with seizure‐free status ranging from 21% to 72% of all treated dogs. In general, many canine refractory idiopathic epileptic patients may benefit from potassium bromide. By allowing a reduction in the use of drugs metabolized by the liver, potassium ­bromide therapy may also reduce the incidence of hepatotoxicity. Bromide is not recommended for use in cats due to potential for life‐threatening allergic asthmatic reaction. Pharmacology  A starting dose of 40 mg/kg/day potas-

sium bromide is slowly metabolized in the dog, with a median elimination half‐life of 15.2 days, Steady‐state concentrations fluctuate between dogs, most likely due to individual differences in clearance and bioavailability. Dietary factors also alter serum drug concentrations, with high‐chloride diets resulting in excessive renal secretion and lower serum concentrations.

Adverse Effects  Potassium bromide is generally well tol-

erated in the dog. The most common adverse effects seen with potassium bromide and phenobarbital combination therapy are polydipsia, polyphagia, increased lethargy, and mild ataxia with increasing serum concentration. Pancreatitis and gastrointestinal intolerance have also been reported. Chronic use at higher serum concentations can result in pelvic limb ataxia and excess sedation.

70  Seizures and Movement Disorders

Administration and  Monitoring  Potassium bromide can be administered at a starting dose of 40 mg/kg/day when used as sole therapy or 30 mg/kg/day when used as an add‐on drug to phenobarbital. Potassium bromide serum concentration should be measured at one month and at the first steady‐state concentration (approximately 8–12 weeks). The recommended goal is to achieve steady‐state trough serum concentrations of 25 μg/mL for phenobarbital and 2000 mg/L for potassium bromide. The range is highly individualized according to the seizure pattern of each dog. Further reductions in phenobarbital can be attempted if a seizure‐free period is maintained for six months. The dosage is adjusted according to the formulae given below. Potassium bromide monotherapy is recommended for dogs with underlying liver disease, less frequent seizure activity (80%). However, wide fluctuations of drug metabolism occur in the dog. The oral dose should be increased when dogs are receiving phenobarbital concurrently as lower serum levels can potentially be related to the ­induction of serum hydrolases. A new extended‐release formulation of levetiracetam has been shown to have a half‐life in excess of seven hours in dogs following oral  administration, giving rise to the potential for twice‐daily administration

Adverse Effects  Levetiracetam is the best tolerated of all the current AEDs used in veterinary medicine. Sedation is the most common initial adverse effect that typically dissipates over time. Dosage and Monitoring  The initial dose of 10–20 mg/ kg orally q12h is gradually incremented to ≥20 mg/kg orally q8h. The therapeutic range is not well defined and drug monitoring is recommended only to establish the pharmacokinetic pattern of the individual patient.

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Section 8  Neurologic Disease

Drug tolerance over time, known as the “honeymoon effect,” can occur. However, wide individual patient drug metabolism may be associated with this phenomenon. As such, this author advocates incremental escalation by 10 mg/kg/dose per week when seizures persist to a maximal tolerated dose. No upper limit dosage exists due to a high therapeutic index (safety) for this  drug. A parenteral formulation is available for intravenous loading at 40–60 mg/kg over 15–30 ­minutes in a 1:1 diluted saline solution.

dependence, compared to the full agonist AED drugs like phenobarbital and diazepam. Imepitoin was recently approved for use in canine epilepsy in Europe and is undergoing clinical investigation for FDA approval in the United States.

Third‐Generation AEDs Lacosamide

Cats and dogs can develop a variety of unusual movement disorders that bewilder even the most seasoned clinicians due to the difficulty in determining their ­neuroanatomic origin and etiology. Movement disorders include a wide range of neurologic disturbances, characterized by either excess (hyperkinesia) or reduced (bradykinesia) movements. These abnormal involuntary movements (AIMs) refer to a number of muscle jerks, twitches, postures, and oscillations that have been ­classified in human neurology with official terms such as tic, chorea, tremor, dystonia, and myoclonus. Uncontrolled muscle contractions may be of muscle or neuronal origin.

This is a functionalized amino acid proven to decrease neuronal discharge frequency and synaptic excitability. The postulated mechanisms of action include selective slow inactivation of sodium channels and novel binding to collapsin response mediator protein‐2. In humans, the drug is well absorbed, has minimal first‐pass effect with predominant renal excretion, low protein binding, favorable drug–drug interactions with other AEDs, and is well tolerated. Clinical trials in humans have demonstrated a comparable decrease in seizure frequency with that of levetiracetam and zonisamide at a dose of 100– 200 mg orally q12h. A parenteral formulation is available for intravenous loading. The author has used lacosamide to successfully treat refractory idiopathic epilepsy in dogs at a dose range of 5–10 mg/kg q12h. Rufinamide

This novel drug is structurally unrelated to any other AED. Its main mechanism of action is related to prolongation of the inactive state of the sodium channel, thus preventing neuronal depolarization. In humans, the drug is absorbed slowly and has low bioavailability. Renal excretion is high and no induction of the hepatic P450 system has been found, although other hepatically metabolized drugs decrease the serum concentration. Of a total of nine double‐blinded studies in humans, five revealed a positive effect of rufinamide to treat refractory partial seizures but not generalized seizures. In beagle dogs, the mean terminal half‐life ranged between five and 14 hours. In people, dose‐dependent adverse effects include sedation, fatigue, and dizziness. Initial dosing in the dog is 20 mg/kg q12h. Serum concentration can be monitored to achieve a therapeutic range of 10–25 μg/ mL. A parenteral formulation is not currently available. Imepitoin

This drug is the first AED that acts as a low‐affinity ­partial agonist at the GABA‐A benzodiazepine receptor site. The advantage is that imepitoin is associated with fewer adverse effects of sedation and ataxia, and is less likely to develop functional drug tolerance or physical

A summary of AED therapy is shown in Table 70.1.

­Movement Disorders

History The saying “A picture is worth a thousand words” is made for movement disorders. The ability to visualize the characteristics of AIMs should be supplemented with a detailed history to characterize the nature of the movement disorder. Information regarding anatomic distribution, rhythmicity, amplitude, frequency, speed of onset and offset of the movement, relationship to posture and activity, situations that alleviate or exacerbate the movement, presence or absence during sleep, and affected ­littermates are all essential to help determine the neuroanatomic localization and potential etiology. If the movement disorder is not present at the time of the examination, owners should be encouraged to video the events for future review. Many times, the initial few minutes of observation will solidify the clinical perspective, allowing an accurate diagnostic and therapeutic course of action to proceed. Etiology and Pathophysiology The process of initiation and completion of voluntary movement is a complex one that relies on coordination of serial communications between multiple components of the central and peripheral nervous system. Failure of any of these components to complete their assigned tasks can result in either loss of the desired movement or excessive, abnormal movements. A summary of movement disorders in dogs is provided in Box 70.2.

Table 70.1  Summary of antiepileptic drug therapy in the dog Clinical pharmacology

Vd (L/ kg)

Protein binding (%)

Drug

T1/2 (h)

Tss (d)

Therapeutic range

Dosage

IV load dose

Monitor

Guidance advice

Bromide

20–46 days

100– 200

0.45

0

Monotherapy: 1000–3000 mg/L; with phenobarbital at 1500–2500 mg/L

40–60 mg/ kg orally q24h

3 % Sodium bromide 800 mg/kg/24h

At 1 and 3 months and then q6 months or q1 month post dose change

Synergistic effect with PB No hepatotoxicity Inexpensive

Clorazepate

5–6

1–2

1.6

85

20–75 μg/mL (nordiazepam)

2–4 mg/kg PO q12h

No

NR

Longer acting than diazepam Complex partial seizure therapy Inexpensive

Felbamate

5–6

1–2

1.0

25

25–100 mg/L

20 mg/kg PO q8h

No

NR

Complex partial seizure therapy Risk of blood dyscrasia Expensive

Gabapentin

2–4

1

0.2

0

4–16 mg/L

10–20 mg/ kg PO q8–12h

No

NR

For use with liver compromised patients

Levetiracetam

4–8 and 2–4 with PB

2–3

0.5

156 mEq/L (dogs) or >161 mEq/L (cats). Causes include: ●●

excess water loss: diabetes insipidus (central or nephrogenic), burns, fever, osmotic diuresis (acute/chronic renal failure, diabetes mellitus, diuretics, or IV solute administration such as mannitol, glucose or urea), osmotic diarrhea (lactulose therapy, malabsorption syndromes, infectious enteritides), hot weather

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Section 8  Neurologic Disease ●●

●●

excess salt intake: salt poisoning, administration of IV hypertonic solutions (NaCl), sodium bicarbonate, or saline emetics insufficient water intake: lack of access, inability to drink, CNS disease resulting in primary adipsia (absence of thirst), mental depression, congenital adipsia.

Marked hypernatremia may induce cerebral signs, such as depression, weakness, irritability, uncharacteristic aggression, confusion, propulsive circling, dementia, seizures, coma, and death as the result of cellular dehydration of neurons. Neurologic signs may not occur until serum Na levels exceed 170–175 mEq/L (>350 mOsm/kg), although the severity of neurologic signs depends especially on the rate of increase, with signs being most severe in animals with rapidly developing hyperosmolality. Treatment depends on the rate at which hypernatremia develops; overrapid correction can lead to worsening clinical signs (due to a relative hyponatremia and movement of water from the vascular spaces into the brain). Replacement of the water deficit should be undertaken using 5% dextrose with the aim of correction of the deficit over 24 hours (or longer). In animals with acute‐onset hypernatremia, correction of Na levels at a rate of 1.0 mEq/h is acceptable. However, if the hypernatremia is more chronic, Na levels should not be allowed to change at more than 0.5 mEq/h. In animals with excess salt intake, diuretics should be given in conjunction with fluid therapy to avoid pulmonary edema. Hyponatremia

Hyponatremia occurs when serum Na levels are less than 146 mEq/L in dogs (151 mEq/L in cats). Hyponatremia can be further divided into true hyponatremia (associated with total body water in excess of Na – serum osmolality is low, Ca++ and an efflux of potassium. This region is also closely associated with a high density of sodium channels in order to promote and amplify the signal to assure the propagation of an action potential to generate muscle contraction. As the sodium ions move into the muscle membrane, local depolarization occurs in what is termed an endplate potential (EPP) which spreads across the surface of the sarcolemma, with the ensuing excitation‐contraction coupling allowing the muscle fiber to contract. The magnitude of the EPP is dependent on the

amount of ACh released and the amount of nAChRs but there is normally an abundance of both, resulting in a large safety factor for neuromuscular transmission. Acetylcholinesterase then breaks down the ACh (producing choline and esterase) and the ACh diffuses away from the synapse, allowing the sodium channels to close. New ACh molecules are synthesized when the choline is transported, through reuptake into the presynaptic terminal. The clinical presentation in animals with neuromuscular transmission disease is very similar regardless of whether the disorder is presynaptic (i.e., botulism) or postsynaptic (i.e., myasthenia gravis). There is usually a symmetric progressive generalized weakness in all limbs. Exercise intolerance is often seen as well. Tendon reflexes may or may not be intact. There is often facial muscle, laryngeal, pharyngeal or esophageal weakness associated with these disorders. Sensory function is unaffected, as would be expected.

­Myasthenia Gravis Myasthenia gravis (MG) is a neuromuscular transmission disorder caused by a reduction in the number of functional nAChRs on the postsynaptic membrane. It is usually the result of an acquired autoimmune disease and much less commonly a congenital disorder. Clinical Presentation Myasthenia gravis should be a consideration in any animal being examined for focal or generalized neuromuscular weakness. The clinical signs may be focal in nature and limited to esophageal dilation leading to regurgitation, pharyngeal dysfunction causing dysphagia and laryngeal paresis/paralysis causing dysphonia (voice change). Multiple cranial nerve abnormalities may also occur in

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 8  Neurologic Disease

the absence of generalized muscle weakness. One study demonstrated that 43% of dogs with a confirmed diagnosis of MG (positive AChR antibody titer) did not have clinically detectable limb muscle weakness. The remaining 57% of dogs had generalized weakness, with 13% of these having no esophageal or pharyngeal dysfunction. The classic presentation of a dog with MG is generalized muscle weakness that worsens with activity but not all animals display this exercise intolerance. Dogs are more likely than humans or cats to develop megaesophagus because the canine esophagus contains more skeletal muscle than smooth muscle. If megaesophagus is present there is a higher risk of aspiration pneumonia. An acute fulminating form of MG (5 Hz) rate or incremental response at high (>50 Hz) rate. EMG may show some spontaneous activity such as fibrillation potentials and positive sharp waves but it is often normal. Treatment of botulism consists of supportive care, with mildly or moderately affected dogs recovering over a period of several days. Intensive management may be required for dogs needing ventilatory support or having megaesophagus and dysphagia as they are at high risk for

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Section 8  Neurologic Disease

aspiration pneumonia. Recumbent nursing care, which includes frequent turning, good padding to prevent decubital ulcers and bladder care, is essential for a good recovery and to avoid secondary complications. Oral antibiotics are of no benefit, unless toxico‐infectious botulism is suspected and in that case penicillins or ­metronidazole would be indicated. Administration of C. botulinum antitoxin needs to be type specific and would not be beneficial unless a specific type C‐antitoxin were available. The antitoxin administration only helps to inactivate circulating unbound toxin and not toxin already fixed at the site of neuromuscular junction damage so, depending on the timing of administration, it could be too late in the course of the disease to use. Complete recovery is possible with proper care and should occur within 1–3 weeks. Dogs with aspiration pneumonia and respiratory paralysis may have a more guarded prognosis depending on severity and the ability to obtain respiratory support. Postsynaptic Disorders Organophosphate and Carbamate Toxicity

Organophosphates (OP) and carbamates are commonly used compounds found in many pesticides and insecticides. They are long‐acting anticholinesterases that irreversibly bind acetylcholinesterase (AChE) in nervous tissue and muscle, allowing accumulation of ACh to cause continuous cholinergic stimulation. This occurs in central, muscarinic, and nicotinic cholinergic synapses, resulting in clinical signs that include autonomic overstimulation (salivation, lacrimation, urination, defecation) and neuromuscular dysfunction. A delayed OP‐induced polyneuropathy has also been described. Clinical toxicity in veterinary medicine is usually associated with inappropriate or accidental dosing, with cats being more susceptible to these compounds than dogs. Historical information is imperative to reaching the diagnosis. Although serum cholinesterase concentrations can be measured, the results are usually not available in a timely fashion so a presumptive diagnosis based on history and clinical signs is enough to begin treatment. Clinical signs of OP intoxication are usually dominated by the autonomic effects, but neuromuscular signs include a stiff rigid gait, muscle tremors, and fasciculation (except for fenthion). Treatment for these toxicities consists of reducing further exposure or absorption by bathing or gastric lavage as appropriate. Atropine sulfate can be titrated to effect to counteract the parasympathetic signs. Pralidoxime chloride (2‐PAM or protopam chloride) and pralidoxime mesylate (P2S) can be used in OP toxicity but not in carbamate toxicity (except for aldicarb). These oxime reactivators cause release the bound AChE from the OP compound.

Drug‐Induced Neuromuscular Blockade Certain drugs have been shown to induce NMJ blockade in veterinary medicine although most reports are experimental, with very few documented clinical cases. The most commonly associated antibiotics are aminoglycosides, lincomycin, penicillamine, polymyxins, and tetracyclines. The mechanism of action in aminoglycoside NMJ blockage is presynaptic inhibition of ACh release as a result of blocked calcium influx. There is also blockage of calcium influx on the postsynaptic membrane. The degree of transmission impairment with aminoglycosides depends on the specific drug and the balance between the pre‐ and postsynaptic effects. The most potent effects, in descending order, are from neomycin, kanamycin, amikacin, gentamicin, and tobramycin. Penicillamine has been documented to cause an immune‐ mediated MG in people. Polymyxins have their most profound effect on the postsynaptic membrane by acting as a noncompetitive nondepolarizing agent. Tetracyclines have the proposed effect of chelating calcium ions and thereby depressing the effect of ACh on the postsynaptic muscle fiber. It is important to remember that these drugs can potentiate neuromuscular blocking agents used during surgical procedures or may worsen or unmask preexisting disorders of neuromuscular transmission. The use of these agents should be avoided in the treatment of other neuromuscular disorders such as MG patients. Other drugs that have a reported effect on neuromuscular transmission are antiarrhythmic agents, phenothiazines, methoxyflurane, magnesium, and the antiprotozoals chloroquine and quinine. Methimazole has been documented to cause a reversible drug‐induced autoimmune MG in hyperthyroid cats. Black Widow Spider Envenomation The black widow spider (Lactrodectus spp.) has a worldwide distribution. The most potent component of its venom is the purified fraction B known as alpha‐ latrotoxin. The toxin causes a massive release of neurotransmitter and loss of synaptic vesicles. The synaptic vesicle loss is caused by increased fusion and docking of the synaptic vesicles with the plasma membrane and inhibition of new vesicle formation. The cat appears to be more susceptible to envenomation and the clinical signs usually occur within the first eight hours. There are fine muscle tremors, muscle spasticity, abdominal rigidity, pain, and profound flaccid muscle weakness. Hypertension and electrolyte abnormalities may also occur. Treatment consists of administering the appropriate antivenin and supportive care. A rapid response to the antivenin is reported in people.

74  Diseases of the Neuromuscular Junction

Snake Envenomation There are four families of snakes that produce venom which causes neuromuscular blockade: Atractaspididae (African mole viper), Colubridae (bloomsang, twig snake), Elapidae, and Viperidae. The Elapidae have three major subfamilies: Elapidae (cobras, mambas, coral snakes), Hydrophiinae (sea snakes), and Laticaudinae (sea kraits). The Viperidae have two major subfamilies: Crotalinae (pit vipers, copperheads, cotton mouths, and rattle snakes) and Viperinae, the “Old World” vipers (carpet vipers, adders, desert vipers). The pit viper toxin that causes neuromuscular weakness is the Mojave toxin. It causes a presynaptic block of ACh release which can then lead to complete neuro-

muscular blockade when there is a facilitatory release of neurotransmitter leading to complete cessation of neurotransmitter release. The Elapidae family, such as the coral snake, produce at least three neurotoxins (taipoxin, beta‐bungarotoxin, and notexin). These ­ toxins have phospholipase A2 activity and damage synaptic vesicles. There is also a change in the morphologic structure of the motor axon terminal, including a decreased number of synaptic vesicles. The clinical signs of snake envenomation are of a progressive neuromuscular weakness with cranial nerve involvement, with death usually resulting from respiratory paralysis. If the snake can be identified then the use of antivenin has a positive effect on outcome, especially the coral snake.

­Further Reading Dewey C. A Practical Guide to Canine and Feline Neurology, 2nd edn. Ames, IA: Iowa State University Press, 2008. Khorzad R, Whelan M, Sisson A, Shelton GD. Myasthenia gravis in dogs with an emphasis on treatment and

critical care management. J Vet Emerg Crit Care 2011; 21(3): 193–208. Shelton GD. Myasthenia gravis and disorders of neuromuscular transmission. Vet Clin North Am Small Anim Pract 2002; 32(1): 189–206, vii.

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75 Myopathies Marguerite F. Knipe, DVM, DACVIM (Neurology) School of Veterinary Medicine, University of California Davis, Davis, CA, USA

­Etiology/Pathophysiology

­Signalment

Myocytes are specialized cells, with contractile elements (actin and myosin) comprising the majority of the intracellular space, along with the other organelles (mitochondria, lysosomes, etc.) Disruption of myocytes (myofibers) will result in decreased ability to contract the muscle with appropriate strength. Myopathies occur secondary to either intrinsic pathology within the myofiber itself (noninflammatory myopathy) or infiltration and damage by inflammatory cells (myositis) from infectious or immune‐mediated disease. Any disruption of the muscle membrane or intracellular structures results in myofiber dysfunction, whether through accumulation of storage products, through cellular infiltration, or dysfunction of the sarcolemma, other cytoskeletal proteins, or membrane ion channels.

Inherited myopathic processes (Box  75.1) typically exhibit clinical signs in young animals (50  000  IU/L). CK is specifically released secondary to damage to the myofiber sarcolemma. Not all myopathies have elevations in CK, so a normal CK does not rule out a myopathic process; however, with appropriate clinical signs, elevated serum CK will raise the suspicion of a myopathy. Canine masticatory muscle myositis (MMM) is an immune‐mediated disease against the 2M myofibers, exclusively located in the muscles of mastication (temporalis, masseter, pterygoids, rostral digastricus). A s­ erologic assay for 2M antibodies is available through the Comparative Neuromuscular Laboratory (http://vetneuromuscular.ucsd. edu/), and is highly sensitive and specific. Functional Electromyography (EMG) performed on the anesthetized patient assesses the stability of the muscle membrane. Normal muscle maintains electrical silence on EMG, while myofibers with membrane instability will depolarize spontaneously, resulting in abnormal spontaneous activity (fibrillation potentials, positive sharp waves, complex repetitive discharges). One EMG abnormality specific for myotonia is the presence of myotonic discharges which spontaneously wax and wane, but more often than not, EMG alone is not sufficient to diagnose a myopathy. Abnormal spontaneous activity on EMG indicates either a myopathic process or denervation of that muscle, so motor nerve conduction velocities should be done to assess nerve function, as well as muscle and nerve biopsies to assess muscle histology.

75 Myopathies

Electromyography is useful to confirm that the motor unit is affected and helps the clinician determine the extent of the disease process (focal, generalized). EMG may be especially helpful to screen for subclinical disease; a common recommendation for EMG examination is to assess a minimum of two limbs, usually the thoracic and pelvic limbs on one side of the patient. Often, only one side is examined with EMG to maintain the opposite limbs free of iatrogenic mechanical injury from the exploring needle so biopsies can be obtained. Sometimes an additional limb is examined (e.g., both pelvic limbs and a thoracic limb), especially if the clinician wants to assess the contralateral limb for comparison. Histology A definitive diagnosis of a myopathy is based on histologic abnormalities of the muscle. For the clinician, biopsy selection, handling, and processing are extremely important. Prior to biopsy, clinicians must decide whether they will submit fresh muscle to a specialized laboratory for processing, or will take samples to submit in formalin for routine histopathologic evaluation. Fresh muscle must be packed appropriately and shipped overnight to the lab, so the clinician must schedule sampling and shipping so that the muscle arrives in good condition. For generalized disease processes, biopsies of the triceps, biceps femoris/quadriceps, and cranial tibial muscles permit evaluation of both proximal and distal muscles, as well as the thoracic and pelvic limbs. Temporalis muscle, if affected, is another muscle ­frequently sampled, particularly if there is a clinical suspicion of MMM (although measuring serum 2M antibody titer is less invasive  –  see Hematology/Serology earlier). If a different, specific muscle is the only one affected, then the clinician should consider sampling that as well. By sampling multiple muscles, the clinician increases the likelihood of obtaining a biopsy with pathologic changes, since only a small portion (about 1 cm3) of any muscle is taken, and focal or multifocal processes may be missed. Two laboratories process international veterinary fresh muscle samples, and information on shipping is available on their websites (see Resources below). Muscle histopathology is broadly categorized into inflammatory (true myositides) or noninflammatory (dystrophic) myopathies (see Boxes 75.1 and 75.2). Macrophages are present for myofiber phagocytosis from any cause, but true myositis is characterized by excessive cellular infiltration in the endo‐ and perimysium, with lymphocytes, neutrophils, and eosinophils, as well as macrophages. If the myositis is secondary to an infection, the causative agent may be apparent in the

biopsy, but again, the small sample makes it highly likely that microorganisms may be missed. Noninflammatory myopathies may have vacuoles, abnormal accumulations within the myofibers, loss of normal cytoskeletal architecture (cores), necrosis and phagocytosis with macrophages only, may have nonspecific abnormalities, or may be completely normal on histopathology.

­Therapy For inflammatory myopathies, specific treatment for the infectious process or immunosuppression is indicated, but for most noninflammatory myopathies, unless an underlying or concurrent treatable disease is identified that could cause the myopathy (e.g., electrolyte imbalance, endocrine disease, etc.), there are no definitive treatments. Nonspecific Therapy Nutritional supplementation with L‐carnitine (50 mg/kg PO q12h), co‐enzyme Q10 (100 mg PO q24h), and vitamin B complex for “muscle support” has been recommended for many noninflammatory myopathies, and there are only anecdotal reports of clinical improvement with these medications. Inflammatory Myopathies If an infectious etiology (parasitic, bacterial, protozoal, etc.) is identified on biopsy or suspected clinically, appropriate antimicrobial therapy should be instituted. In small animal medicine, immune‐mediated myositides are likely more common than infectious, and immunosuppressive therapy is required. In some cases, immune‐ mediated myositis may be secondary to a chronic infection. Clinicians may choose from several different immunomodulatory drugs, including glucocorticoids, azathioprine, and cyclosporine.

­Prognosis Prognosis will depend on the specific underlying myopathy, and can range from grave to excellent. Some inherited myopathies may have tolerable clinical signs and function throughout the life of the patient (e.g., some myotonias), while others progress relentlessly to markedly impair quality of life (e.g., muscular dystrophies). For most acquired myopathies, if the underlying cause is identified and appropriately treated, patients may have an excellent outcome.

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Resources http://vetneuromuscular.ucsd.edu/ – 2M antibody titers; accepts fresh muscle (and nerve) biopsies from canine and feline patients.

https://neurology.vetmed.ucdavis.edu/ndl – accepts fresh muscle (and nerve) biopsies from canine, feline, equine, and food animal patients.

Further Reading Eminaga S, Cherubini GB, Shelton GD. Centronuclear myopathy in a Border collie dog. J Small Anim Pract 2012; 53(10): 608–12. Evans J, Levesque D, Shelton GD. Canine inflammatory myopathies: a clinicopathologic review of 200 cases. J Vet Intern Med 2004; 18: 679–91. Haley AC, Platt S, Kent M, et al. Breed‐specific polymyositis in Hungarian Vizsla dogs. J Vet Intern Med 2011; 25: 393–7.

Shelton GD. What’s new in muscle and peripheral nerve diseases? Vet Comp Ortho Traumatol 2007; 20: 249–55. Sykes JE, Dubey A, Lindsay L, et al. Severe myositis associated with Sarcocystis spp. infection in 2 dogs. J Vet Intern Med 2011; 25: 1277–83.

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76 Myelopathy Joan R. Coates, DVM, MS, DACVIM (Neurology) Department of Veterinary Medicine and Surgery, College of Veterinary Medicine, University of Missouri, Columbia, MO, USA

Myelopathy is a nonspecific term representing pathologies that result in signs of spinal cord dysfunction. It is important for the clinician to recognize the neurologic signs of myelopathy and to differentiate from or avoid misdiagnosis of other mimicking orthopedic and neuromuscular (nerve, muscle, neuromuscular junction) disorders. Signalment, history, and physical and neurologic examination findings will establish presence of a myelopathy, provide neuroanatomic localization and consideration of differentials. Ultimately, with neurodiagnostics to narrow the differential diagnosis, appropriate therapies can be selected for treatment or management of patients with spinal cord disease.

­History An accurate clinical history is important in defining the cause of the myelopathy. Time of onset (sudden or insidious), rate of progression (static, gradual or rapid), and temporal relation (intermittent/episodic, stable or chronic) can be established. Acute disorders such as fibrocartilaginous embolism (FCE), aortic thromboemboli, trauma, discospondylitis, and Hansen type I intervertebral disc herniation and meningomyelitis often present with sudden onset and can be rapidly progressive (Table 76.1). Chronic disorders such as spinal canal/spinal cord neoplasia, degenerative lumbosacral syndrome, cervical spondylomyelopathy, degenerative myelopathy, and Hansen type II intervertebral disc herniation cause insidious signs of myelopathy (Table 76.2).

­Pathophysiology The spinal cord serves as a conduit for the upper motor neuronal fibers, sensory pathways and the origin of the

motor units of the cervical, thoracic, lumbar, sacral, and caudal segments. The severity of motor and sensory ­deficits from spinal cord disease is dependent upon the rapidity of disease onset, extent of lesion involvement, and the area within the vertebral column. Spinal cord injury occurs through primary mechanical and secondary injury mechanisms. Primary injury results from extrinsic or intrinsic injury processes causing laceration, compression, contusion, ischemia, and inflammation of the tissue. The acute stage consists of primary tissue effects after injury such as central hemorrhage after a severe compression/concussive injury. Typically, myelopathic changes initially involve the gray matter with centrifugal spread to the white matter. Sequential hemorrhage, edema, and neuronal necrosis depend on severity and type of injury. The primary injury initiates a cascade of vascular, ionic, and biochemical events, associated with ischemia that contributes to ­secondary spinal cord injury processes and irreversible neuronal damage. Physical attributes of the chronic stage subsequent to secondary injury mechanisms include cavitation, presence of apoptosis, and Wallerian degeneration. Neurodegenerative diseases related to underlying inherited, metabolic or toxic causes result in intrinsic neuronal degeneration involving the cell body, axon, and/or myelin.

­Clinical Signs The primary objective of the neurologic examination in spinal cord disease is to determine an accurate neuroanatomic localization (C1–5, C6–T2, T3–L3, L4–S3, and caudal segments) (Table  76.3) and lesion distribution (focal, multifocal, diffuse). Careful interpretation of the neurologic examination is essential to avoid diagnostic procedures that are inappropriate for lesion determination and potentially harmful to the patient.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 76.1  Differential diagnosis for acute‐onset myelopathy Differential category

Differentials

Comments

Degenerative

Hansen type I intervertebral disc herniation

Asymmetric or symmetric

Anomalous

Atlantoaxial instability

Usually symmetric

Neoplastic

Metastatic, vertebral body tumors, pathologic fracture, histiocytosis, lymphoma, other hematopoietic neoplasia

Often asymmetric

Inflammatory (infectious)

Bacterial, protozoal, and fungal meningomyelitis, canine distemper virus, feline infectious peritonitis, discospondylitis, spinal cord epidural empyema

Asymmetric or symmetric

Inflammatory (noninfectious)

Steroid‐responsive meningitis‐arteritis, granulomatous meningoencephalomyelitis

Usually symmetric

Trauma

Fracture/subluxation, acute noncompressive intervertebral disc herniation

Asymmetric or symmetric; nonprogressive

Vascular

Fibrocartilaginous embolism, aortic thromboembolism

Usually asymmetric; nonprogressive

Italicized differentials indicate diseases associated with hyperesthesia on spinal palpation.

Table 76.2  Differential diagnosis for chronic myelopathy Differential category

Differentials

Comments

Degenerative (spinal cord)

Degenerative myelopathy, myelinopathy, neuronopathy, axonopathy, subarachnoid diverticulum, storage disorders

Symmetric

Degenerative (vertebral column)

Hansen type II intervertebral disc herniation, cervical spondylomyelopathy (wobbler syndrome), degenerative lumbosacral syndrome, spinal synovial cyst, diffuse idiopathic skeletal hyperostosis, spondylosis deformans, mucopolysaccharidosis, osteochondromatosis

Symmetric or asymmetric

Anomalous

Chiari‐like malformation (syringohydromyelia), atlantoaxial instability, vertebral malformation, myelodysplasia, spinal stenosis, dermoid sinus

Usually symmetric

Neoplastic

Meningioma, primary spinal cord (oligodendroglioma, astrocytoma, ependymoma), nephroblastoma

Often asymmetric

Inflammatory (infectious)

Bacterial meningomyelitis, canine distemper virus, feline infectious peritonitis, protozoal, fungal

Asymmetric or symmetric

Italicized differentials indicate diseases associated with hyperesthesia on spinal palpation.

General Proprioceptive Ataxia

Paresis/Plegia

General proprioceptive (GP) ataxia reflects the lack of information reaching the central nervous system (CNS) responsible for the awareness of movement and position of neck, trunk, and limbs in space. Due to anatomic proximity within the spinal cord of the proprioceptive pathways and motor tracts, proprioceptive abnormalities often overlap with those caused by upper motor neuron (UMN) weakness. A deficit of GP appears as misplacement or knuckling of the foot, which may not occur with every step, and poorly controlled swaying of the body. The feet may be crossed or placed too far apart. Overall movement of the limbs lacks coordination.

Paresis/plegia is a deficit/loss of ability to carry out voluntary motor function. The terms lower motor neuron (LMN) and UMN are applied to differentiate two basic types of neurologic weakness. Clinical signs of UMN weakness are characterized by spasticity with normal to increased spinal reflexes and muscular hypertonia. LMN weakness manifests as flaccidity, hyporeflexia to areflexia and muscle atrophy that is severe and rapid in onset. Neuroanatomic localization to a specific spinal cord region is upon UMN or LMN signs (see Table 76.3). In addition, the affected limbs have inadequate or absent voluntary motion, which may be described as monoparesis/plegia (involvement of one limb), paraparesis/­

76 Myelopathy

Table 76.3  Neurologic signs associated with spinal cord regions Clinical signs Evaluations

C1 to C5

C6 to T2

T3 to L3

L4 to S3 and caudal segments

Mental status

Normal

Normal

Normal

Normal

Posture

Normal; wide‐based stance all limbs; +/‐ recumbency; horizontal neck carriage

Normal; wide‐based stance all limbs, recumbent; +/− horizontal neck carriage

Normal; falling; acute – Schiff– Sherrington posture,

Normal; falling; tail tone decreased or flaccid

Gait

Normal, GP/UMN ataxia (PL > TL), spasticity all limbs, tetraparesis/ plegia, hemiparesis/ plegia

Normal, spastic PL, GP/ UMN ataxia (PL > TL), shorten stride TL, tetraparesis/plegia, hemiparesis/ plegia

GP/UMN PL ataxia, symmetric or asymmetric spastic paraparesis/plegia

GP PL ataxia, weakness may vary; symmetric or asymmetric (more often with cauda equina) paraparesis/ plegia

Cranial nerves

Normal; bilateral or ipsilateral Horner syndrome

Normal; bilateral or ipsilateral Horner syndrome

Normal

Normal

Postural reactions

Normal, mild‐severe deficits (PL > TL); absent

Normal, mild‐severe deficits (PL > TL); absent

Mild‐severe deficits; absent

No deficits; mild‐severe deficits; absent

Spinal reflexes

Normal, hyperreflexia all limbs

Normal, hyporeflexia or absent reflexes TL; normal to hyperreflexia PL

Normal to hyperreflexia PL

Hypo‐ to areflexia; pseudohyperreflexic patellar reflex with sciatic nerve dysfunction

Spinal hyperesthesia

None; mild‐severe on palpation; resists neck movements

None; usually mild; may resist neck movement

None; mild‐severe on palpation

None; mild‐severe on palpation; accentuate on tail extension

Sensation (nociception)

Usually normal; severe lesions may show mild‐severe sensory loss

Usually normal; severe lesions may show mild‐severe sensory loss

Mild‐severe sensory loss; absent

None; mild‐severe (more common with intumescence lesion) sensory loss

Micturition

Usually intact; may have detrusor areflexia; sphincter hypertonia

Usually intact; may have detrusor areflexia; sphincter hypertonia

Usually affected with loss of motor function, detrusor areflexia‐sphincter hypertonia

None; mild‐severe detrusor areflexia; sphincter hypotonia

C, cervical; GP, general proprioceptive; L, lumbar; LMN, lower motor neuron; PL, pelvic limb; S, sacral; T, thoracic; TL, thoracic limb; UMN, upper motor neuron.

plegia (involvement of pelvic limbs), tetraparesis (involvement of all limbs) or hemiparesis (involvement of the ipsilateral thoracic and pelvic limbs). Gait is assessed with regard to coordination, voluntary motor functions, and direction. The action of gait is characterized by swing (flexion) and stance (extension) phases. The voluntary motor system involves recruitment of the flexor muscles to initiate the swing phase and the postural system recruits extensor muscles for the stance and propulsive phases. Animals with UMN weakness will often have a gait of long stride length, whereas with LMN weakness the stride length is shortened. The gait in animals with spinal pain may have a shortened stride length. Animals with joint, muscle, or meningeal pain often appear to be “walking on eggshells.” Thoracic or pelvic limb lameness may represent radicular pain (nerve root signature).

Postural Abnormalities Abnormal postures can be specific for vertebral column abnormalities and spinal cord dysfunction. Ventroflexion of the neck is a common phenomenon in animals with disorders of neuromuscular weakness (especially cats) or pain. Animals with spinal malformations can show kyphosis (dorsal deviation), scoliosis (lateral deviation), and lordosis (ventral deviation). A stiff neck or an arched back is often reflective of cervical or generalized spinal pain. Neck pain may manifest with horizontal neck carriage, increased muscle tone, and intermittent spasms/ jerks. The animal tends to move by turning the entire body when changing directions. When the arched back is centered to the thoracolumbar region, this may indicate pain localized to the caudal thoracic and lumbar regions. Arching of the back with low head carriage also

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can be observed in animals with neck pain. An animal with pain localized to the lumbosacral area will have its pelvic limbs tucked under the caudal abdomen. Postural Reaction Deficits

Various postural reactions (paw replacement, hopping, hemiwalking, wheelbarrow, extensor postural thrust) can further define the location and symmetry of the weakness. Asymmetric weakness is common with vascular, inflammatory, and compressive myelopathies. Spinal Reflex Abnormalities Spinal, myotatic, and withdrawal reflexes can assist with neuroanatomic localization to specific spinal regions (see Table  76.1). Presence of hypo‐ to areflexia of limb reflexes can localize the lesion to within an intumescence. The cutaneous trunci reflex can further assist with localization of a thoracolumbar spinal cord lesion. Hyperesthesia Hyperesthesia denotes an unpleasant behavioral response to a nonnoxious stimulus. As part of a routine neurologic examination, spinal hyperesthesia is evaluated by deep palpation of the spinal epaxial musculature and by detecting resistance with flexion‐extension and lateral movements. Testing for spinal hyperesthesia should be performed near the end of the neurologic examination. Spinal palpation should begin distal to the suspected lesion localization. Cervical spinal hyperesthesia can be elicited by deep palpation of the cervical spinal musculature near the vertebral transverse processes. Clinical signs include caudal flinching of the ears, twitching spinal musculature, and behavioral signs of discomfort. The neck is manipulated by flexion, extension, and lateral movements. Normal animals have full range of movement with no resistance. Resistance or behavioral reluctance to move is evidence of spinal pain. Meningeal pain often is diffuse but will commonly localize to the cervical spine. Joint and muscle pain are assessed during palpation and evaluating range of motion of the limbs. Differentials associated with spinal hyperesthesia include those associated with inflammation or compression (e.g., intervertebral disc herniation, neoplasia). Anatomic structures with nociceptors include the meninges, nerve roots, outer one‐third of the disc, joints, periosteum, and muscle. Tissue damage or inflammation produces pain through stimulation of nociceptors that are sensitive to mechanical, thermal, and chemical stimuli. Neuropathic pain occurs with injury to neural tissue and represents abnormalities in transmission and somatosensory processing in the peripheral and central nervous

s­ ystems. Some disease processes encompass both nociceptive/inflammatory and neuropathic pain mechanisms. Loss of Nociception Nociception (pain sensation) is considered the most important prognostic indicator for functional recovery of myelopathy. In dogs with thoracolumbar Hansen type I intervertebral disc herniation, the majority of those with intact nociception have an excellent prognosis. Dogs with loss of nociception longer than 24–48 hours prior to surgery have a poorer prognosis for return of ambulatory function. If surgery is performed within 12–36 hours, the prognosis is better for more rapid and complete recovery. In spinal fractures, however, animals with loss of nociception are assumed to have complete injury of neural tissues and have a poor prognosis for recovery.

­Acute Spinal Cord Dysfunction It is important to recognize specific physical and neurologic examination findings associated with acute spinal cord injury. Neurogenic shock is a systemic complication associated with severe cervical or cranial thoracic injury to the spinal cord. This syndrome results from sympathetic loss (decreased blood pressure and heart rate resulting from unopposed vagal tone) and continual vagal tone. This phenomenon results in loss of spinal cord blood flow regulation and subsequent ischemia. Neurogenic shock resolves with fluid therapy and pressor agents. Spinal shock usually manifests as flaccidity of the limbs distal to the lesion. The spinal reflexes are depressed to absent. The bladder may be flaccid with urine retention and the sphincter hypotonic. This phenomenon may mislead a neuroanatomic localization if neurologic examination is not reassessed. The duration of spinal shock is proportional to the degree of species encephalization and thus may last only a few hours in quadrupeds. Cause may be cessation of tonic input of spinal neurons by excitatory impulses in descending pathways. The Schiff–Sherrington posture is characterized by increased extensor tone of the thoracic limbs and flaccid paralysis of the pelvic limbs after acute T3–L3 spinal cord lesions. “Border cells,” which exert inhibitory influences on extensor motor neurons of the thoracic limbs via the fasciculus proprius, are predominantly located in the L2–4 spinal cord segments. Damage to these cells or interruption of the fasciculus proprius as it ascends through the thoracolumbar spinal cord causes release of thoracic limb extensor motor neurons and hypertonia. Despite the increase in extensor tone, the thoracic limbs are neurologically normal. Schiff–Sherrington posture does not indicate that the spinal cord lesion is irreversible.

76 Myelopathy

A traction injury of the spinal cord involving tethering of nerves often is traumatic in origin. This injury commonly involves the nerves associated with the sacrococcygeal and cervicothoracic spinal cord segments. Traction of nerves/nerve roots causes injury of associated spinal cord segments such as tail‐pull and brachial plexus avulsion. For example, in cats with tail‐pull injury, injury to the sacral segments affects the pelvic nerve and its innervation of the bladder, resulting in loss of bladder contraction. Ascending and descending hemorrhagic myelomalacia should be suspected in dogs with thoracolumbar intervertebral disc disease (IVDD) that have an ascending loss of the cutaneous trunci reflex. Other neurologic signs of myelomalacia include loss of nociception caudal to the lesion, ascending and descending flaccidity, weakness and areflexia, which can lead to tetraplegia, hyperthermia, and respiratory distress. Death results from asphyxia from intercostal and diaphragmatic muscle paralysis. Clinical signs of ascending and descending myelomalacia may manifest in hours to several days from onset of paraplegia.

­Diagnostic Approach Diagnostic evaluation of an animal with spinal cord disease begins with bloodwork (complete blood count, serum biochemistry), urinalysis, and survey spinal radiographs. Thoracic radiography is recommended in ­animals older than 5 years or when neoplasia is considered a disease differential. If a diagnosis cannot be determined through these routine procedures, referral to a specialty practice is recommended. Conventional radiography of the vertebral column can assist with recognition of obvious abnormalities such as congenital malformation, discospondylitis, fracture/luxation, and bone neoplasia. Meticulous technique, collimation, and proper patient positioning, usually under anesthesia or heavy sedation, are essential to detection of subtle changes that are often the key to a diagnosis. Although many lesions are obvious with cursory inspection, subtle changes may be difficult to appreciate. However, in the majority of instances, the diagnostic capability of plain radiography of the vertebral column is limited. The diagnostic capabilities are improved with myelography, injection of iodinated contrast media into the subarachnoid space. Myelography is accurate and sensitive for identifying compressive lesions. Dynamic studies (traction, extension, flexion) also can be performed. Masses that occupy space in the vertebral canal (e.g., tumors, abscesses, disc herniations) cause alterations in the myelographic contrast column. The extradural, intra-

dural/extramedullary, or intramedullary location can be determined by the type of distortion occurring in the contrast column. Cross‐sectional imaging includes computed tomography (CT) and magnetic resonance imaging (MRI) and is more likely to provide a definitive diagnosis. These imaging modalities are now available at many specialty ­practices. Lesion descriptions affecting the spinal cord are similar to those described for myelography. Administration of contrast media can improve visualization of pathology on MRI and CT. Intravenous administration of contrast media can increase the conspicuity of a lesion due to breakdown in the blood–brain barrier and better define its borders by creating greater contrast between the lesions and surrounding tissue. Computed tomography is a cross‐sectional imaging modality in which images are constructed based on the attenuation of X‐rays through tissue. The main benefit of CT over conventional radiography is that with CT, there is greater soft tissue differentiation and lack of superimposition of overlying structures which greatly improves the evaluation of the vertebral column. CT also can assist with determining lesion extent after myelography. CT images should be evaluated for alterations in attenuation of tissues. Dark or relatively black areas are hypodense or hypoattenuating while light or relatively white areas are hyperdense or hyperattenuating. CT may provide more useful information than MRI in cases of vertebral fractures or subluxation where bony detail is important. Taking advantage of the density of dystrophic mineralized intervertebral disc material, CT can be used in the evaluation of chondrodystrophic dogs with suspected IVDD. Due to the inferior soft tissue contrast, assessment of soft tissues within spinal cord tissue is limited. Magnetic resonance imaging is the preferred imaging modality for assessment of the spinal cord. The primary reason for this is the superior soft tissue contrast that MRI provides, allowing differentiation of anatomic structures. No other imaging procedure provides conspicuity of soft tissue discrimination along with exceptional resolution. Imaging can be performed in different planes (transverse, sagittal, dorsal) without loss of resolution, enabling examination of complex anatomic regions. MRI is based on the magnetic properties of hydrogen atoms, which are extremely abundant in tissues containing water, protein, and fat. Lesions are characterized based on number (single or multiple), intensity (hypo‐, hyper‐, isointense in relationship to an adjacent normal area), distribution of the intensity (homogeneous or heterogeneous), lesion borders (well or poorly defined), anatomic location of the lesion as well as lesion location in reference to the spinal cord and meninges. Similarly, lesions are characterized based on pattern of contrast enhancement.

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Cerebrospinal fluid (CSF) is contained within the subarachnoid space, which lies between the arachnoid and pia mater. Cerebrospinal fluid analysis will assist with determining the presence of inflammatory disease. Abnormalities in CSF cytology and protein are sensitive indicators of CNS disease but rarely specific for a disease process. On occasion, infectious organisms or neoplastic cells may be identified. The CSF profile, including ­protein content, cell counts and cytology, will assist the clinician to narrow the differential diagnosis. Additional diagnostic procedures include electrodiagnostic evaluation (electromyography and nerve conduction studies), CSF protein electrophoresis, serology, and exploratory surgery.

cord disorders by cross‐sectional imaging and CSF a­ nalysis combined with results for the SOD1 mutation analysis. A dog with progressive T3–L3 myelopathy that has normal neurodiagnostics and is homozygous for the SOD1 mutation is likely to have degenerative myelopathy. Degenerative conditions of the vertebral column such as intervertebral disc herniation, cervical spondylomyelopathy, and degenerative lumbosacral stenosis secondarily cause myelopathy by spinal cord compression. Other less common degenerative conditions of the ­vertebral column include synovial cysts, canal stenosis, and diffuse idiopathic skeletal hyperostosis (DISH). Anomalies

­Differential Diagnosis Myelopathies can be classified according to the DAMNIT V scheme (degenerative, anomalous, metabolic, neoplastic, nutritional, inflammatory‐infectious/noninfectious, traumatic, toxic, vascular) (see Tables 76.1 and 76.2). Degenerative Degenerative conditions of the spinal cord can occur in both young and adult animals. Myelinopathies, axonopathies, and neuronopathies present initially with progressive signs of paraparesis and eventual tetraparesis. Many of these conditions are breed specific and inherited. Recent identification of gene mutations has enhanced our understanding of the underlying pathogenesis for many degenerative conditions associated with ion channel defects, abnormalities in protein aggregation, loss of enzyme function (storage disease), and dysfunction in protein metabolism and growth factors. Canine degenerative myelopathy (DM) is an adult‐ onset neurodegenerative disease. Most dogs are at least 8 years old before onset of clinical signs. The initial signs of degenerative myelopathy typically include asymmetric GP ataxia and spastic paresis in the pelvic limbs. At this stage, segmental spinal reflexes are indicative of UMN loss. Dogs become nonambulatory paraparetic within one year after onset. When euthanasia is delayed, weakness can ascend to the thoracic limbs with emergence of LMN signs, dysphagia, and widespread muscle atrophy. Recently, a missense mutation in the gene superoxide dismutase 1 (SOD1) was established as a genetic “risk factor” for dogs developing DM. Dogs that are homozygous (two copies of the mutant allele) for the SOD1 mutation are considered at risk for developing degenerative myelopathy. Because a variety of common acquired compressive spinal cord diseases can mimic early DM, a presumptive diagnosis is made by ruling out other spinal

Dogs and cats with congenital spinal malformations often have abnormalities involving multiple segments of the vertebral column and spinal cord. Malformations associated with the spinal cord are often referred to as spinal dysraphism that involve defects in closure of the neural tube. Such defects may also involve the vertebral column. Spinal dysraphism is a result of abnormal midline fusion and disrupts the normal pattern of spinal cord circuits. Other defects include syringomyelia, dermoid sinus, spina bifida, and caudal vertebral hypoplasia. Genetic mutations and exposure to toxic and infectious agents at certain stages of development can lead to these malformations. Abnormal formation of vertebral bodies by itself does not cause spinal cord dysfunction but some conditions can cause myelopathy from spinal cord compression or spinal canal stenosis. Congenital anomalies affecting the craniocervical junction include atlantoaxial instability and atlantooccipital overlapping, which also can occur simultaneously. The subsequent spinal cord compression results in signs of C1–5 myelopathy or manifests as cervical spinal pain. Metabolic Endocrocrine‐related peripheral neuropathies such as diabetes mellitus, hypothyroidism, and insulinoma paraneoplastic neuropathy reflect as sciatic nerve disease. Clinical signs appear in the pelvic limbs first and may mimic myelopathy. Neoplastic Tumors affecting the spinal cord are described as extradural, intradural‐extramedullary, and intramedullary based on location with respect to the spinal cord. Intramedullary tumors include gliomas (astrocytoma, oligodendroglioma, and ependymoma) and neuronal‐ derived tumors. Common intradural‐extramedullary

76 Myelopathy

tumors include meningiomas, neuroepitheliomas, and nerve sheath tumors. Extradural tumors include primary bone tumors, metastatic tumors, and hematopoietic tumors (e.g., histiocytic disease, lymphoma, plasma cell tumors). Tumors that involve the vertebral body cause secondary spinal cord compression. Metastatic disease of the spinal cord can occur from spread within the CNS (drop metastasis) or from extraneural tissues via direct extension or hematogenously. Hemangiosarcoma, melanomas, and carcinomas are the most common tumors to metastasize to the spinal cord and vertebral column. Inflammatory (Infectious, Noninfectious) Meningitis (inflammation of the meninges) and meningomyelitis (inflammation of the meninges and spinal cord) cause focal or diffuse signs of myelopathy and severe spinal pain. Onset is acute, peracute or insidious. Signs typically progress but can wax and wane. Neurologic signs are variable and related to the area of spinal cord affected. Common clinical signs include GP ataxia, limb paresis, and paraspinal hyperesthesia. Lesion distribution can be focal or multifocal, causing asymmetric neurologic deficits. Animals with meningomyelitis could also have signs of encephalitis. Spinal cord MRI combined with CSF analysis is the most reliable diagnostic approach for identifying the presence and extent of CNS inflammation. Some infectious agents affect other organ systems in addition to the CNS. Serology and molecular techniques screen for infectious etiologies. Disease confirmation often requires biopsy or necropsy examination. Common infectious diseases causing myelopathy include viral (e.g., feline coronavirus, feline infectious peritonitis, canine distemper virus, feline immunodeficiency virus, feline leukemia virus), protozoal, rickettsial, algal, and fungal diseases. In the dog, canine distemper virus and protozoa are the most frequently identified agents. Infectious meningomyelitis seems to be the most common cause underlying myelopathy in the cat. Granulomatous meningoencephalomyelitis (GME) and steroid‐responsive meningitis‐arteritis are noninfectious inflammatory diseases that predominate in the dog. Inflammation/infections of the vertebral column can involve the vertebra (osteomyelitis, physitis) or intervertebral disc space (discospondylitis). Spinal cord epidural empyema is defined as an extensive accumulation of purulent material in the epidural space of the vertebral column related to direct extension of osteomyelitis or discospondylitis. Discospondylitis is associated with bacterial or fungal infection of the intervertebral disc and contiguous vertebrae. Hyperesthesia of the vertebral column is the primary clinical sign. Radiographic features of discospondylitis include lysis and sclerosis of the

vertebral endplates. Pathologic changes and lesion extent can be more conspicuous with CT and MRI. Due to its zoonotic potential, Brucella canis screening is indicated. Antimicrobial selection should be based on susceptibility testing of bacteria isolated from urine or blood or from infected tissue. Trauma Animals with spinal fractures and luxations are assessed with minimal manipulation to prevent further injury and displacement of the spine. Nociception is assessed in plegic animals to assist with determining prognosis. Spinal fracture/luxation in dogs and cats is most commonly associated with severe external trauma and results in ­spinal cord dysfunction. Diagnosis is based on radiography or cross‐sectional imaging of the entire spine. Nonsurgical or surgical management depends upon presence of instability. The prognosis for recovery from a spinal fracture/­ luxation with loss of nociception is considered poor. Vascular Occlusive Disorders Fibrocartilaginous embolic myelopathy is the most common cause of vascular occlusion of the spinal cord in dogs and infrequently can occur in cats. FCE most commonly affects the younger, larger dog breeds but also occurs in small dog breeds, with the miniature schnauzer overrepresented. Onset of signs is often peracute or acute with little progression. Key neurologic signs include asymmetric paresis with lack of paraspinal hyperesthesia. Localization frequently reflects the segments of the cervicothoracic and lumbosacral intumescences; however, segments within L4–S3 and T3–L3 are most commonly reported. The pathogenesis still remains enigmatic. Spinal cord arteries become occluded with fibrocartilage that originates from the nucleus pulposus of the intervertebral disc. The clinical presentation is sometimes difficult to distinguish from the concussive, noncompressive extrusions of the intervertebral disc. Prognosis is dependent on the severity of neurologic deficits, lesion location and extent, and owner’s commitment to nursing care. The prognosis is favorable in patients that show improvement within two weeks of onset. In cats with acute onset of asymmetric paresis/ paralysis, aortic thrombosis is a primary differential. The pelvic limbs are commonly affected, with signs of loss of femoral pulse, pain and firmness in the muscles, and loss of nociception distally. Hypertrophic cardiomyopathy is the most frequent underlying disease. Diagnosis is suspected based upon clinical signs, elevated creatine kinase concentration, and evidence of cardiac disease. Initial therapy involves management of the cardiac disease, preventing further clot

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formation and supportive care. Long‐term prognosis is guarded during the early recovery phase.

­Treatment Treatment of myelopathy is determined based on the underlying disease diagnosis. Specific treatment

approaches focus on surgical decompression of compressive lesions, excision/debulking of compressive masses (neoplasia, cyst), stabilization procedures and appropriate therapy of infectious and inflammatory diseases. Principles of appropriate supportive care and physical rehabilitation assist with expediting recovery and prevention of secondary complications of a recumbent patient.

­Further Reading Da Costa RC, Samii VF. Advanced imaging of the spine in small animals. Vet Clin North Am Small Anim Pract 2010; 40: 765–90. De Risio L, Adams V, Dennis R, McConnell FJ, Platt SR. Association of clinical and magnetic resonance imaging findings with out‐come in dogs suspected to have ischemic myelopathy: 50 cases (2000–2006). J Am Vet Med Assoc 2008; 233: 129–35. Granger N, Smith PM, Jeffery ND. Clinical findings and treatment of non‐infectious meningoencephalomyelitis in dogs: a systematic review of 457 published cases from 1962 to 2008. Vet J 2010; 184: 290–7. Griffin JF, Levine JM, Levine GJ, Fosgate GT. Meningomyelitis in dogs: a retrospective review of

28 cases (1999 to 2007). J Small Anim Pract 2008; 49: 509–17. Marioni‐Henry K. Feline spinal cord diseases. Vet Clin North Am Small Anim Pract 2010; 40: 1011–28. Pancotto TE, Rossmeisl JH, Zimmerman K, Robertson JL, Were SR. Intramedullary spinal cord neoplasia in 53 dogs (1990–2010): distribution, clinicopathologic characteristics, and clinical behavior. J Vet Intern Med 2013; 27: 1500–8. Tipold A, Stein VM. Inflammatory diseases of the spine in small animals. Vet Clin North Am Small Anim Pract 2010; 40: 871–9.

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77 Neuroophthalmology Bradford J. Holmberg, DVM, MS, PhD, DACVO Animal Eye Center, Little Falls, NJ, USA

Neuroophthalmology is a complex discipline involving the intricate relationship between the eye and the central nervous system (CNS). A thorough understanding of afferent and efferent pathways is crucial in the loca­ lization and diagnosis of neuroophthalmic disorders. A  review of normal anatomy and physiology is beyond the scope of this chapter and can be found elsewhere. Abnormalities in the afferent or efferent arm may pre­ sent as changes in vision, eye position, tear production, eyelid carriage, function, sensation, and pupil shape, size or response. This chapter will focus on common neu­ roophthalmic diseases.

­Amaurosis Amaurosis is defined as partial or complete vision loss without observable ophthalmic pathology. Clinical signs of vision loss include a reduced or absent menace response, inability to track a cotton ball, failure to navi­ gate around stationary objects, and possibly an absent dazzle reflex or pupillary light reflex (PLR). Amaurosis may be caused by primary retinal disease, such as sudden acquired retinal degeneration syndrome (SARDS), ret­ robulbar optic nerve disease, or any lesion along the optic chiasm, optic tract, lateral geniculate nucleus, optic radiation, or visual cortex. Electroretinography (ERG) is  essential in differentiating retinal disease from CNS disease as ERG will identify a retina that appears normal on examination but fails to function properly. Frequently, magnetic resonance imaging (MRI) and cer­ ebrospinal fluid (CSF) analysis are instrumental in diag­ nosing a primary CNS cause of vision loss. Neoplasia is the most common CNS cause of vision loss in dogs and cats. Prognosis for return of vision is grave with SARDS and poor with CNS disease, depending on the underlying etiology.

­Optic Neuritis Inflammation of the optic nerve is usually bilateral and results in complete vision loss with the absence of pupil­ lary light reflexes. On ophthalmic exam, the optic disc is enlarged and hyperemic with focal hemorrhages and peripapillary retinal detachment (Figure 77.1). If only the retrobulbar optic nerve is affected, the ophthalmic exam will be normal and diagnosis is confirmed through a normal ERG and abnormal MRI. T2‐weighted MRI may reveal an enlarged, moderately hyperintense optic nerve. CSF analysis in dogs with optic neuritis is variable and may reveal a marginal to marked increase in protein or cellularity. Optic neuritis can be secondary to infectious, inflam­ matory, and neoplastic disease. In dogs, infectious causes include viral (distemper and tick‐borne encephalitis virus), fungal (cryptococcosis, blastomycosis, histoplas­ mosis), protozoal (toxoplasmosis), and bacterial (ehrli­ chiosis). Granulomatous meningoencephalitis is the most common inflammatory disorder, although idio­ pathic and necrotizing meningoencephalitides have also been reported. Primary optic nerve tumors and space‐ occupying retrobulbar masses may trigger secondary nerve inflammation. In cats, optic neuritis is often sec­ ondary to feline infectious peritonitis, but may be due to other infectious diseases, including cryptococcosis and toxoplasmosis. Most cases of optic neuritis are consid­ ered idiopathic due to lack of an apparent cause even with a thorough systemic medical work‐up. Treatment of optic neuritis is directed at the underlying cause when feasible. If no underlying cause is identified, immunosuppressive doses of corticosteroids are used to reduce inflammation. Even with appropriate therapy, prognosis for return of vision is poor and optic nerve atrophy is an unfortunate common end‐result.

Clinical Small Animal Internal Medicine Volume I, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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positive with C‐type virus particles present within the ciliary nerves or ganglia. Unilateral miosis in dogs and cats can be secondary to anterior uveitis, Horner’s syndrome (discussed later), or pharmacologic agents such as pilocarpine or topical prostaglandin analogs. Anterior uveitis is differentiated from Horner’s syndrome based on concurrent ophthal­ mic clinical signs including conjunctival hyperemia, episcleral congestion, corneal edema, blepharospasm, aqueous flare, keratic precipitates, and ocular hypotony.

­Internal and External Ophthalmoplegia

Figure 77.1  Idiopathic optic neuritis in a dog. Note the optic disc is edematous and hyperemic with peripapillary retinal detachment.

­Anisocoria Pupil diameter is regulated by autonomic control of the iris sphincter and iris dilator muscles. Anisocoria is defined as unequal pupils. Physiologic anisocoria may occur with heterochromia iridis or age‐related iris atrophy with the affected pupil dilated. When not physi­ ologic, determination of which pupil is abnormal can be challenging. Examination of the patient in bright, ambient, and dim light conditions facilitates diagnosis. An abnormal mydriatic pupil will not constrict com­ pletely in bright light compared to the normal pupil. In contrast, an abnormal miotic pupil will not dilate completely in dim light compared to the normal pupil. Unilateral mydriasis in dogs and cats can have primary ocular or neurogenic causes. Ocular causes include iris atrophy, glaucoma, end‐stage retinal degeneration, optic nerve atrophy, and pharmacologic dilation with agents such as tropicamide or atropine. Neurogenic causes include a lesion in the parasympathetic nerve, oculomo­ tor nerve, or brainstem. Due to the unique innervation of the cat sphincter muscle by two short ciliary nerves con­ trolling either side of the iris, parasympathetic denerva­ tion may result in hemiplegia and a “D” shaped or reverse “D” shaped pupil. Iris sphincter hemiplegia has been associated with trauma and iris stromal infiltration by lymphoma. Mydriasis alternating between eyes is second­ ary to feline spastic pupil syndrome. Cats affected with feline spastic pupil syndrome tend to be feline leukemia

Unilateral mydriasis may be secondary to an afferent or efferent defect. Afferent defects involve the retina, optic nerve, or optic tract. Efferent defects involve the oculo­ motor nucleus, oculomotor nerve, or iris sphincter mus­ cle. Vision and PLR testing are vital in distinguishing afferent from efferent defects. Animals with afferent lesions are nonvisual, have an absent direct PLR, and an absent consensual (affected to nonaffected eye) PLR. Animals with efferent lesions are visual, have an absent direct PLR, and a normal consensual (affected to nonaf­ fected eye) PLR. The presence of a normal consensual PLR from the nonaffected eye to the affected eye rules out an efferent arm lesion. Efferent lesions may involve the oculomotor nucleus, oculomotor nerve, or iris sphincter muscle. Peripheral parasympathetic fibers along the medial aspect of the oculomotor nerve are responsible for pupillary constric­ tion. The oculomotor nerve also innervates extraocular muscles (superior, medial, and inferior rectus, inferior oblique) and the levator palpebrae superioris muscle. Lesions affecting just the peripheral aspect of the nerve will result in mydriasis only, termed internal ophthalmoplegia (Figure  77.2). Lesions affecting the entire nerve will result in lateral strabismus and ptosis, adding the diagnosis of external ophthalmoplegia. External ophthal­ moplegia is defined as paralysis of one or more of the extraocular muscles and may only involve the oculomo­ tor nerve or may also involve the trochlear and abducens nerves. When evaluating mydriasis, it is important to distin­ guish ophthalmoplegia from a pupillomotor defect. Pupillomotor defects are usually secondary to iris atro­ phy or previous topical anticholinergic medication administration. Animals with iris atrophy may have a slight direct PLR or may have a characteristic scalloped appearance to the pupillary margin. When these signs are absent, pharmacologic testing is useful in differenti­ ating a pupillomotor defect from ophthalmoplegia.

77 Neuroophthalmology

Other CNS signs may be evident depending on the spe­ cific localization of the lesion. The presence of concurrent internal and external ophthalmoplegia should prompt the clinician to evaluate for intracranial disease to explain involvement of multiple cranial nerves. Imaging (MRI) is the diagnostic tool of choice. Treatment is limited unless the underlying cause is found. Prognosis is poor as most cases progress to involve adjacent CNS sites.

­Trigeminal Nerve Deficit

Figure 77.2  Internal ophthalmoplegia in a dog. The right pupil is normal. The left pupil is mydriatic with an absent direct and consensual (right to left) PLR. Topical application of 2% pilocarpine resulted in pupil constriction within five minutes, confirming an efferent pathway deficit.

Pharmacologic testing has classically involved the use of both indirect‐ and direct‐acting parasympathomimetics. Indirect acting agents, such as topical 0.5% physostig­ mine, will cause rapid pupillary constriction in cases of a preganglionic (proximal to the cilary ganglion) lesion due to release of acetylcholine stores at the postgangli­ onic axon terminals. Direct‐acting agents, specifically 1% to 2% pilocarpine, are more commonly used. One drop of pilocarpine acts directly on the iris sphincter muscle by binding to and activating acetylcholine receptors, inducing pupil constriction within 30 min­ utes. Use of a direct‐acting agent will allow pupil con­ striction with both pre‐ and postganglionic lesions and therefore is useful in differentiating ophthalmoplegia from a pupillomotor defect but does not differentiate a pregan­ glionic from a postganglionic site. Lack of pupillary con­ striction following administration of a topical parasympathomimetic confirms a pupillomotor defect. In a young dog or one without clinical evidence of iris atrophy, pharmacologic mydriasis is commonly the cause. A thorough history will aid in diagnosis. Internal and external ophthalmoplegia occur as a result of oculomotor nerve, ganglion, or nucleus compression. They are a result of neoplasia, trauma, or inflammation at the level of the midbrain, brainstem, or cavernous sinus.

Two of the three branches of the trigeminal nerve pro­ vide sensory innervation to the eye, adnexa, and perioc­ ular skin. Lateral sensation is primarily via the maxillary branch and medial sensation via the ophthalmic branch. Corneal sensation is derived through afferent fibers within the long ciliary nerves that originate in the ophthalmic branch. Decreased or absent function is diagnosed through an absent palpebral reflex (i.e., blink response) and an absent corneal reflex. Lack of sensation must be distinguished from lack of motor function (facial nerve deficit) where the patient is unable to blink. Differentiation is accomplished by attempting to elicit a menace response or dazzle reflex while avoid­ ing contact with the skin. A patient with trigeminal nerve dysfunction will blink in response whereas animals with a facial nerve defect will not have a palpebral reflex or normal menace response. The long ciliary nerves derive from the nasociliary nerve, a branch of the ophthalmic branch of the trigemi­ nal nerve. Inadequate function of the long ciliary nerves results in corneal anesthesia or hypoesthesia. With loss of these neurons, there is a loss of normal regulatory neuromediators such as acetylcholine, substance P, calcitonin gene‐related peptide, and neuropeptide Y ­ which are instrumental in maintaining normal corneal integrity. Neurotrophic keratitis describes pathology resulting from a trigeminal nerve defect. The decrease in corneal sensation leads to reduced corneal epithelial cell proliferation and delayed healing with corneal ulcera­ tion. Neurotrophic keratitis is likely secondary to trigem­ inal nerve trauma, but as in humans, diabetes mellitus and herpes virus may be predisposing factors. The mandibular branch of the trigeminal nerve has both motor and sensory components. It is responsible for motor innervation to the masticatory muscles and sensation to the mouth, lower face, and ear. Bilateral dis­ ruption of the mandibular branch results in the inability to close the mouth. Ninety percent of cases are idiopathic and clinical signs typically resolve over three weeks. Prolonged cases may develop atrophy of the muscles of mastication. When the pterygoid muscles are affected,

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Figure 77.3  Neurotrophic and neuroparalytic keratitis in a dog. Note the left eye is enophthalmic, the third eyelid is elevated, the ocular surface is dry, and a deep, infected stromal corneal ulcer is present. Also note the severe masseter and temporalis muscle atrophy. Neuroophthalmic localization of the lesion is at the petrosal temporal bone.

secondary enophthalmos and passive third eyelid eleva­ tion may occur (Figure 77.3). Treatment for trigeminal nerve defects is supportive. Topical lubricating agents are recommended to support surface ocular health and symptomatic treatment for secondary corneal ulceration is indicated when necessary. As most cases are secondary to trauma or are idiopathic, prognosis for return of function is fair. Other reported causes of trigeminal dysfunction include neoplasia and polyneuritis of unknown origin.

­Facial Nerve Paralysis The efferent arm of the palpebral, menace, corneal, and dazzle reflex are all mediated by the facial nerve. The facial nerve arises at the facial nucleus in the brainstem and courses with the vestibulocochlear nerve through the petrosal bone before branching. Ultimately, the zygomatic branch innervates the orbicularis oculi and retractor anguli oculi muscles and controls the blink­ ing response. Clinical signs associated with facial nerve paralysis may be restricted to the eye or may also involve other structures, depending on the site affected. Nonocular clinical signs associated with facial nerve paralysis may include deviation of the nasal septum toward the unaffected side, drooping of the ipsilateral lip or pinna, and lack of sensation to the inner pinna. Ocular signs of facial nerve paralysis include an absent palpebral reflex, neuroparalytic keratitis, and, potentially, neuro­ genic dry eye. Differentiation of facial nerve paralysis from trigeminal nerve dysfunction can be difficult. Animals with facial

nerve paralysis still have normal sensation and may retract their eye and elevate the third eyelid in response to a menacing gesture or painful stimulus applied to the adnexa. Animals with a trigeminal nerve defect have a normal menace response. Without the ability to close the eyelids completely, neuroparalytic keratitis may arise due to chronic cor­ neal exposure and tear film evaporation. With progres­ sion, corneal vascularization, scarring, and ulceration ensue (see Figure 77.3). Corneal protection is the treat­ ment of choice and involves the use of topical lubri­ cants, topical antibiotics, and possibly placement of a temporary ­tarsorrhaphy. If the parasympathetic fibers that run with the facial nerve to the lacrimal gland are affected, neurogenic dry eye will occur. Approximately 20% of dogs with facial nerve paralysis develop neuro­ genic dry eye that can be diagnosed by a routine Schirmer tear test. Concurrent neurogenic dry eye hastens corneal deterioration. Treatment with oral or topical parasympathomimetics will aid in return of tear production due to denervation hypersensitivity and upregulation of acetylcholine receptors at the lac­ rimal gland. Facial nerve paralysis is reported to be idiopathic in 75% of dogs and 25% of cats. Other causes include ­surgery, trauma, neoplasia, hypothyroidism, and otitis media. Total ear canal ablation and bulla osteotomy surgery results in facial nerve paralysis or paresis in 49% of dogs and cats. Median time to restoration of function is 2–4 weeks. Neoplasia is an uncommon cause in dogs, but the most common in cats. Inflam­ mation of the middle or inner ear is often present with facial nerve paralysis. With primary or secondary pet­ rosal bone inflammation, other neuropathies may become apparent. The trigeminal, vestibulocochlear, glossopharyngeal, and oculosympathetic and parasym­ pathetic nerves course through the petrosal bone. Therefore, additional clinical signs may include ocular and facial anesthesia or hypoesthesia, neurotrophic keratitis, masticatory muscle atrophy, vestibular dis­ ease, loss of taste sensation, and Horner’s syndrome. Fifty to sixty percent of dogs and cats with facial nerve paralysis will have additional signs, with vestibular dis­ ease most common in cats and hypothyroidism or ves­ tibular disease most common in dogs. Treatment of facial nerve paralysis involves addressing the underlying cause when present. All patients with facial nerve paralysis should have a thorough otic exam and a thyroid level completed. If an underlying cause is not found, and true idiopathic facial nerve paralysis exists, supportive care to involve ocular lubrication and protection is indicated. Prognosis for return to function is guarded, with some cases improving in several weeks and others never.

77 Neuroophthalmology

­Horner’s Syndrome Sympathetic fibers to the eye originate in the paraven­ tricular nucleus of the hypothalamus, course within the spinal cord and exit between T1 and T3 into the thoracic cavity, forming the sympathetic trunk. These fibers travel through the mediastinum and join the vagosympathetic trunk in the neck before synapsing at the cranial cervical ganglion located adjacent to the tympanic bulla. All fib­ ers prior to this synapse are considered preganglionic and those that course from the cranial cervical ganglion to the eye are considered postganglionic. Any disruption of nerve conduction along this circuitous route will lead to the classic clinical signs of Horner’s syndrome. Clinical signs include ptosis secondary to loss of tone in Müller’s muscle of the upper and lower eyelid, miosis due to loss of sympathetic tone in the iris dilator muscle, and enophthalmos with secondary third eyelid elevation due to loss of smooth muscle tone of the periorbita (Figure 77.4). These four clinical signs are diagnostic for Horner’s syndrome. Depending on where nerve conduc­ tion is disrupted, these may be the only clinical signs, or additional clinical signs may be present, including altered behavior/mentation, altered gait or paresis, spinal or neck pain, dysphagia, coughing, regurgitation, otitis media, vestibular disease, keratoconjunctivitis sicca, or facial nerve paralysis. Localization of the lesion is impor­ tant for both prognosis and potential treatment. Pharmacologic testing along with identifying clinical signs are useful in differentiating preganglionic from post­ ganglionic Horner’s syndrome. Topical application of  indirect‐acting sympathomimetic drugs such as 1%  hydroxyamphetamine will induce norepinephrine release from postganglionic neurons, allowing immediate pupil dilation. This positive test confirms preganglionic Horner’s syndrome. There is no effect of an indirect‐acting

Figure 77.4  Horner’s syndrome in a cat, demonstrating ptosis of the left upper eyelid, enophthalmos, third eyelid elevation, and miosis.

sympathomimetic with cases of postganglionic Horner’s syndrome. With the difficulty in obtaining these agents and abuse potential, this testing is rarely completed. Direct‐acting sympathomimetic agents such as 0.001% epinephrine or dilute (1%) phenylephrine can be used to confirm a postganglionic lesion due to denervation hyper­ sensitivity of smooth muscle. With a postganglionic defect, application of these agents will allow improvement of the clinical signs within 5–20 minutes. Lack of a response is suggestive, but not diagnostic, of a preganglionic lesion. Preganglionic lesions carry a worse prognosis than postganglionic. The majority of preganglionic cases are secondary to neoplasia, vascular events, or significant head, neck, brachial plexus, or chest trauma. A diagnosis of preganglionic Horner’s requires further diagnostics to localize the lesion and should include a complete blood count, serum biochemistry, thyroid level, thoracic radi­ ography, MRI of brain and spinal cord, and CSF analysis. Treatment is directed toward the underlying cause. Prognosis for return of function is guarded. Postganglionic lesions tend to be idiopathic or second­ ary to otitis media, but can be secondary to retrobulbar disease or trauma. If there is no evidence of ear disease, an idiopathic cause is assumed. Golden retrievers and Labrador retrievers between the ages of 10 and 12 years are overrepresented. Most cases of idiopathic, postgan­ glionic Horner’s syndrome resolve without treatment within two months.

­Cavernous Sinus Syndrome The cavernous sinus incorporates two venous sinuses located at the base of the cranial vault extending from the orbital fissure to the petrooccipital canal. The sinuses are joined by several anastomoses across the midline. Structures of clinical significance that pass through or are in close proximity to the cavernous sinus include the oculomotor nerve, trochlear nerve, ophthalmic and maxillary branches of the trigeminal nerve, abducens nerve, and postganglionic sympathetic axons. Dogs and cats with cavernous sinus syndrome present with a multitude of clinical signs related to the affected cranial nerves, including internal and external ophthal­ moplegia (mydriasis, ptosis, lateral or ventrolateral strabismus), corneal anesthesia, periorbital and facial anesthesia, and enophthalmos. Most cases are unilateral, with rare bilateral cases reported. Mean age of affected dogs and cats is 9 and 10 years of age, respectively. Diagnosis is obtained through observation of the clinical signs. Advanced imaging, preferably MRI, is nec­ essary to confirm the diagnosis. Most reported cases in dogs occur secondary to primary or metastatic neopla­ sia. In cats, intracranial neoplasia and infectious disease

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have been reported. Long‐term prognosis is poor even with treatment aimed at the underlying cause.

­Dysautonomia (Key–Gaskell or Dilated Pupil Syndrome) Dysautonomia is a rare, idiopathic condition in which there is generalized loss of autonomic nervous system function secondary to degeneration of parasympathetic and sympathetic neuronal ganglia. Ophthalmic clinical signs may be the first observed and include bilateral mydriasis with absent PLR, elevated third eyelids, ptosis, and decreased tear production. Dysautonomia can be readily differentiated from Horner’s syndrome by the presence of mydriasis rather than miosis. Nonocular clinical signs may include xeromycteria, xerostomia, dys­ phagia, vomiting, regurgitation, gastric distension, ano­ rexia, diarrhea, bradycardia, constipation, and urinary bladder distension.

Both cats and dogs can be affected, although the disease is much more prevalent in cats. A geographic distribu­ tion has been suggested, with most affected cats living in the United Kingdom and most affected dogs living in the American Midwest. Animals tend to be young and have a rapid onset (two days to two weeks). Although there is no known cause, 50% of cats in one study did test positive for Clostridium botulinum type C. Diagnosis of dysautonomia is generally accomplished through the overt clinical signs. Pharmacologic testing as described previously for internal ophthalmoplegia and Horner’s syndrome can be completed. In cases of dysau­ tonomia, application of dilute pilocarpine will induce pupillary constriction and application of dilute epineph­ rine or phenylephrine will allow third eyelid retraction. Treatment for dysautonomia is supportive. Prognosis for survival is poor, with 75–85% of animals succumbing to the disease or being euthanized due to the difficulty in mainte­ nance of supportive care. Some cats have spontaneously recovered, although this is rare and should not be expected.

­Further Reading Harkin KR, Andrews GA, Nietfeld JC. Dysautonomia in dogs: 65 cases (1993–2000). J Am Vet Med Assoc 2002; 220(5): 633–9. Mayhew PD, Bush WW, Glass EN. Trigeminal neuropathy in dogs: a retrospective study of 29 cases (1991–2000). J Am Anim Hosp Assoc 2002; 38(3): 262–70. Nell B. Optic neuritis in dogs and cats. Vet Clin North Am Small Anim Pract 2008; 38(2): 403–12.

Theisen SK, Podell M, Schneider T, Wilkie DA, Fenner WR. A retrospective study of cavernous sinus syndrome in 4 dogs and 8 cats. J Vet Intern Med 1996; 10(2): 65–71. Webb A, Cullen C. Neuro‐ophthalmology. In: Veterinary Ophthalmology. Ames, IA: John Wiley & Sons, 2012, pp. 1821–68.

Clinical Small Animal Internal Medicine

Clinical Small Animal Internal Medicine Volume 2 Edited by David S. Bruyette, DVM, DACVIM (SAIM) Chief Medical Officer, Anivive Lifesciences, Long Beach, CA, USA

SECTION EDITORS Nick Bexfield, BVetMed, PhD, DSAM, DipECVIM‐CA, PGDipMEdSci, PGCHE, FHEA, MRCVS University of Cambridge

Johnny D. Chretin, DVM, DACVIM (O) TrueCare for Pets

Linda Kidd, DVM, PhD, DACVIM (SAIM) Western University of Health Sciences

Stephanie Kube, DVM, DACVIM (N)

Veterinary Neurology and Pain Management

Catherine Langston, DVM, DACVIM (SAIM) Ohio State University

Tina Jo Owen, DVM, DACVS Washington State University

Mark A. Oyama, DVM, MSCE, DACVIM‐Cardiology University of Pennsylvania

Nathan Peterson, DVM, DACVECC VCA West Los Angeles Animal Hospital

Lisa V. Reiter, DVM, DACVD McKeever Dermatology Clinics

Elizabeth A. Rozanski, DVM, DACVIM (SAIM), DACVECC Tufts University

Craig Ruaux, BVSc (Hons), PhD, MACVSc, DACVIM (SAIM) Massey University

Sheila M.F. Torres, DVM, MS, PhD University of Minnesota

This edition first published 2020 © 2020 by John Wiley & Sons, Inc. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions. The right of David S. Bruyette to be identified as the author of the editorial material in this work has been asserted in accordance with law. Registered Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA Editorial Office John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print‐on‐demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty The contents of this work are intended to further general scientific research, understanding, and discussion only and are not intended and should not be relied upon as recommending or promoting scientific method, diagnosis, or treatment by physicians for any particular patient. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of medicines, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each medicine, equipment, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Cataloging‐in‐Publication data applied for ISBN 9781118497067 Cover image: Cell images (left, middle) © Kateryna Kon/Shutterstock, Cell image (right) © David Bruyett, Dog and Cat image © David Bruyette Cover design by Wiley Set in 10/12pt Warnock by SPi Global, Pondicherry, India 10 9 8 7 6 5 4 3 2 1

­Dedication To CB Chastain, Dick Nelson and Ed Feldman for trying their best to make me a better clinician. For someone interested in internal medicine and endocrinology I could not have had better mentors. To all of my former students, interns and residents, thanks so much for always keeping me honest and forcing me to try to stay ahead of those I was supposed to be teaching. To all the wonderful folks at Wiley (Erica Judisch, Gillian Whitley, Purvi Patel, Holly Regan-Jones, and Katrina Maceda) that made this book possible and tried their best to keep me on track. You turned the idea into reality and improved the book at every step in the process. To my amazing children McKenna, Taylor, Ivy and Cole, thanks for letting me be a part of your lives. Being your Dad is the best job in the world. To my amazing wife and best friend Maya. You make my life complete and I look forward to sharing every day with you.

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Contents Preface  xix List of Contributors  xxi About the Companion Website  xxix VOLUME I Section 1 

Evaluation and Management of the Patient  1

  1 The Concept of One Medicine  3 Lonnie J. King   2 Statistical Interpretation for Practitioners  7 Philip H. Kass   3 Using Data for Clinical Decision Making  17 Philip H. Kass Section 2 

Endocrine Disease  27

  4 Principles of Endocrinology  29 Robert Kemppainen  5 Neuroendocrinology  35 Maya Lottati   6 Feline Acromegaly  43 David S. Bruyette   7 Pituitary‐Dependent Hyperadrenocorticism in Dogs and Cats  49 David S. Bruyette   8 Polyuria and Polydipsia  65 Jennifer L. Garcia   9 Canine Hypothyroidism  71 David S. Bruyette 10 Feline Hyperthyroidism  75 David S. Bruyette

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11 Hypoadrenocorticism in Dogs and Cats  81 Patty Lathan 12 Diabetes Mellitus in Dogs and Cats  93 Jacquie S. Rand 13 Hypoglycemia in Patients without Diabetes Mellitus: Evaluation and Management  103 Rhett Nichols 14 Canine Autoimmune Polyglandular Syndromes  113 Deborah Greco Section 3 

Cardiovascular Disease  117

15 Approach to the Patient with Suspected Cardiovascular Disease  119 Ingrid Ljungvall and Jens Häggström 16 Imaging in Cardiovascular Disease  127 Valérie Chetboul 17 Electrocardiography 165 Anna R.M. Gelzer and Marc S. Kraus 18 Pathophysiology of Heart Failure  175 Barret J. Bulmer 19 Management of Heart Failure  185 Steven Rosenthal and Mark A. Oyama 20 Ventricular Arrhythmias  199 Amara H. Estrada and Romain Pariaut 21 Supraventricular Arrhythmias  205 Romain Pariaut and Amara H. Estrada 22 Systemic Hypertension  219 Rebecca L. Stepien 23 Pulmonary Hypertension  225 Heidi B. Kellihan 24 Congenital Heart Disease  231 Brian A. Scansen 25 Valvular Heart Disease  245 Michele Borgarelli 26 Canine Myocardial Disease  253 M. Lynne O’Sullivan 27 Feline Myocardial Disease  267 Virginia Luis Fuentes

Contents

28 Pericardial Disease  275 Ashley B. Saunders and Sonya G. Gordon Section 4 

Respiratory Disease  287

29 A Respiratory Pattern‐Based Approach to Dyspnea  289 Christopher G. Byers 30 Feline Bronchial Asthma  297 Christine M. Serafin 31 Canine Chronic Bronchitis  305 Kevin Kumrow 32 Pulmonary Thromboembolism  313 Robert Goggs 33 Surgical Approaches to Thoracic Disease  325 Raegan Wells 34 Pleural Effusion  333 Ashley Allen and Gareth Buckley Section 5 

Critical Care Medicine  345

35 Approach to the Patient in the Critical Care Setting  347 Sarah Allen 36 Fluid Therapy  355 Teresa Rieser 37 Cardiopulmonary Resuscitation  365 Nathan Peterson 38 Respiratory Monitoring in Critical Care  373 Mathew Mellema 39 Acute Respiratory Failure  381 Matthew Mellema 40 Mechanical Ventilation  393 Kate Hopper and Mathew Mellema 41 Approach to the Patient with Shock  403 James W. Barr 42 Cardiogenic Shock  413 Nathan Peterson and James W. Barr 43 Septic Shock  421 James W. Barr

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44 Disorders of Heat and Cold  431 Sarah Allen 45 Acute Poisoning  437 Ben O’Kelley 46 Medical Management of Trauma and Burns  445 Nathan Peterson 47 Venomous Snake Bites  459 Nathan Peterson Section 6 

Gastrointestinal Disease  467

48 Gastrointestinal Imaging  469 Susanne M. Stieger‐Vanegas 49 Gastrointestinal Endoscopy  507 Craig Ruaux 50 Diseases of the Oral Cavity and Salivary Glands  533 Maria M. Soltero‐Rivera and Alexander M. Reiter 51 Gastritis and Gastric Ulceration in Dogs and Cats  547 Katie Tolbert and Emily Gould 52 Disorders of the Esophagus  557 Silvia Funes and Craig Ruaux 53 Motility Disorders of the Alimentary Tract  563 Reto Neiger and Silke Salavati 54 Exocrine Pancreatic Insufficiency in Dogs and Cats  583 Panagiotis G. Xenoulis 55 Pancreatitis in the Dog  591 Caroline Mansfield 56 Pancreatitis in the Cat  601 Craig Ruaux 57 Rectoanal Diseases – Medical and Surgical Management  609 Craig Ruaux and Milan Milovancev 58 Dysbiosis and the Use of Pre‐, Pro‐ and Synbiotics  621 Jan S. Suchodolski 59 Diagnosis and Management of Chronic Enteropathies  627 Karin Allenspach Section 7 

Diseases of the Liver, Gallbladder, and Bile Ducts  639

60 Approach to the Patient with Liver Disease  641 Emma O’Neill

Contents

61 Imaging in Hepatobiliary Disease  659 Esther Barrett 62 Metabolic, Toxic, and Neoplastic Diseases of the Liver  677 Jan Rothuizen 63 Feline Inflammatory Liver Disease  687 Nicki Reed 64 Canine Inflammatory Liver Disease  695 Nick Bexfield 65 Cirrhosis and its Consequences  705 Katherine Scott 66 Portosystemic Shunts and Microvascular Dysplasia  713 Geraldine Hunt 67 Diseases of the Gallbladder and Extrahepatic Biliary Ducts  721 Ben Harris Section 8 

Neurologic Disease  727

68 The Neurologic Examination  729 Alexander de Lahunta 69 Central Nervous System Trauma  741 Simon R. Platt 70 Seizures and Movement Disorders  759 Michael Podell 71 Disorders of the Forebrain  773 Sam N. Long 72 Vestibular Disease  789 Tammy Stevenson 73 Meningoencephalitis and Meningomyelitis  795 Christopher L. Mariani 74 Diseases of the Neuromuscular Junction  803 David Lipsitz 75 Myopathies 811 Marguerite F. Knipe 76 Myelopathy 815 Joan R. Coates 77 Neuroophthalmology 823 Bradford J. Holmberg

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VOLUME II Section 9  Part 1 

Infectious Disease  829

Diagnostic Considerations  831

78 Epidemiology of Infectious Disease  833 Peggy L. Schmidt and Helen T. Engelke 79 Laboratory Diagnosis of Infectious Diseases  839 Laia Solano‐Gallego and Gad Baneth Part 2 

Select Infectious Diseases and Disease Agents  849

80 Canine Distemper  851 David S. Bruyette 81 Canine Herpesvirus  855 Yvonne Drechsler 82 Canine Viral Enteritis  857 Margaret C. Barr 83 Viral Papillomatosis  861 Margaret C. Barr 84 Canine Influenza Virus  865 Ellen Collisson 85 Feline Parvovirus  869 Margaret C. Barr 86 Feline Coronavirus  873 Yvonne Drechsler 87 Feline Leukemia Virus  877 David S. Bruyette 88 Feline Immunodeficiency Virus  883 Tom Phillips 89 Feline Viral Upper Respiratory Tract Disease  887 Yvonne Drechsler 90 Rabies in Dogs and Cats  891 Emily Beeler and Karen Ehnert 91 West Nile Virus  899 Tracey McNamara 92 Ebola Virus  901 Linda Kidd

Contents

  93 Ehrlichiosis and Anaplasmosis  903 Pedro P. Vissotto de Paiva Diniz   94 Salmon Poisoning Disease  913 Pedro P. Vissotto de Paiva Diniz  95 Wolbachia pipientis Infection  917 Pedro P. Vissotto de Paiva Diniz  96 Bartonellosis  919 Pedro P. Vissotto de Paiva Diniz  97 Hemotropic Mycoplasma  927 Séverine Tasker  98 Nonhemotropic Mycoplasma, Ureaplasma, and L‐Form Bacteria  931 Joachim Spergser   99 Spotted Fever and Typhus Group Rickettsia  937 Linda Kidd 100 Lyme Borreliosis  941 Meryl P. Littman 101 Leptospirosis 945 Katharine F. Lunn 102 Yersiniosis 951 Maria Grazia Pennisi 103 Tularemia 955 Linda Kidd 104 Q Fever  959 Linda Kidd 105 Brucellosis 963 Erin E. Runcan and Marco A. Coutinho da Silva 106 Tetanus and Botulism  967 Andrea Fischer 107 Anthrax 971 Wayne E. Wingfield and Jerry J. Upp 108 Actinomycosis, Nocardiosis, and Mycobacterial Infections  977 Joanna Whitney and Vanessa R. Barrs 109 Fungal Infections  985 Jane E. Sykes 110 Protozoal and Protozoa‐Like Infections  1003 Gad Baneth and Laia Solano‐Gallego

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111 Coccidia 1023 Chris Adolph 112 Surgical, Traumatic, and Bite Wound Infections  1029 Laura L. Nelson 113 Canine Infectious Respiratory Disease Complex  1035 Jonathan Dear Part 3 

Therapeutic Considerations  1039

114 Antimicrobial Therapy in Dogs and Cats  1041 Katrina R. Viviano 115 Antifungal Therapy  1049 Daniel S. Foy Part 4 

Special Topics  1055

116 Nosocomial and Multidrug‐Resistant Infections  1057 Jason W. Stull and J. Scott Weese 117 Management of Infectious Disease in Kennels and Multicat Environments: Creating a Culture of Compliance  1063 Frank Bossong Section 10 

Renal and Genitourinary Disease  1067

118 Disorders of Sodium and Water Homeostasis  1069 Julien Guillaumin and Stephen DiBartola 119 Disorders of Phosphorus and Magnesium  1079 Rosanne Jepson 120 Acute Kidney Injury  1089 Adam E. Eatroff 121 Glomerular Disease  1101 Shelly L. Vaden 122 Obstructive Uropathy  1109 Edward Cooper and Brian A. Scansen 123 Urolithiasis in Small Animals  1123 Alice Defarges, Michelle Evason, Marilyn Dunn, and Allyson Berent 124 Prostatic Diseases  1157 Serge Chalhoub 125 Management of Chronic Kidney Disease  1165 Jessica Quimby

Contents

126 The Role of Dialysis  1175 Adam E. Eatroff 127 Micturition and Associated Disorders  1181 Julie K. Byron 128 Urinary Tract Infections  1189 Nicole Smee Section 11 

Oncologic Disease  1197

129 Approach to the Cancer Patient  1199 Lisa DiBernardi 130 Biology of Cancer and Cancer Genetics  1205 Mary‐Keara Boss 131 Endocrine Manifestations of Cancer: Ectopic Hormone Production  1213 Cory Brown 132 Paraneoplastic Syndromes  1217 Cory Brown 133 Lymphoid Leukemias, Myeloid Neoplasia, and Myelodysplastic Syndrome  1223 Angela R. Kozicki 134 Lymphomas 1231 Kristine Elaine Burgess 135 Plasma Cell Disorders  1241 Orna Kristal 136 Central Nervous System Tumors in Dogs and Cats  1247 David Ruslander 137 Cancer of the Nose and Mouth  1253 Lauren Askin Quarterman 138 Tumors of the Eye and Ocular Adnexa  1261 Erin M. Scott and Paul E. Miller 139 Cancer of the Heart  1271 Nick A. Schroeder and Lisa DiBernardi 140 Cancer of the Airway and Lung  1277 Joanne L. Intile 141 Cancer of the Esophagus and Stomach  1283 Avenelle I. Turner 142 Cancer of the Small and Large Intestine  1287 Edwin Brodsky

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143 Exocrine Pancreatic Cancer  1293 Avenelle I. Turner 144 Pancreatic Endocrine Tumors  1297 Karen Eiler 145 Liver and Biliary Tract Tumors  1303 Amanda K. Elpiner 146 Tumors of the Urinary Tract  1307 Pedro A. Boria 147 Tumors of the Male Reproductive System  1311 Trina Hazzah 148 Gynecologic Cancers  1317 Trina Hazzah 149 Mammary Cancer  1321 Julie Bulman‐Fleming 150 Tumors of Bone and Joint  1327 Stephanie L. Shaver, William T.N. Culp, and Robert B. Rebhun 151 Soft Tissue Sarcomas  1333 Lauren Askin Quarterman 152 Hemangiosarcoma 1339 Christine B. Oakley and John D. Chretin 153 Melanoma 1347 Philip J. Bergman 154 Nonmelanoma Skin Cancers  1353 Barbara E. Kitchell 155 Mast Cell Neoplasia  1359 Zachary M. Wright 156 Apheresis in Companion Animals  1369 Steven Suter Section 12 

Skin and Ear Diseases  1373

157 Approach to the Patient with Dermatologic Disease  1375 Lisa V. Reiter 158 Principles of Therapy of Dermatologic Diseases  1397 Sandra N. Koch 159 Atopic Dermatitis  1403 Alison Diesel

Contents

160 Allergic Skin Diseases  1413 Patrick Hensel 161 Cutaneous Adverse Food Reactions  1419 Ralf S. Mueller 162 Autoimmune and Immune‐Mediated Skin Diseases  1423 Nicole A. Heinrich 163 Approach to Alopecia  1433 Linda A. Frank 164 Canine Sterile Papular and Nodular Skin Diseases  1441 Sandra Diaz 165 Parasitic Skin Diseases  1449 Elizabeth E. Toops 166 Bacterial Pyodermas  1461 Jennifer R. Schissler 167 Otitis 1471 Sue Paterson 168 Cutaneous Manifestations of Systemic Disease  1481 Kinga Gortel 169 Superficial Necrolytic Dermatitis  1491 Mitchell D. Song 170 Miscellaneous Skin Diseases  1495 Lisa V. Reiter and Sheila M.F. Torres Section 13 

Diseases of Bone and Joint  1507

171 Skeletal Development and Homeostasis  1509 Matthew J. Allen and Gert J. Breur 172 Metabolic Bone Diseases  1521 Keren E. Dittmer 173 Osteoarthritis in Small Animals  1529 Steven A. Martinez 174 Developmental Orthopedic Diseases  1537 Gert J. Breur, Nicolaas E. Lambrechts, and Heather A. Towle Millard Section 14 

Social and Ethical Issues in Veterinary Medicine  1553

175 Canine and Feline End of Life Care  1555 Robin Downing

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Section 15 

Preventive Care  1565

176 Role of Immunization  1567 Melissa Kennedy 177 Behavior Triage for Internists and the General Practitioner  1571 Karen Lynn C. Sueda Section 16  Index  1583

Laboratory Support  1581

xix

Preface Internal medicine is hard. Progress in science and technology makes it almost impossible to keep abreast of recent advances and on top of that, all of us want to balance our life both at work and home. Is there enough time in the day for everything? There are a variety of ways in which we learn and the path is not the same for everyone. For those of us with an interest in internal medicine, and I’m assuming that includes the reader of this book, we rely on a number of resources. Our colleagues, continuing education seminars, the literature, professional educational networks, and reference textbooks. A concern of mine over the past few years has been the increasing reliance on technology to arrive at a clinical diagnosis rather than emphasizing the need to understand physiology and the value of a complete history and a thorough physical examination. We feel increasing pressure to arrive at a specific and definitive diagnosis and, more importantly, to arrive at that diagnosis almost immediately. Your goal in internal medicine is not to arrive at a diagnosis. I would much prefer to see a pet respond to my treatment and recover without a definitive diagnosis than to arrive at a definitive diagnosis at necropsy. If you are successful in achieving a diagnosis, that can be very rewarding. However, your goal should be to accurately identify problems and address those problems in a logical, timely, and cost‐effective manner, always weighing the risk/benefit of running myriad diagnostics versus improving the quality of life of your patient and the pet owner. While not every pet owner will be able to afford or desire to follow each and every one of our recommendations, it is our job to make sure that whatever decision the owner makes is based on being fully informed. Illness is really physiology gone awry. If you have an understanding of what is normal, it makes your job of identifying the abnormal much easier. The body has a limited repertoire of responses to an insult so often many diseases will have very similar clinical presentations.

While we all like to make lists of the 20 differential diagnoses for a given patient’s abnormalities, it’s important to recognize that in clinical practice, common things occur commonly and those differentials should be at the top of your rule‐out list. There are numerous excellent textbooks on the market and I use many of them on a daily/weekly basis. Some serve as definitive reference works for the topic under discussion. Some are brief, bullet point, clinically oriented texts that one can use to find something quickly. What I thought was currently lacking was a text that would be continually updated, provide enough background physiology to help the reader understand normal versus abnormal, and provide useful and, more importantly, clinically relevant material to help you with your patients. The goal of this text was to first identify section editors who were recognized experts in their field both academically and clinically. The section editors then identified topics they felt were of greatest clinical importance and selected authors who could translate this information into a text that would be used every day. I hope we have achieved that goal. We will be updating the text online with new information and updated references on a quarterly basis, adding additional sections and chapters with future editions, and uploading podcasts consisting of interviews with the authors to highlight and emphasize the material in the text and any recent advances in the field. We hope that you will find the text helpful and all credit for that success goes to the authors. Any omissions or errors lie with me so please let me know both the good and the bad so we can improve things going forward. As my favorite philosopher once said “It’s a magical world, Hobbes, ol’ buddy, let’s go exploring.” All the best and enjoy exploring the mysteries of medicine. Dave

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List of Contributors Chris Adolph, DVM, MS, PhD, DACVM (Parasitology)

Zoetis Inc. Tulsa, OK, USA Ashley Allen, DVM, DACVECC

College of Veterinary Medicine University of Florida Gainesville, FL, USA Matthew J. Allen, VetMB, PhD

Department of Veterinary Medicine University of Cambridge Cambridge, UK Sarah Allen, DVM, DACVECC

Massachusetts Veterinary Referral Hospital Woburn, MA, USA Karin Allenspach, Dr.Med.Vet., PhD, DECVIM‐CA

College of Veterinary Medicine Iowa State University Ames, IA, USA Gad Baneth, DVM, PhD, DECVCP

School of Veterinary Medicine Hebrew University Rehovot, Israel James W. Barr, DVM, DACVECC

BluePearl Veterinary Partners Tampa, FL, USA Margaret C. Barr, DVM, PhD

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Vanessa R. Barrs, BVSc (Hons), MVetClinStud, MACVSc (Small Animal), FACVSc (Feline), GradCertEd (Higher Ed)

University of Sydney Sydney, Australia Emily Beeler, DVM, MPH

Veterinary Public Health Program Los Angeles County Department of Public Health Los Angeles, CA, USA Allyson Berent, DVM, DACVIM (SAIM)

The Animal Medical Center New York, USA Philip J. Bergman, DVM, PhD, DACVIM (Oncology)

Katonah‐Bedford Veterinary Center Clinical Studies, VCA Antech Bedford Hills, NY, USA Nick Bexfield, BVetMed, PhD, DSAM, DECVIM‐CA, PGDipMEdSci, FHEA, MRCVS

The Queen’s Veterinary School Hospital University of Cambridge Cambridge, UK Michele Borgarelli, DVM, PhD, DECVIM (Cardiology)

Virginia‐Maryland Regional College of Veterinary Medicine Blacksburg, VA, USA Pedro A. Boria, DVM, MS, DACVIM (Oncology)

Blue Pearl Veterinary Partners Northfield, IL, USA Mary‐Keara Boss, DVM, PhD, DACVR (Radiation Oncology)

Colorado State University Fort Collins, CO, USA Frank Bossong, DVM

Esther Barrett, MA, VetMB, DVDI, DECVDI, MRCVSt

Wales and West Imaging Chepstow, UK

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

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­List of Contributors

Gert J. Breur, DVM, MS, PhD, DACVS

Johnny D. Chretin, DVM, DACVIM (Oncology)

College of Veterinary Medicine Purdue University West Lafayette, IN, USA

TrueCare for Pets Studio City, CA, USA

Edwin Brodsky, DVM, DACVIM (Oncology)

College of Veterinary Medicine University of Missouri Columbia, MO, USA

Veterinary Medical Center of Long Island West Islip, NY, USA Cory Brown, DVM, DACVIM (SAIM)

Joan R. Coates, DVM, MS, DACVIM (Neurology)

Ellen Collisson, MS, PhD

VetScan Mobile Diagnostics Powell, OH, USA

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

David S. Bruyette, DVM, DACVIM (SAIM)

Edward Cooper, VMD, MS, DACVECC

Anivive Lifesciences Long Beach, CA, USA Gareth Buckley, MA, VetMB, MRCVS, DACVECC

College of Veterinary Medicine University of Florida Gainesville, FL, USA Julie Bulman‐Fleming, DVM, DACVIM (Oncology)

Veterinary Cancer Group Tustin, CA, USA Barret J. Bulmer, DVM, MS, DACVIM (Cardiology)

Tufts Veterinary Emergency Treatment & Specialties Walpole, MA, USA Kristine Elaine Burgess, DVM, DACVIM (Oncology)

Cummings School of Veterinary Medicine Tufts University North Grafton, MA, USA Christopher G. Byers, DVM, DACVECC, DACVIM (SAIM), CVJ

CriticalCareDVM.com Omaha, NE, USA Julie K. Byron, DVM, MS, DACVIM (SAIM)

College of Veterinary Medicine Ohio State University Columbus, OH, USA

College of Veterinary Medicine Ohio State University Columbus, OH, USA William T.N. Culp, VMD, DACVS, ACVS Founding Fellow of Surgical Oncology

School of Veterinary Medicine University of California, Davis Davis, CA, USA Jonathan Dear, DVM, DACVIM (SAIM)

School of Veterinary Medicine University of California, Davis Davis, CA, USA Alice Defarges, DVM, DACVIM (SAIM)

Ontario Veterinary College University of Guelph Guelph, ON, Canada Alexander de Lahunta, DVM, PhD, DACVIM (Neurology), DACVP

College of Veterinary Medicine Cornell University Ithaca, NY, USA Sandra Diaz, DVM, MS, DACVD Department of Veterinary Clinical Sciences

Ohio State University Columbus, OH, USA

Serge Chalhoub, DVM, DACVIM (SAIM)

Stephen DiBartola, DVM, DACVIM (SAIM)

Faculty of Veterinary Medicine University of Calgary Calgary, AB, Canada

Ohio State University Columbus, OH, USA

Valérie Chetboul, DVM, PhD, DECVIM‐CA (Cardiology)

Lisa DiBernardi, DVM, DACVIM (Oncology), DACVR (Radiation Oncology)

National Veterinary School at Alfort Maisons‐Alfort, France

Gulf Coast Veterinary Specialists Houston, TX, USA

­List of Contributors

Alison Diesel, DVM, DACVD

Michelle Evason, DVM, DACVIM (SAIM)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Atlantic Veterinary College University of Prince Edward Island Charlottetown, PE, Canada

Pedro P. Vissotto de Paiva Diniz, DVM, PhD

Andrea Fischer, DVM, DECVN, DACVIM (Neurology)

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Keren E. Dittmer, PhD, BVSc, DACVP

School of Veterinary Science Massey University Palmerston North, New Zealand Robin Downing, DVM, MS, DAAPM, DACVSMR

The Downing Center for Animal Pain Management, LLC Windsor, CO, USA Yvonne Drechsler, PhD

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Marilyn Dunn, DVM, DACVIM (SAIM)

Centre Hospitalier Universitaire Vétérinaire University of Montreal Saint‐Hyacinthe, QC, Canada Adam E. Eatroff, DVM, DACVIM (SAIM)

ACCESS Specialty Animal Hospitals Culver City, CA, USA Karen Ehnert, DVM, MPVM, DACVPM

Veterinary Public Health Program Los Angeles County Department of Public Health Los Angeles, CA, USA Karen Eiler, DVM, MS, DACVIM (SAIM)

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Helen T. Engelke, BVSc, MPVM, DACVPM, MRCVS

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Ludwig‐Maximilians University Munich, Germany Daniel S. Foy, MS, DVM, DACVIM (SAIM), DACVECC

College of Veterinary Medicine Midwestern University Glendale, AZ, USA Linda A. Frank, DVM, MS, DACVD

College of Veterinary Medicine University of Tennessee Institute of Agriculture Knoxville, TN, USA Virginia Luis Fuentes, MA, VetMB, PhD, CertVR, DVC, MRCVS, DACVIM (Cardiology), DECVIM (Cardiology)

Royal Veterinary College University of London Hatfield, Herts, UK Silvia Funes, DVM, MS, DACVIM (SAIM)

VCA Bay Area Veterinary Specialists and Emergency Hospital San Leandro, CA, USA Jennifer L. Garcia, DVM, DACVIM (SAIM)

Sugar Land Veterinary Specialists and VetCompanion Houston, TX, USA Anna R. M. Gelzer, DVM, PhD, DACVIM (Cardiology), DECVIM‐CA (Cardiology)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA Robert Goggs, BVSc, PhD, DACVECC, DECVECC

Cornell University College of Veterinary Medicine Companion Animal Hospital Ithaca, NY, USA Sonya G. Gordon, DVM, DVSc, DACVIM (Cardiology)

VCA Great Lakes Veterinary Specialists Cleveland, OH, USA

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Amara H. Estrada, DVM, DACVIM (Cardiology)

Kinga Gortel, DVM, MS, DACVD

University of Florida Gainesville, FL, USA

Tri Lake Animal Hospital and Referral Centre Lake Country, BC, Canada

Amanda K. Elpiner, DVM, DACVIM (Oncology)

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Emily Gould, DVM, MS

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Deborah Greco, DVM, PhD, DACVIM (SAIM)

Desert Veterinary Specialists Palm Desert, CA, USA Julien Guillaumin, DVM, DACVECC

Colorado State University Fort Collins, CO, USA Jens Häggström, DVM, PhD, DECVIM‐CA (Cardiology)

Swedish University of Agricultural Sciences Uppsala, Sweden Ben Harris, MA, VetMB, CertSAM, MRCVS

Northwest Veterinary Specialists Sutton Weaver, Cheshire, UK

Rosanne Jepson, BVSc, MVetMed, PhD, DACVIM (SAIM), DECVIM-CA, PGCertVetEd, FHEA, MRCVS

Royal Veterinary College University of London London, UK Philip H. Kass, BS, DVM, MPVM, MS, PhD

Population Health and Reproduction Davis, CA, USA Heidi B. Kellihan, DVM, DACVIM (Cardiology)

University of Wisconsin Madison, WI, USA Robert Kemppainen, DVM, PhD

Auburn University Auburn, AL, USA Melissa Kennedy, DVM, PhD, DACVM

College of Veterinary Medicine University of Tennessee Knoxville, TN, USA Linda Kidd, DVM, PhD, DACVIM

Trina Hazzah, DVM, DACVIM (Oncology)

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Nicole A. Heinrich, DVM, DACVD

McKeever Dermatology Clinics Eden Prairie, MN, USA Patrick Hensel, Dr. Med.Vet., DACVD, DECVD

Tierdermatologie Basel, Switzerland Bradford J. Holmberg, DVM, MS, PhD, DACVO

Animal Eye Center Little Falls, NJ, USA Kate Hopper, BVSc, DACVECC, PhD

School of Veterinary Medicine University of California, Davis Davis, CA, USA Geraldine Hunt, BVSc, MVetClinStud, PhD, FACVSc

School of Veterinary Medicine University of California, Davis Davis, CA, USA Joanne L. Intile, DVM, DACVIM (Oncology) College of Veterinary Medicine

North Carolina State University Raleigh, NC, USA

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Lonnie J. King, DVM, MS, MPA, DACVPM

The Ohio State University Columbus, OH, USA Barbara E. Kitchell, DVM, PhD, DACVIM (Internal Medicine, Oncology)

VCA Veterinary Care Animal Hospital and Referral Center Albuquerque, NM, USA Marguerite F. Knipe, DVM, DACVIM (Neurology)

School of Veterinary Medicine University of California, Davis Davis, CA, USA Sandra N. Koch, DVM, MS, DACVD

College of Veterinary Medicine University of Minnesota St Paul, MN, USA Angela R. Kozicki, DVM, DACVIM (Oncology)

Bluepearl Veterinary Partners Southfield, MI, USA Marc S. Kraus, DVM, DACVIM (SAIM, Cardiology), DECVIM‐CA (Cardiology)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA

­List of Contributors

Orna Kristal, DVM, DACVIM (Oncology)

Veterinary Specialty and Emergency Hospital Wan Chai, Hong Kong Stephanie Kube, DVM, DACVIM (Neurology)

Veterinary Neurology and Pain Management Center of New England Walpole, MA, USA Kevin Kumrow, DVM, DACVIM (SAIM)

Orchard Park Veterinary Medical Center Orchard Park, NY, USA Nicolaas E. Lambrechts, DVM, DECVS, DACVSMR

College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO, USA Catherine Langston, VM, DACVIM (SAIM)

Department of Veterinary Clinical Sciences Ohio State University Columbus, OH, USA Patty Lathan, VMD, DACVIM (SAIM)

College of Veterinary Medicine Mississippi State University Mississippi State, MS, USA David Lipsitz, DVM, DACVIM (Neurology)

Veterinary Specialty Hospital San Diego, CA, USA Meryl P. Littman, VMD, DACVIM (SAIM)

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA Ingrid Ljungvall, DVM, PhD, DECVIM‐CA (Cardiology)

Swedish University of Agricultural Sciences Uppsala, Sweden Sam N. Long, BVSc, PhD, DipECVN

Centre for Animal Referral and Emergency Melbourne, Australia Maya Lottati, DVM, PhD, DACVIM (SAIM)

Caroline Mansfield, BSc, BVMS, MVM, PhD, MANZCVS, DECVIM-CA

Melbourne Veterinary School University of Melbourne Melbourne, Australia Christopher L. Mariani, DVM, PhD, DACVIM (Neurology)

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA Steven A. Martinez, DVM, MS, DACVS, DACVSMR

College of Veterinary Medicine Washington State University Pullman, WA, USA Tracey McNamara, DVM, DACVP

College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA Mathew Mellema, DVM, PhD, DACVECC

School of Veterinary Medicine University of California, Davis Davis, CA, USA Heather A. Towle Millard, DVM, MS, DACVS‐SA

Blue Pearl Pet Hospital Overland Park, KS, USA Paul E. Miller, DVM, DACVO

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA Milan Milovancev, DVM, DACVS‐SA

School of Veterinary Medicine Oregon State University Corvallis, OR, USA Ralf S. Mueller, Dr. Med.Vet., DACVD, FANZCVSc

Centre for Clinical Veterinary Medicine Ludwig‐Maximilians‐University of Munich Munich, Germany Reto Neiger, Dr.Med.Vet., PhD, DACVIM (SAIM), DECVIM-CA

TrueCare for Pets Studio City, CA, USA

Small Animal Clinic Hofheim Hofheim, Germany

Katharine F. Lunn, BVMS, MS, PhD, MRCVS, DACVIM (SAIM)

Laura L. Nelson, DVM, MS, DACVS

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA

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­List of Contributors

Rhett Nichols, DVM, DACVIM (SAIM)

Antech Diagnostics Farmingdale, NY, USA Christine B. Oakley, DVM

VCA Veterinary Specialists of the Valley Woodland Hills, CA, USA Ben O’Kelley, DVM, DACVECC

BluePearl Veterinary Partners Tampa, FL, USA Emma O’Neill, BSc, BVSc, PhD, DSAM, DECVIM‐CA, MRCVS

School of Veterinary Medicine University College Dublin Dublin, Eire M. Lynne O’Sullivan, DVM, DVSc, DACVIM (Cardiology)

Atlantic Veterinary College University of Prince Edward Island Charlottetown, PEI, Canada Tina Jo Owen, DVM, DACVS

Washington State University Pullman, WA, USA Mark A. Oyama, DVM, MSCE, DACVIM (Cardiology) Department of Clinical Sciences and Advanced Medicine

University of Pennsylvania Philadelphia, PA, USA

Romain Pariaut, DVM, DACVIM (Cardiology), DECVIM-CA (Cardiology)

College of Veterinary Medicine Cornell University Ithaca, NY, USA Sue Paterson, MA, VetMB, DVD, DECVD, FRCVS

Veterinary Dermatologicals Altrincham, Cheshire, UK Maria Grazia Pennisi, DVM, PhD

Specialist Applied Microbiology University of Messina Messina, Italy Nathan Peterson, DVM, DACVECC

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Tom Phillips, DVM, MS, PhD

Deceased Formerly College of Veterinary Medicine Western University of Health Sciences Pomona, CA, USA

Simon R. Platt, BVM&S, FRCVS, DACVIM (Neurology), DECVN

College of Veterinary Medicine University of Georgia Athens, GA, USA Michael Podell, MSc, DVM, DACVIM (Neurology)

Chicago Veterinary Emergency and Specialty Center Chicago, IL, USA Lauren Askin Quarterman, DVM, DACVR (Radiation Oncology)

PetCure Oncology San Jose, CA, USA Jessica Quimby, DVM, PhD, DACVIM (SAIM)

Ohio State University Veterinary Medical Center Columbus, OH, USA Jacquie S. Rand, BVSc, DVSc, MANZVS, DACVIM (SAIM)

School of Veterinary Science University of Queensland Gatton, Queensland, Australia and Australian Pet Welfare Foundation Kenmore, Queensland, Australia Robert B. Rebhun, DVM, PhD, DACVIM (Oncology)

Department of Surgical and Radiological Sciences University of California, Davis Davis, CA, USA Nicki Reed, BVM&S, CertVR, DSAM (Feline), DECVIM‐CA, MRCVS

Veterinary Specialists West Lothian, Scotland, UK Alexander M. Reiter, Dipl.Tzt., Dr.Med.Vet., DAVDC, DEVDC

School of Veterinary Medicine University of Pennsylvania Philadelphia, PA, USA Lisa V. Reiter, DVM, DACVD

McKeever Dermatology Clinics Eden Prairie and Inver Grove Heights, MN, USA Teresa Rieser, DVM, DACVECC

Veterinary Specialty Care Mt. Pleasant, SC, USA Steven Rosenthal, DVM, DACVIM (Cardiology)

CVCA Cardiac Care for Pets Towson, MD, USA

­List of Contributors

Jan Rothuizen, DVM, PhD, DECVIM‐CA

Erin M. Scott, DVM, DACVO

Department of Clinical Science of Companion Animals Utrecht University Utrecht, The Netherlands

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA

Elizabeth A. Rozanski, DVM, DACVIM (SAIM), DACVECC

VCA Alameda East Veterinary Hospital Denver, CO, USA

Tufts University Medford, MA, USA Craig Ruaux, BVSc (Hons), PhD, MACVSc, DACVIM (SAIM)

School of Veterinary Science Massey University Palmerston North, New Zealand Erin E. Runcan, DVM, DACVT

Department of Veterinary Clinical Sciences Ohio State University Columbus, OH, USA David Ruslander, DVM, DACVIM (Oncology), DACVR (Radiation Oncology)

Veterinary Specialty Hospital of the Carolinas Cary, NC, USA Silke Salavati, Dr.Med.Vet., PhD, DipECVIM‐CA, MRCVS

Royal School of Veterinary Studies University of Edinburgh Edinburgh, Scotland Ashley B. Saunders, DVM, DACVIM (Cardiology)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Brian A. Scansen, DVM, MS, DACVIM (Cardiology)

Katherine Scott, DVM, DACVIM (SAIM)

Christine M. Serafin, DVM, DACVIM (SAIM)

The Animal Surgical Center of Michigan Flint, MI, USA Stephanie L. Shaver, DVM, DACVS

School of Veterinary Medicine University of California, Davis Davis, CA, USA Marco A. Coutinho da Silva, DVM, MS, PhD, DACT

Ohio State University Columbus, OH, USA Nicole Smee, DVM, MS, DACVIM

Las Vegas Veterinary Specialty Center Las Vegas, NV, USA Laia Solano‐Gallego, DVM, PhD, DECVCP

Facultat de Veterinária Universitat Autònoma de Barcelona Barcelona, Spain Maria M. Soltero‐Rivera, DVM, DAVDC

VCA San Francisco Veterinary Specialists San Francisco, CA, USA Mitchell D. Song, DVM, DACVD

Colorado State University Fort Collins, CO, USA

VETMED Specialty Referral and 24‐Hour Emergency Care Veterinary Hospital Phoenix, AZ, USA

Jennifer R. Schissler, DVM, MS, DACVD

Joachim Spergser, Dipl.Tzt., Dr. Med.Vet., DECVM

College of Veterinary Medicine and Biomedical Sciences Colorado State University Fort Collins, CO, USA Peggy L. Schmidt, DVM, MS, DACVPM

College of Veterinary Medicine Kansas State University Manhattan, KS, USA

Institute of Microbiology University of Veterinary Medicine Vienna, Austria Rebecca L. Stepien, DVM, MS, DACVIM (Cardiology)

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA

Nick A. Schroeder, DVM, DACVIM (Cardiology)

Tammy Stevenson, DVM, DACVIM

LeadER Animal Specialty Hospital Cooper City, FL, USA

Veterinary Specialty Hospital San Diego, CA, USA

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­List of Contributors

Susanne M. Stieger‐Vanegas, DVM, PhD, DECVDI

Avenelle I. Turner, DVM, DACVIM (Oncology)

Carlson College of Veterinary Medicine Oregon State University Corvallis, OR, USA

Veterinary Cancer Group Culver City, CA, USA

Jason W. Stull, VMD, MPVM, PhD, DACVPM

Midtown Animal Hospital Gering, NE, USA

Department of Veterinary Preventive Medicine Ohio State University Columbus, OH, USA Jan S. Suchodolski, MedVet, Dr. Med.Vet., PhD, AGAF, DACVM

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Karen Lynn C. Sueda, DVM, DACVB

VCA West Los Angeles Animal Hospital Los Angeles, CA, USA Steven Suter, VMD, MS, PhD, DACVIM

Canine Bone Marrow Transplant Unit North Carolina State University College of Veterinary Medicine Raleigh, NC, USA Jane E. Sykes, BVSc, PhD, DACVIM (SAIM)

Jerry J. Upp, DVM

Shelly L. Vaden, DMV, PhD, DACVIM (SAIM)

College of Veterinary Medicine North Carolina State University Raleigh, NC, USA Katrina R. Viviano, DVM, PhD, DACVIM (SAIM), DACVCP

School of Veterinary Medicine University of Wisconsin‐Madison Madison, WI, USA J. Scott Weese, DVM, DVSc, DACVIM (SAIM)

Ontario Veterinary College University of Guelph Guelph, ON, Canada Raegan Wells, DVM, DACVECC

Phoenix Veterinary Referral and Emergency Center Phoenix, AZ, USA Joanna Whitney, BSc, BVSc, PhD, MVetStud, MACVS (SMAnimMed, ECC)

University of California, Davis Davis, CA, USA

Small Animal Specialist Hospital North Ryde, NSW, Australia

Séverine Tasker, BSc, BVSc (Hons), PhD, DSAM, DACVIM‐CA, PGCertHE, MRCVS

Wayne E. Wingfield, MS, DVM, DACVS, DACVECC

Bristol Veterinary School University of Bristol Bristol, UK Katie Tolbert, DVM, PhD, ADCVIM (SAIM)

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX, USA Elizabeth E. Toops, DVM, MS, DACVD (Dermatology)

Virginia Veterinary Specialists Charlottesville, VA, USA Sheila M.F. Torres, DVM, MS, PhD, DACVD

College of Veterinary Medicine University of Minnesota St Paul, MN, USA

Department of Clinical Sciences Colorado State University Fort Collins, CO, USA Zachary M. Wright, DVM, DACVIM (Oncology)

VCA Animal Diagnostic Clinic Dallas, TX, USA Panagiotis G. Xenoulis, DVM, Dr. Med.Vet., PhD

College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, TX

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About the Companion Website This book is accompanied by a companion website: www.wiley.com/go/bruyette/clinical The website includes: Podcasts

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Part 1 Diagnostic Considerations

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78 Epidemiology of Infectious Disease Peggy L. Schmidt, DVM, MS, DACVPM1 and Helen T. Engelke, BVSc, MPVM, DACVPM, MRCVS2 1

 College of Veterinary Medicine, Kansas State University, Manhattan, KS, USA  College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA

2

Infectious diseases pose many challenges for veterinarians. How was my patient exposed to the pathogen? When did exposure occur? Where did the disease ­originate? Are there others infected? Does my patient pose a risk to other pets or people? How can I determine if things are getting better or getting worse? Understanding the epidemiology of infectious disease can help veterinarians answer these and other questions which can lead to better informed decisions which then improve patient outcomes and mitigate further spread of disease. Two essential areas of infectious disease epidemiology for the practicing veterinarian to  understand are  performing disease monitoring and surveillance activities and outbreak investigations.

­Disease Monitoring and Surveillance Disease monitoring and surveillance are important activities designed to assess the health and disease status of a population of animals. Monitoring and surveillance, while sounding similar to the lay person, are considered epidemiologically distinct terms. Specific diseases, described syndromes, or general health status are monitored by veterinarians through the systematic collection and recording of data. Disease monitoring activities ­provide information which can inform treatment and prognosis of affected animals, but does not typically elicit an intervention at the population level. Disease surveillance activities may include collection of the same data used for disease monitoring, but activities are more intensive and purposeful. Surveillance is an ongoing, somewhat cyclic event which to be effective occurs over a considerable period of time. Surveillance methods can include testing programs for new additions to populations, such as FeLV testing of new cats entering an animal shelter or cattery, or annual testing for

i­nfectious disease, such as annual heartworm testing for dogs. In surveillance, when a specified threshold disease level is reached, prevention, control, or eradication measures are triggered and applied at the population level. Although most small animal clinical practices deal with individual animals rather than owned populations, disease monitoring and surveillance activities are no less important to the clinic population. For example, veterinarians may erroneously assume low disease prevalence in their area because of historic patterns of disease, thereby preventing early detection of disease in that area. As people, animals, and vectors move more freely and establish new areas of residence, previously rare diseases may incrementally increase in incidence. Small animal veterinarians also have a responsibility to participate in population surveillance programs established for public health. Public health agencies at both the state and federal level maintain a list of reportable diseases which include zoonotic diseases that affect companion animals. While maintaining a focus on human cases of disease, an increasing number of agencies now include reporting of zoonotic disease cases in companion animals. For example, the California Code of Regulations requires veterinarians to report cases of ­category A bioterrorism agents, such as anthrax, plague, and tularemia, as well as zoonotic diseases such as brucellosis (other than B. canis) and rabies to local health officials. Veterinarians should check routinely with their state veterinarian and local health departments to keep current on reporting regulations related to zoonotic disease and potential bioterrorism agents. Small animal veterinarians may serve on the front line for preventing incursions of foreign animal diseases. In 2007, an astute small animal veterinarian in Mississippi found fly larvae on a 16‐year‐old dog imported from Trinidad and reported the findings to the Mississippi state veterinarian. Further diagnostics identified the fly

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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larva as Cochliomyia hominivorax, the New World screwworm, which had been successfully eradicated from the US in the 1960s. Disease monitoring and surveillance activities can be formal or informal and help to identify increases in incidence of specific diseases or disease syndromes. Taking the time to formalize a hunch, collect and analyze data, and report findings to veterinary or public health ­officials may be the key to early identification of disease outbreaks within a practice population.

­Outbreak Investigations Outbreak investigations are parallel to a diagnostic work‐ up of an individual patient. A series of steps, performed in a specific order, provide information which helps describe and diagnose the problem at hand, plus allows insight into measures to mitigate disease and prevent future spread. Sources will vary slightly on the number of steps involved in an outbreak investigation but this chapter will focus on a six‐step process for outbreak investigation. The outbreak investigation process mimics the clinical investigation process for individual animals with disease: recognize there is a problem, define the problem, gather subjective and objective information to better describe the problem, formulate a list of likely differentials, perform diagnostics to rule in or rule out differentials, design a treatment plan, and communicate findings with the client. Goals of an outbreak investigation are to verify the pathogen responsible for the disease, identify the source of disease in the population, determine methods of transmission, and define means to prevent further spread and future occurrences. Pre‐step (or Step Zero): Establish an Investigation Team With outbreaks of food‐borne or infectious disease affecting people, a team of physicians and epidemiologists is established prior to beginning an investigation. In small animal medicine, resources to assist in an investigation may be available through local departments of public health, especially if the outbreak has zoonotic potential or may significantly impact the mental well‐ being of owners or the community. Assistance may also be available from county or state veterinarians or local colleges of veterinary medicine. Step 1: Confirm Presence of Disease Outbreak Before moving forward with an investigation, confirmation of the presence of disease above normal levels must occur. Ideally, this would involve comparing ­current disease levels with historic endemic levels of disease in

the area. Published prevalence studies from the literature may provide insight into endemic disease levels in specific geographic areas or populations. If endemic disease levels are unavailable, levels could be estimated by extracting diagnosis information from electronic medical records in previous years. Step 2: Establish a Case Definition Commonly seen diseases with available accurate diagnostic tests allow for easy establishment of a working case definition. For example, outbreaks of canine parvovirus are not uncommon and are reliably diagnosed by a combination of clinical signs and diagnostic test results. The disease under investigation may only present as a collection of clinical signs rather than through isolation and identification of a specific pathogen. In this situation, the case definition would include significant or unique clinical signs, signalment information, animal location (area of residence), and time of onset of disease. Additional data from animals who meet the case definition should be collected to further inform the ­ investigation. Data should include pertinent information from the patient history, physical exam, and diagnostic procedures. Patient outcomes or response to treatments should also be collected and recorded. As the investigation moves forward, the case definition will be continually refined in response to new information. Step 3: Descriptive Epidemiology Once the working case definition has been established, investigators should review available historical medical records to identify previous animals meeting the case definition. This allows for identification of the time point for the start of the outbreak. Data from previous cases, including species, breed, age, and sex, should be used to calculate prevalence of disease in various populations and further describe the disease epidemiology. Generating hypotheses for risk factors for disease will depend on this information. Key descriptors for the outbreak investigation include temporal and geographic spread of disease. One of the easiest ways to describe these data is through epidemic curves (Figure 78.1). Epidemic curves inform hypotheses for disease introduction and transmission, estimate incubation periods, recognize possible pathogens, and identify possible interventions. Point source epidemic curves are indicative of a common pathogen exposure in a group of animals in a single or relatively brief time point. They are characterized by a sharp slope at the beginning of the outbreak and a more gradual slope at the end. For infectious diseases, spread of the disease is limited to a single incubation period and further spread into naive populations does not occur.

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and risk factors can prevent implementation of successful interventions. This may lead to treatment failure and continued spread of the disease. Observational studies facilitate comparisons between affected and nonaffected populations (cases and controls) with measures of association. Retrospective case– control studies are relatively quick and easy to perform, but may be hampered by incomplete or imperfect data  collection or availability in medical records. Odds ratios can be calculated to identify risk factors for ­disease and support hypothesized control or prevention interventions. Prospective cohort studies are more expensive and time consuming compared to case–control studies, but provide incidence data which lead to more accurate ­estimates of risk through relative risk calculations. Their prospective nature provides an opportunity for more complete data collection in both affected and nonaffected populations which would not be available in ­historical medical records. As with case–control studies, cohort studies also identify potential risk factors for ­disease and support interventions to reduce or prevent further disease spread. Step 5: Implement Control and Prevention Measures Descriptive epidemiologic data from epidemic curves and risk factors determined from observational studies must be carefully analyzed to determine what interventions will provide the greatest benefit and likelihood of success. Measures must be directed at controlling the current outbreak through treatment, if possible, and ­preventing continued spread of the disease into naive animals in the population. In small animal clinical practice, these actions can ­provide significant challenges not encountered in populations under ownership or control of relatively few parties. When a population of animals has a single owner, the potential loss of a few animals is mitigated by the benefits of disease prevention in the larger population. Sacrificing a few animals for diagnostic ­ procedures or to prevent further disease spread often makes economic sense. With companion animals, the value of individual animals does not lie just in dollars and cents, but also in an unquantifiable emotional value as a member of the family. The good of the population does not necessarily override the value of the individual, creating difficult choices for mitigating disease at a population level. Recommendations for treating individual animals are  likely to be accepted and, depending on the cause

of the outbreak, individuals may have a good chance to recover from infection. Treatment options may be cost‐ prohibitive for some owners or treatment may be refused for various reasons. This may have little effect on spread of disease if owners follow strict isolation and quarantine protocols. Unfortunately, some owners may refuse ­quarantine and isolation procedures designed to stop the  spread to naive animals and veterinarians have few options to mandate such procedures without a risk to public health or safety. In these situations, effective and compassionate communication of risk may help increase compliance and mitigate risk of further spread of disease. ­Step 6: Communicate Findings For public health outbreak investigations, communication of findings includes informing the healthcare community of the “who, what, where, when, why, and how” of the outbreak. This can increase awareness of the potential for continued spread of the disease and allow implementation of preventive measures and early detection of the disease in new populations. If the disease outbreak under investigation has a public health implication, and local public health officials were not part of the investigation process, it is important to communicate relevant findings to the local public health department. Findings should include date of onset and conclusion (if outbreak has ended), case definition/­ diagnosis, populations affected, epidemic curves, and outcomes of control and prevention measures. This information should also be shared with other veterinarians in the practice area so results of the investigation can be used to control and prevent disease in the larger ­population of animals in the community. Findings from new or novel disease outbreaks should also be communicated to the larger veterinary audience through publication in journals or presentations at ­continuing education functions. Increased awareness may prevent further outbreaks or mitigate the effects of outbreaks if they spread to new companion animal populations.

­Conclusion Epidemiology is an integral part of the daily practice of veterinary medicine whether veterinarians realize it or not. Tools and techniques for disease monitoring and surveillance and outbreak investigations are readily available and can add to a veterinarian’s epidemiology

78  Epidemiology of Infectious Disease

toolbox. Just as epidemiology is a science focused on populations, veterinarians should not work in isolation when investigating the incidence of disease in their ­practice populations. Local, regional, state, and federal

veterinarians and health professionals should be part of the companion animal healthcare team responsible for rapid identification and resolution of diseases in the clinic and community populations.

­Further Reading California Department of Public Health. 2011. Title 17, California Code of Regulations (CCR) §2500, §2593, §2641.5‐2643.20, and §2800‐2812 Reportable Diseases and Conditions. www.cdph.ca.gov/ Programs/CID/DCDC/CDPH%20Document%20 Library/ReportableDiseases.pdf (accessed June 24, 2019).

California Department of Food and Agriculture. List of Reportable Conditions for Animals and Animal Products. www.cdfa.ca.gov/ahfss/animal_health/pdfs/ CA_reportable_disease_list_poster.pdf (accessed June 12, 2019). JAVMA News. Dog with screwworms brought to Mississippi. J Am Vet Med Assoc 2007; 231(9): 1320–33.

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79 Laboratory Diagnosis of Infectious Diseases Laia Solano‐Gallego, DVM, PhD, DACVP1 and Gad Baneth, DVM, PhD, DACVP2 1 2

Dep. Medicina i Cirurgia Animals, Facultat de Veterinària, Universitat Autònoma de Barcelona, Barcelona, Spain School of Veterinary Medicine, Hebrew University, Rehovot, Israel

Veterinarians use diagnostic testing for infectious ­diseases for two main reasons: ●●

●●

to confirm acute or chronic infection in a dog or cat with clinical signs or clinicopathologic abnormalities compatible with infectious diseases to detect subclinical infection or certify that animals are free of infection.

Detection of infection may be pursued to screen ­clinically healthy animals living in endemic regions, to prevent transmission by blood transfusion, to avoid importation of infected dogs and cats to nonendemic countries, to avoid transmission of disease to people in contact with the animal, including immune‐suppressed owners, to monitor response to treatment, and for research. Different diagnostic procedures can be used depending on the purpose of the diagnostic investigation and the pathogen most likely affecting the dog or cat based on the differential diagnosis list. Test results might also be interpreted according to the aim of the diagnosis, type of pathogen, and clinical status of animals. Accurate diagnosis of infectious diseases requires an integrated routine approach consisting of thorough ­clinical history, physical examination, pertinent routine laboratory tests such as complete blood count (CBC), complete biochemical profile and urinalysis, diagnostic imaging and additional diagnostic testing, depending on the differential diagnoses list for each individual patient. In addition, specific pathogen diagnostic assays should be carried out when infectious diseases are at the top of the differential diagnoses list. This chapter describes general concepts of diagnosis of viral, bacterial, protozoal, and fungal infectious diseases with an overview of the most common diagnostic methods employed for these infections. Specific diagnostic features of common infectious diseases in dogs and cats, geographic distribution, main transmission modes, chief

clinical signs, clinicopathologic abnormalities, treatment, prognosis, and prevention will be described in detail in other chapters in this section. The most commonly employed techniques for the diagnosis of infectious diseases in small animals include microscopic examination of microorganisms in cytologic preparations or histopathologic specimens, serology, polymerase chain reaction (PCR), and culture of the organism in appropriate medium, as described in the following subsections of this chapter. The advantages and disadvantages of these diagnostic techniques are listed in Table  79.1. Sample collection, transport, and preservation of specimens for the different diagnostic tests and pathogens are described in Table 79.2.

­Protozoal and Arthropod‐Borne Infections Microscopic Examination Diagnosis can be based on cytologic or histologic detection of pathogens, either contained inside cells or free in routinely stained smears. Detection by light microscopy of pathogens in cytologic preparations or histopathologic specimens might be difficult, depending on the type of microorganisms causing infection. However, cytology is a more rapid and simple technique for the detection of some microorganisms compared with histopathology. Histopathology commonly requires special staining to detect bacterial, mycobacterial, fungal or protozoal infections. Standard Romanowsky stains such as Giemsa are used for cytologic detection of the majority of organisms (Figure  79.1). Special stains can also be used in cytology for enhancing the visualization of microorganisms but are less frequently needed. Although the specificity of these methods is high, it is important to highlight that identification of species

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Table 79.1  Advantages and disadvantages of common diagnostic methods for the detection of infectious diseases in dogs and cats Diagnostic techniques

Microscopic examination (cytology /histopathology)

Advantages ●●

●●

●●

●●

Serologic testing (antibody detection)  

– Qualitative

●●

●●

●●

Disadvantages

Permits direct detection of the pathogen and nature of the host response. Findings could be suspicious of infection or allow exclusion of other differential diagnoses Cytology is rapid and less invasive than obtaining tissues for histopathology by biopsy Cytology usually permits easier visualization of pathogens when compared with histopathology Detection of specific antibodies against the pathogen causing infection Permits evaluation of seroconversion to confirm recent infection Rapid in‐clinic test

●●

●●

●●

●● ●● ●●

●●

●● ●●

●●

– Quantitative (IFA, ELISA)

Serologic testing (antigen detection) Molecular testing

●●

●●

●● ●● ●●

Culture

●●

●●

Determines the antibody level which is of major importance in some diseases such as leishmaniasis where high antibody levels in the presence of compatible clinical signs and/or clinicopathologic abnormalities are conclusive of clinical leishmaniasis Allows the detection of pathogen

●●

●●

●●

●●

Allows the detection of pathogen DNA and RNA High sensitivity and specificity for target loci Allows determination of pathogen load (by real‐time PCR) Permits the isolation of pathogens and their maintenance for further comparisons and analysis Facilitates in‐depth identification of pathogens

●●

●●

●● ●●

●●

Relatively low sensitivity for the detection of pathogens in tissues or body fluids Requires the performance of other diagnostic tests such as immunohistochemistry and/or PCR when pathogens are not visualized Does not distinguish between morphologically similar organisms Requires expertise Does not detect the pathogen itself Typically does not discriminate between vaccinated and naturally infected dogs or cats Serologic cross‐reactivity between related organisms is possible Provides only positive or negative results Variable sensitivities and performance with risk of false‐negative results due to conservative cut‐off level and use of recombinant proteins A positive result will benefit from additional validation by quantitative serology Performance and accuracy of cut‐off vary between laboratories Frequent lack of sufficient standardization of techniques between laboratories Low antibody levels frequently require further testing May not be sensitive when low numbers of pathogens are present False‐positive results possible due to DNA contamination or to amplification of erroneous targets including host DNA Different techniques used by diagnostic laboratories and lack of standardization Time‐consuming and laborious Requires special equipment and biohazard conditions and therefore restricted to specialized laboratories May require up to one month to provide a result

Source: Modified from Solano‐Gallego and Baneth (2016). Reproduced with permission of John Wiley & Sons. ELISA, enzyme‐linked immunosorbent assay; IFA, immunofluorescence assay; PCR, polymerase chain reaction.

based solely on morphology is often not possible and molecular analysis is required for speciation. Sensitivity will depend on the time spent searching for microorganisms, type of pathogen, tissue sampled, amount of organism in the tissue (parasite load), and clinical status of the patient. Generally speaking, higher diagnostic sensitivity is found in sick animals compared with subclinically infected dogs or cats. The diagnosis of chronically infected and carrier animals remains a diagnostic ­challenge due to low and often intermittent parasitemia,

or low tissue pathogen load, which frequently make pathogen observation by microscopic evaluation difficult. Therefore, the use of molecular diagnostic assays is strongly recommended in those cases. In addition, identification of protozoa in formalin‐fixed, paraffin‐ ­ embedded sections of different tissues may be facilitated by immunohistochemical methods such as immunoperoxidase staining or in situ hybridization techniques. The optimal tissue or body fluid for sampling will depend on types of pathogen involved and lesions found

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Table 79.2  Collection, processing, and transport recommendations for different diagnostic tests in small animal infectious diseases. Follow all governmental guidelines relevant to a given infectious disease agent and clinical specimen Diagnostic test

Specimen type

Collection and processing

Shipment

Histology

Any tissue specimen

Samples collected for histology should never be >1 cm thick (preferably 5–7 mm thick). Fix in 10% buffered formalin (10× volume)

Double bag and leak‐proof container with adequate fixative

Cytology

Any tissue specimen or fluid analysis

Air‐dried smear should be prepared immediately after the sample has been collected to minimize cell deterioration. Alcohol fixation is not needed. Analysis of various effusions and fluids usually includes determination of protein content, total cell concentration, and cytologic examination. A sample of effusion/fluid should be collected into an EDTA tube for routine analysis. A second sample should be collected into a serum tube if any biochemical analyses are to be performed or if a bacterial culture is desired

Blood or cytologic smears should never be mailed to the laboratory in the same package with formalin‐fixed tissues because formalin vapors will produce artifacts in the specimen. Cytologic preparation should be transported at room temperature in a plastic container. Fluid samples should be shipped chilled but not frozen

Antibody or antigen testing

Blood, plasma, serum, cerebrospinal fluid, urine samples or feces

Serology generally requires serum, but plasma is often satisfactory. For serum samples, the blood should be drawn into a plain tube or a separator tube. The sample should be held at room temperature for 20–30 min to allow complete clot formation and retraction. The sample should then be centrifuged at high speed (~1000 g) for 10 min. In some instances, paired samples may be required for an adequate diagnosis

Specimens can be refrigerated for up to a week prior to shipment to the laboratory. Specimens should be frozen at −20 °C or −80 °C if stored for a longer period. Specimens can be stored frozen for years without loss of antibody levels. For specimens collected for assays for antigen detection, results may be more susceptible to variation with long specimen storage. The laboratory should be contacted in order to determine specimen storage and handling requirements

Molecular testing

Any specimen

Collect aseptically to prevent contamination. For blood or bone marrow and DNA analysis, use preferably EDTA or acid citrate dextrose (ACD) anticoagulant

For DNA analysis: submit specimens held at room temperature within 24 h of collection. Specimens can be stored at 2–8 °C for 72 h, or at −20 °C or ≤−70 °C for prolonged storage. For RNA analysis: use stabilization solution or send immediately to the laboratory on wet ice. For storage, freeze specimens at or below −70 °C

Isolation‐ culture

Any specimen

Collect aseptically to prevent contamination. Some agents might have special requirements such as anaerobic culture or special media. Culture from blood should be collected in special media bottles designed to prevent coagulation and neutralize antibiotic residues if animal was treated

Normally, specimens should be refrigerated and not frozen and delivered directly to the laboratory within 24 h

on physical examination. Blood‐borne organisms such as Babesia (Figure 79.2) can be found in blood smears, concentrated and stained buffy coat or splenic ­specimens. Other important tissue samples used for the detection of these pathogens are lymph node aspirates, skin touch impressions, bone marrow, joint fluid, cerebrospinal fluid (CSF) or other body fluids and tissues. Serologic Testing (Antibody and Antigen Detection) Antibody Detection

Several serologic methods can be used to detect specific serum antibodies directed against protozoal and arthro-

pod‐borne pathogens. Methods include the ­indirect fluorescent antibody (IFA) test, the enzyme‐linked immunosorbent assay (ELISA), immunochromato­ graphic rapid in‐house devices, direct agglutination assays, and western blotting. The most common serologic tests employed in clinical practice are quantitative assays (IFA and ELISA) and qualitative tests (rapid in‐house devices). In general, most of these methods have good sensitivities and specificities but sensitivity and specificity greatly depend on the antigens employed. Whole‐parasite extracts are sensitive for the detection of subclinical or clinical canine and feline infections but provide lower specificity. On the other hand, assays that employ recombinant protein antigens are very specific but may lack

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Figure 79.1  Lymph node aspirate from a dog. Note the increased numbers of plasma cells and macrophages with abundant intracellular Leishmania amastigotes (modified Wright stain, original magnification ×400).

s­ensitivity for the detection of infection, depending on the antigen employed and the level of serum antibodies. Serologic cross‐reactivity is common between pathogens of the same genus or otherwise closely related organisms. Therefore, cross‐reactivity between antibodies directed at different pathogens is possible with some serologic tests, especially those based on whole‐parasite antigen, and is less likely to occur when using recombinant protein antigens. It is important to note that the use of antigens of similar species will frequently result in cross‐reaction at a high level. For example, antibodies against Babesia canis will cross‐react with Babesia vogeli antigen. Pathogens of closely related different genera are more likely to cross‐react at low levels, such as the cross‐ reactivity between antibodies to Trypanosoma cruzi and Leishmania infantum. Antibody detection can indicate past exposure or present acute or persistent infection. Acute diseases ­ may be difficult to diagnose due to the lack of detectable antibody production and low pathogen load in blood or other tissues. Therefore, false‐negative results are possible

in peracute or acute diseases. Evidence of seroconversion is fundamental in the diagnosis of some acute diseases. In these cases, the measurement of acute and convalescent antibody levels (paired samples) is c­ onfirmatory for acute infection. Antibody level determination may vary between different laboratories, and therefore, it is recommended to use the same laboratory and assay for comparison of antibody levels. Furthermore, variations are also likely between assays carried out on different days. Therefore, ideally, paired samples should be tested at the same time. Seroconversion is considered when a fourfold or greater increase in antibody level is demonstrated in paired samples over a 2–4‐week period, and in some ­particular infections longer than that, for instance in L. infantum infection where a serologic response ­develops months after initial infection. In addition, a­ntibodies induced by vaccination may be detected by serologic assays, making it impossible to discriminate between vaccinated and naturally infected animals. Examples of this include vaccination against B. canis and L. infantum. High antibody levels are associated with high tissue parasite loads and disease in some protozoal and arthropod‐borne chronic diseases. In contrast, cases of animals with suspected clinical signs and compatible clinicopathologic abnormalities and low antibody levels require the use of additional detection methods to exclude or confirm the disease, because low antibody levels may also be detected in subclinical carriers suffering from a different clinical disease. Canine leishmaniasis is a good example of a disease in which moderate to severe clinical disease is usually manifested by medium to high antibody levels, whereas subclinical infection is more often associated with lower antibody levels or seronegativity. Antigen Detection

The detection of circulating plasma or blood antigen specific for a pathogen is an additional and usually very specific serologic technique. This type of assay detects the presence of a circulating plasma or blood antigen. For example, the most common detection technique of  canine heartworm infection caused by Dirofilaria Figure 79.2  Blood smears from two different dogs. (Right) A large form of Babesia. (Left) Small forms of Babesia (modified Wright stain, original magnification ×1000).

79  Laboratory Diagnosis of Infectious Diseases

immitis is based on detection of a circulating secretory antigen of the adult female worm. Molecular Diagnostics Polymerase chain reaction is a sensitive and specific diagnostic technique frequently employed for the diagnosis of protozoal and arthropod‐borne diseases. It is used for diagnostic purposes, monitoring during and after treatment, research studies, and for screening of blood donors. Moreover, it is particularly useful for detection of infection in animals with low parasitemia levels and for speciation of pathogens. PCR can be ­carried out on DNA extracted from tissues, blood, body fluids, conjunctival and oral swabs, or even cytologic preparations or histopathologic specimens. A large number of PCR assays and protocols using a variety of gene targets have been described for the detection of protozoal and arthropod‐borne infections of dogs and cats. It is important to highlight that for diagnostic purposes, the best DNA target will often be the locus or gene with the largest number of copies per organism. For instance, the kinetoplast DNA (kDNA) of L. infantum is an excellent target as it has about 10 000 copies in each Leishmania amastigote. Other targets are more useful for distinguishing between species such as the leishmanial ribosomal internal transcribed spacer 1 (ITS1). Due to the fact that in some endemic areas, more than one species co‐exist and infect animals, molecular techniques have been developed to discriminate DNA from different species of the same genus or related genera. These techniques include semi‐nested PCR, reverse line blotting, PCR restriction fragment length polymorphism (RFLP), and high‐resolution melting curve quantitative fluorescence resonance energy transfer PCR. Sequencing may also reveal infections with novel organisms that have not been described before. It is important to highlight that negative results by molecular techniques only indicate that specific DNA was not detected under the assay conditions and should not be interpreted as absolute evidence for the absence of infection. In addition, false‐positive results are possible due to DNA contamination or the amplification of DNA from other sources which may not be noticed if sequencing is not performed. Controls should be included in each step of the assay to ensure that DNA contamination has not occurred. Isolation Diagnosis may also be established by culture of the infectious agent. However, this technique is usually tedious and time‐consuming. It sometimes requires special ­conditions and a long duration for obtaining results and

therefore is less commonly used in clinical practice for parasites and arthropod‐borne pathogens. This technique is employed in research studies because it permits the identification and maintenance of pathogens. It is important to highlight that some organisms, such as Babesia species, are difficult to culture. The culture method is pathogen specific and usually requires special media and temperature conditions. Special culture medium is used for the isolation of some protozoal ­pathogens, such as L. infantum.

­Viral Infections Microscopic Examination Most viral infections are not diagnosed by microscopy. Viruses are too small to be detectable by light microscopy, but typical cytoplasmatic or intranuclear inclusion bodies caused by viruses may be detected in blood or ­tissue cells. Electron microscopy may be used for the detection of certain viruses such as canine parvovirus in feces of dogs shedding the virus, although this is a ­cumbersome, time‐consuming technique which is often unable to specify the exact species involved in infection, as it relies on morphology. Serologic Testing (Antibody and Antigen Detection) Serologic tests are frequently used for the detection of viral infections in cats and dogs. There are a number of basic serologic techniques such as ELISA, IFAT, virus neutralization, complement fixation, agglutination, agar gel immunodiffusion, and western blotting. Serology can be aimed at detecting specific antibodies produced against a certain virus in the infected animal, as in the most common tests for feline immunodeficiency virus (FIV) which detect antibodies to the p24 core protein. It can also be aimed at the actual detection of a certain viral antigen in a body fluid such as the blood, as in the common feline leukemia virus (FeLV) antigen test based on the detection of the viral p27 capsid protein. Rapid commercial tests are available for many of the common viral infections, and they are also frequently based on the detection of either antibodies against the virus or virus antigen. Rapid commercial tests are often based on immunochromatographic devices which produce a color response when antibodies against a virus are detected, or when the actual viral antigens are detected, if the assay is geared to the detection of antigen. These types of assays are often used for in‐house rapid diagnosis in veterinary clinics and hospitals. Several manufacturers have produced such rapid assays for the detection of antibodies against FIV, antigenemia with FeLV, or the

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presence of parvoviral antigen in the feces of dogs shedding this virus. The rapid assays that detect antibodies against a virus are often not quantitative and only produce a positive or negative result, in contrast to tests done in a specialized laboratory which frequently ­provide an antibody titer or quantitative measure related to the level of antibodies, such as the optical density in the ELISA. Molecular Diagnostics Molecular diagnosis of viruses includes the detection of specific nucleic acid sequences present in the targeted virus. Viruses may contain DNA or RNA, depending on the viral family to which they belong. PCR assays are commonly associated with the amplification of the nucleic acid sequence selected as the target for the assay and visualization of the product on a gel in conventional PCR, or its amplification and release of measurable fluorescence by real‐time PCR which can also be quantitative. Amplification of nucleic acids from RNA viruses may require an additional reverse transcription step in which RNA is translated into cDNA and then amplified. RNA is less stable than DNA and is susceptible to rapid degradation unless stored in an RNA stabilization solution and frozen at –70 °C or below. A multiplex reverse transcription‐nested PCR assay has been developed to differentiate between wild‐type and vaccine strains of canine distemper virus. An additional molecular diagnostic technique is the use of nucleic acid probes which attach specially to nucleic acid sequences in viruses and do not involve amplification of the virus’s DNA or RNA. Isolation Isolation of virus from blood, secretions, or organ tissues is a common method of diagnosis for viral infections of dogs and cats. Isolation requires the use of appropriate media to transport the tested sample, a specialized laboratory, and rapid transportation to the laboratory in refrigeration. Viruses are usually taken from the transport media or the original fluid in which they were submitted (e.g., blood or CSF) and grown in various cell culture lines. Often virus isolation is attempted in multiple cell lines simultaneously as the host cell preference of the particular virus strain is unknown. Virus isolation from animals is usually performed with the addition of antibiotics and antifungal drugs to cell lines to prevent contamination with bacteria and fungi. While feline retroviruses such as FIV and FeLV are usually isolated from blood, isolation of canine distemper virus is usually attempted from the CSF, bronchoalvelor lavage, and transtracheal lavage specimens.

­Bacterial Infections Microscopic Examination Most bacteria are detectable by high‐power light microscopy of cytology specimens (Figure 79.3) but this is usually not sufficient for species identification and may serve as a relatively insensitive measure for the presence of bacteria in the sample. Furthermore, some anatomic sites such as the skin, oral cavity, gastrointestinal tract, and upper airway normally contain bacteria and it is usually difficult to distinguish between the normal flora and infection with pathogenic bacteria just by microscopic examination of the organisms. Gram stain, acid‐fast staining, and other staining methods are ancillary ­techniques that can assist in the initial assessment of bacterial infection. Dark‐field microscopy is helpful in the detection of spirochetes such as leptospires in urine samples. Serologic Testing Serology can be useful in the detection of some bacterial infections of dogs and cats. Detection of specific antibodies is helpful in the diagnosis of leptospirosis and a battery of serogroup antigens is used to find which serovar demonstrates the highest antibody titer and is either responsible for infection or closest to the Leptospira serovar present in the animal. Serology is also useful for the detection of exposure to intracellular bacteria such as Ehrlichia canis or Anaplasma phagocytophilum. Isolation and Identification (Including Molecular Testing) Culture has been the major technique for detection and characterization of bacterial pathogens for the past century. There are a number of different media for growth of bacteria from various groups with specific metabolic requirements. Often, one type of medium is used for the transport or initial growth of bacteria and other media are subsequently used for subculture with enrichment and selection of the bacteria suspected as cause of infection. The MacConkey agar medium selects for gram‐ negative bacteria. Cultures from an infection site are often placed initially into more than one medium when a broad spectrum of infective agents is suspected, for instance in aerobic and anaerobic specific media which also require specific growth conditions such as an ­anaerobic environment enriched with carbon dioxide for anaerobes. Identification and characterization of isolated bacteria are usually done based on biochemical qualities and tests such as the catalase test and coagulase production.

79  Laboratory Diagnosis of Infectious Diseases

Commercial biochemistry test strips enable the identification of cultured bacteria and automated commercial identification systems are often used in large bacteriology laboratories for rapid bacteria identification. Some bacteria such as Anaplasma platys and hemotrophic mycoplasms (e.g., Mycoplasma hemofelis, Candidatus Mycoplasma hemominutum) are unculturable. The detection of these bacteria relies mainly on molecular biology techniques such as PCR amplifying DNA sequences which are specific for the suspected bacteria. Other bacteria such as Rickettsia spp., E. canis, and A. phagocytophilum are strictly intracellular and can only be grown in appropriate cell culture lines, restricting their isolation to research laboratories with expertise. Molecular techniques can also be useful for the identification of bacteria that have been propagated in culture and require classification to the species and strain levels. These can be sometimes be achieved by multigenic sequencing and characterization, with phylogenetic analysis. Providing the bacteriology laboratory with a good clinical history, physical examination and initial blood test work‐up and cytology findings can be very helpful in the selection of different isolation techniques. Sampling the correct organ sites is imperative and should be based on clinical judgment. Common body fluids and anatomic sites sampled for bacteriology from dogs and cats include urine, blood, feces, bronchoalveolar lavage, skin and ear, abdominal and pleural fluids, CSF, and cornea. Interpretation of Culture Results

The results of culture should be interpreted cautiously, considering that the bacterium isolated, when culture is successful, may not actually be the cause of infection, or that in some other cases, culture is negative despite bacterial infection. This can be due to several reasons including technical difficulties in transporting the culture with infectious agent, use of incorrect isolation technique or culture site, intermittent shedding of ­bacteria to the blood or other body fluid, and previous antibiotic treatments. Antimicrobial Susceptibility Tests

Determination of minimum inhibitory concentrations (MIC) for isolated bacteria can be helpful in deciding which antibiotic should be used for treatment of susceptible infections. This is also important in the detection of antimicrobial resistance, if present. The standard techniques for antibiotic susceptibility testing involve either gel diffusion or dilution methods. Although they can provide an idea of the antibiotics to which the isolated bacteria is susceptible in vitro, this does not always conform with the actual susceptibility due to in vivo factors such as accessibility to the infected tissue (e.g., decreased

accessibility to abscesses), concentration of antibiotic in the affected tissue, and inhibition of antibiotics by body proteins.

­Fungal Infections Fungal agents have a remarkably varied spectrum of clinical picture: primary skin disease, single to multiple subcutaneous masses, mass lesions within body cavities, disseminated disease with multiple organ dysfunction, and disseminated disease that presents as single‐organ dysfunction or as a cause of sudden death. Clinicopathologic findings are not specific but mild anemia of chronic disease, leukocytosis, hypergammaglobulinemia, hypoalbuminemia, and hypercalcemia can be present in some of these diseases. Primary (i.e., Blastomyces, Coccidioides, Cryptococcus, Histoplasma) and opportunistic pathogens, mainly saprophytic soil fungi (i.e., Aspergillus, Basidiobolus, Cladosporium, Conidiobolus), can be responsible for these diseases. Opportunistic fungi can cause disease by trauma and often present as a single subcutaneous mass. When dissemination occurs, it is often associated with ­ decreased immunocompetence. Microscopic Examination Suspicion of fungal diseases should be made when macrophagic or macrophagic‐neutrophilic inflammation is noted in any tissue by cytologic or histopathologic ­specimens. Bacterial inflammatory lesions (Figure 79.3) that are unresponsive to appropriate medical therapy should be investigated further to rule out underlying mycotic disease due to the fact that bacterial and fungal co‐infections do occur. Cytological evaluation of any tissue or body cavity fluid specimen is very useful for identification of fungal disease in dogs and cats. Cytology is a more rapid and simple technique for the detection of some fungal microorganisms than histopathology. Histopathology commonly requires special staining such as periodic acid–Shiff (PAS) or Gomori methenamine silver (GMS) to detect fungal infections. In addition, some laboratories offer immunohistochemistry for fungal organisms within tissues submitted for routine histopathology. Yeasts, different types of spores, and hyphae can be observed in cytologic specimens (Figure 79.4). Cytologic features of most fungal pathogens are described elsewhere. Routine Romanowsky staining adequately stains most fungal organisms within cytologic preparations. Organisms that do not stain well appear as negatively stained elements (Figure 79.5).

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Figure 79.3  Kidney aspirate from a cat. Note neutrophilic‐ macrophagic inflammation with infection of rod‐shaped bacteria (modified Wright stain, original magnification ×1000).

Figure 79.5  Inguinal subcutaneous mass in a dog. Note the marked neutrophilic inflammation and the evidence of unstained and partially pale basophilic hyphae (modified Wright stain, original magnification ×400).

Serologic Testing (Antibody and Antigen Detection)

Figure 79.4  Thoracic mass from a German Shepherd dog. Note the necrotic background, increased numbers of neutrophils and Aspergillus terreus hyphae (modified Wright stain, original magnification ×1000).

India ink staining is commonly used for the identification of Cryptococcus species. The expertise of the individual evaluating the slide is the main variable in using cytology in the clinical setting. The sensitivity of cytologic and histologic evaluation varies considerably although specificity is high. Cytologic and histopathologic features cannot definitively identify the species and therefore culture and biochemical characterization or molecular analysis are mandatory for identification. It is important to highlight that morphologic features of fungi in tissue differ when grown on fungal media.

Serology is used in the clinical setting to diagnose fungal disease in dogs and cats. However, interpretation of serologic results in some fungal diseases might be difficult and will benefit from additional cytologic or molecular evidence or culture. Limited studies are available regarding sensitivity and specificity of serologic testing of some fungal diseases. The majority of fungal serologic tests currently in use in veterinary medicine detect the antibody response to infection. A few tests detect fungal antigens, providing a definitive diagnosis if the antigen is sufficiently specific (such as the antigen‐based latex agglutination test available for Cryptococcus neoformans). As with any serological test, cross‐reactivity can exist between fungal agents. Positive serology provides evidence of exposure and supports the clinical findings. Paired serum samples (2–4 weeks apart) should be run at the same time. A fourfold increase or decrease in the antibody level is strong evidence of active disease. More details on current serologic tests available for fungal disease are described elsewhere in this section and in the literature. Isolation and Identification (Including Molecular Testing) Culture

The specific identification of fungal organism is based on culture of fluids or tissue. However, successful culture of fungal agents depends on several variables, including the

79  Laboratory Diagnosis of Infectious Diseases

fungal organism itself, concentration of fungal elements in the sample, sample integrity, culture requirements of  the fungal agent, and expertise of the laboratory ­performing the culture. Obtaining and submitting the appropriate sample for  culture and providing the laboratory with detailed history and clinical diagnosis are essential for a successful culture. Samples of urine, exudates, and body cavity fluids can be submitted in sterile serum collection tubes. Up to 10 mL of fluid is recommended. Samples of fresh tissue from a lesion can be submitted in a sterile container for culture. Identification of fungal agents in cultures is time‐ consuming. The culture has to grow and form characteristic fruiting bodies, conidia, or arthrospores to allow morphologic identification of the fungus. Microscopic morphology of fungal reproductive structures is the most useful criterion for identification. It is strongly advised that fungal culture (other than for dermatophytosis) is performed in a reference diagnostic laboratory.

Molecular Testing

Molecular testing is less commonly employed for the diagnosis of fungal diseases in tissues or body fluids in veterinary medicine as it is for bacterial, viral, and protozoal diseases. Molecular techniques are the same as reported for other infectious diseases and include conventional, nested, and real‐time PCR and sequencing. Common target genes are the 18S or 5.8S ribosomal gene or internal transcribed spacer (ITS) regions of ribosomal DNA. Fungal identification based on DNA sequencing is not always definitive because results depend on whether or not the DNA of the fungus in question has been previously sequenced and submitted to the genomic databases. Therefore, fungal identification based on DNA sequencing should include at least two DNA regions for comparison. In contrast, PCR‐based methods are commonly used to confirm identification of fungi culture in diagnostic laboratories. Commercial chemiluminescent‐labeled DNA probes are available for some fungal agents such as Blastomyces, Coccidioides, and Histoplasma.

­Further Reading Goldstein RE. Canine leptospirosis. Vet Clin North Am Small Anim Pract 2010; 40: 1091–101. Schulz BS, Strauch C, Mueller RS, Eichhorn W, Hartmann K. Comparison of the prevalence of enteric viruses in healthy dogs and those with acute haemorrhagic diarrhoea by electron microscopy. J Small Anim Pract 2008; 49: 84–8. Si W, Zhou S, Wang Z, Cui SJ. A multiplex reverse transcription‐nested polymerase chain reaction for

detection and differentiation of wild‐type and vaccine strains of canine distemper virus. Virol J 2010; 7: 86. Solano‐Gallego L, Baneth G. Diagnosis of protozoal and arthropod‐borne diseases. In: Villiers E, Ristic J, Blackwood L, eds. BSAVA Manual of Canine and Feline Clinical Pathology. Gloucester, UK: BSAVA, 2016. Solano‐Gallego L, Koutinas A, Miro G, et al. Directions for the diagnosis, clinical staging, treatment and prevention of canine leishmaniosis. Vet Parasitol 2009; 165: 1–18.

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80 Canine Distemper David S. Bruyette, DVM, DACVIM Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology Canine distemper virus (CDV) is a large (100–250 nm) ssRNA virus belonging to the genus Morbillivirus of the family Paramyxoviridae. Mutations affecting the CDV H protein required for virus attachment to host cell receptors are associated with virulence and disease emergence in novel host species. CDV has a lipoprotein envelope and a nonsegmented negative‐stranded RNA genome consisting of six genes that code for a single envelope‐ associated protein (matrix [M]), two glycoproteins (hemagglutinin [H] and fusion [F] proteins), two transcriptase‐associated proteins (phosphoprotein [P] and large protein [L]) and the nucleocapsid (N) protein, which encapsulates the viral RNA. Only one serotype of CDV is recognized with several co‐circulating genotypes based on variation in the H protein. Canine distemper has affected dogs worldwide for centuries, with descriptions of disease outbreaks in the European literature dating back to the 17th century. It was first formally described in dogs in 1905. Despite the introduction of modified live vaccines in the 1950s and their extensive uptake, the disease remains prevalent. Systemic CDV infection, resembling distemper in domestic dogs, can also be found in wild canids (e.g., wolves, foxes), procyonids (e.g., raccoons, kinkajous), ailurids (e.g., red pandas), ursids (e.g., black bears, giant pandas), mustelids (e.g., ferrets, minks), viverrids (e.g., civets, genets), hyaenids (e.g., spotted hyenas), and large felids (e.g., lions, tigers). In addition, besides infection with the closely related phocine distemper virus, seals can become infected by CDV. In some CDV outbreaks, including the mass mortalities among Baikal and Caspian seals and large felids in the Serengeti Park, terrestrial carnivores including dogs and wolves have been suspected as vectors for the infectious agent. Lethal infections have been described in noncarnivore species such as peccaries

and nonhuman primates demonstrating the remarkable ability of the pathogen to cross species barriers. Canine distemper virus infection of dogs is characterized by a systemic and/or nervous clinical course and viral persistence in selected organs, including the central nervous system and lymphoid tissue. Main manifestations include respiratory and gastrointestinal signs, immunosuppression and demyelinating leukoencephalomyelitis (DL). The primary mode of infection is via inhalation. After initial exposure, dogs may mount a robust immune response and recover. It is estimated that as many as 75% of infections may actually be subclinical. Initially, CDV replicates in lymphoid tissue of the upper respiratory tract. Monocytes and macrophages are the first target cells which propagate the virus. Impaired immune function, associated with depletion of lymphoid organs, consists of a viremia‐associated loss of lymphocytes, especially of CD4+ T cells, due to lymphoid cell apoptosis in the early phase. After clearance of the virus from the peripheral blood, an assumed diminished antigen presentation and altered lymphocyte maturation cause an ongoing immunosuppression despite repopulation of lymphoid organs. Following a variable incubation period (1–4 weeks), animals develop a characteristic biphasic fever. During the first viremic phase, generalized infection of lymphoid tissues with lymphoid depletion, lymphopenia, and transient fever are observed. Profound immunosuppression is a consequence of leukocyte necrosis, apoptosis, and dysfunction. The second viremia is associated with high fever and infection of parenchymal tissues such as the respiratory tract, digestive tract, skin, and CNS. The early phase of DL is the result of direct virus‐mediated damage and infiltrating CD8+ cytotoxic T cells associated with an upregulation of proinflammatory cytokines such as interleukin (IL)‐6, IL‐8, tumor necrosis factor (TNF)‐ alpha, and IL‐12 and a lack of response of immunomodulatory cytokines such as IL‐10 and transforming growth

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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factor (TGF)‐beta. A CD4+‐mediated delayed‐type hypersensitivity and cytotoxic CD8+ T cells contribute to myelin loss in the chronic phase. Additionally, upregulation of interferon‐gamma and IL‐1 may occur in advanced lesions. Moreover, an altered balance between matrix metalloproteinases and their inhibitors seems to play a pivotal role for the pathogenesis of DL. Histologic manifestations include polioencephalitis and DL. The observations of p75NTR‐expressing bipolar Schwann cell‐like glia in CDV‐DL highlight the potential of Schwann cell‐mediated remyelination and raise the hope for a beneficial regenerative process in CDV‐DL. Though the central role of axonal damage as a triggering event is established in CDV‐DL, the role of macrophage polarization in CDV‐DL is currently unknown. In light of the fact that distinct macrophage responses have been linked to axonal regeneration, degeneration, demyelination, and remyelination, the elucidation of a potential macrophage polarization during CDV‐DL appears to represent a promising field for future investigations and the development of new treatment strategies. Recovery from CDV depends on the host immune response. A strong and effective cellular immune response can eliminate the virus prior to infection of parenchymal tissues, while weak and delayed cellular and humoral immune responses lead to virus spread and persistence, respectively.

­Epidemiology Canine distemper is a fatal disease of dogs with a worldwide distribution. With a mortality rate of around 50%, it is second only to rabies when it comes to virus‐induced fatality in dogs.

­Signalment There is no breed or sex predilection. Animals 4 months   Healthy dogs often shed CCoV

Enterocytes Moderate villous atrophy, deepening of crypts Pantropic variant has increased virulence and causes systemic disease

Usually mild, self‐limiting diarrhea and vomiting   Pantropic variant – fever, lethargy, hemorrhagic diarrhea, lymphopenia, neurologic signs

A distantly related Beta‐ coronavirus causes respiratory disease in dogs similar to SARS‐CoV in humans

Canine distemper virus (CDV) (Morbillivirus, Paramyxoviridae)

ssRNA, enveloped; intracytoplasmic & intranuclear inclusions   Closely related to human measles & rinderpest viruses

Domestic and wild canids; wild felids; variants infect raccoons, skunks, ferrets, marine mammals

3–6 months Macrophages, lymphocytes, epithelial cells, neurons, glial cells (pancytotropic)   Mild‐moderate villous atrophy, hyperkeratosis, lymphoid and thymic depletion, encephalomyelitis

Multisystemic disease: lethargy, fever, vomiting, diarrhea, coughing, dyspnea, nasal and ocular discharge, conjunctivitis, keratitis, photophobia, seizures, dementia, ataxia, hyperesthesia, hyperkeratosis

“Old dog” encephalitis – rare form of CDV disease in older dogs years after primary infection

Canine rotavirus (Rotavirus, Reoviridae)

Segmented (11) dsRNA, non‐enveloped; reassortment with A rotaviruses

Domestic and wild canids  See Additional information column

Neonates

Mature enterocytes   Mild villous atrophy

Usually mild, self‐limiting diarrhea

Interspecies transmission and zoonotic potential with reassorted viruses

Canine kobuvirus (Kobuvirus, Picornaviridae)

ssRNA non‐enveloped, icosohedral capsid

Unknown Domestic dogs and foxes, likely other canids – very few epidemiologic studies reported

Not yet characterized

Has been isolated from stool samples of diarrheic dogs, but pathogenicity is currently unknown

Genetically similar to human Aichi virus A, a gastroenteritis transmitted through consumption of oysters

Dog circovirus (Circovirus, Circoviridae)

Domestic dogs; wild ssDNA, non‐enveloped, canids unknown icosohedral capsid   Closely related to porcine circovirus

Not yet characterized

Unclear – has been associated with hemorrhagic gastroenteritis, vasculitis and vascular necrosis Granulomatous lymphadenitis, thrombocytopenia

Similar prevalence in healthy and sick dogs; probably most important as a co‐pathogen with CPV‐2 – increased mortality

6 weeks–6 months

82  Canine Viral Enteritis

derived immunity wanes and remain susceptible until they acquire active immunity, either through natural infection or through vaccination. The outcome of infection, ranging from subclinical infection to severe disease, seems to depend on individual host factors, with some breeds (notably American pit bull terriers, Rottweilers, Doberman pinschers, and German shepherds) having increased risk for severe enteritis.

­History and Clinical Signs Most puppies infected with CPV‐2 present with a history of lethargy and inappetence. These signs are often accompanied by fever, abdominal pain, vomiting and diarrhea, which may lead to substantial dehydration. Although profuse, hemorrhagic diarrhea is a typical finding, many dogs with CPV enteritis will produce scant mucoid feces instead. Extensive damage to the small intestinal mucosa may result in coliform bacterial septicemia and endotoxic shock. Lymphopenia is a consistent clinicopathologic finding; other changes may include neutropenia, anemia, thrombocytopenia, electrolyte imbalances (especially h ­ ypokalemia), hypoglycemia, and hypoalbuminemia. Serum C‐reactive protein levels are often elevated in critically ill dogs and correlate loosely with increased risk of mortality. Myocarditis occurs with perinatal infection of puppies born to naive bitches; puppies may present with dyspnea, weakness, and crying or experience sudden death. This syndrome is not commonly seen in vaccinated and/ or CPV‐endemic populations.

­Diagnosis Although parvovirus infection is the likely diagnosis in an unvaccinated puppy presenting with enteritis, fever, and lymphopenia, other enteric viruses may have a similar presentation. Diagnosis of parvoviral enteritis is best accomplished by observation of clinical signs along with detection of CPV‐2 antigen (point‐of‐care enzyme‐ linked immunosorbent assay [ELISA] or hemagglutination) or DNA (polymerase chain reaction [PCR]) in feces or rectal swab specimens. PCR‐based tests are more sensitive than antigen tests and can be used to identify the variant of the infecting CPV.

­Therapy Ideally, puppies infected with CPV‐2 should be hospitalized and kept in isolation. Therapy for CPV‐2 enteritis consists of aggressive supportive care to correct hydration, hypoglycemia, and electrolyte abnormalities such as

hypokalemia, control pain and maintain nutrition. Antimicrobials should be administered to prevent and treat secondary bacterial infections (gram‐negative and anaerobic) that arise from bacterial translocation across the compromised gastrointestinal tract or from neutropenia. Frequent (q8h) monitoring of blood glucose is necessary and supplementation of intravenous (IV) fluids with dextrose is often needed. Colloid therapy with hetastarch or plasma may be necessary in some patients, particularly those with severe hypoalbuminemia (Figure 82.1). Red cell transfusion may be necessary in patients with significant blood loss. While treatment with recombinant human granulocyte stimulating factor and hyperimmune serum has not been shown to be of benefit, feline interferon‐omega has shown promising results.

­Prognosis Puppies with severe enteritis have a very poor survival rate without treatment due to hypovolemic and septic shock and electrolyte/acid–base disturbances, while hospitalization with fluid therapy greatly improves their prognosis for recovery. A protocol for outpatient treatment of puppies whose owners cannot afford hospital care has been developed and is showing promise for improving the prognosis in these patients.

­Prevention Prevention of CPV‐2 infection is accomplished by limiting contact of susceptible animals with potentially infected animals and premises while stimulating immunity with vaccination. Shelters, kennels, clinics, and other high‐density units need to practice good sanitation, disinfection, and appropriate use of quarantine protocols and isolation facilities. Because maternally derived antibody can interfere with immunization, vaccination with modified live virus vaccine is recommended every 3–4 weeks beginning at 6 weeks of age until the puppy is 16 weeks old (or 20 weeks old for high‐risk breeds). Although current vaccines, which are composed of CPV‐2 or CPV‐2b, have been effective in protecting dogs against CPV‐2c infection under experimental conditions, it is still unclear how protective these vaccines are under some field conditions.

­Public Health Implications Canine parvovirus‐2 variants do not infect people, but feces should be treated as if other infectious agents that cause intestinal disease in people are present.

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83 Viral Papillomatosis Margaret C. Barr, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

Dogs and cats, like many other species, are susceptible to development of papillomatous growths (warts) on the skin and mucous membranes. Most of the time, viral papillomas are benign and transient lesions but they can significantly affect the health of an animal if they pro­ gress to premalignant or malignant neoplastic lesions. Canine and feline papillomas and the viruses that cause them share many features and will be discussed together in this chapter.

­Etiology/Pathophysiology Papillomaviruses (PV) are small nonenveloped viruses with a circular double‐stranded DNA genome enclosed in an icosohedral capsid. They belong to the large and diverse viral family Papillomaviridae, which currently includes over 280 types of papillomaviruses infecting most (if not all) mammalian species and many nonmam­ malian species. Each PV type demonstrates a strong host and tissue/site preference, with only a few examples of interspecies transmission. Papillomaviruses replicate in epithelial cells of skin and mucous membranes, beginning their life cycle in the basal layer and progressing as the cells mature. Viral assembly and particle release are restricted to the stra­ tum granulosum and stratum corneum. This pattern of replication is orchestrated by the complex expression of a series of early and late genes. Infections are often subclinical, with frequent detec­ tion of papillomavirus DNA in normal skin of healthy dogs and cats. The typical presentation of papillomatosis (cutaneous or oral warts) occurs when high levels of PV early gene expression induce rapid expansion of the basal epithelial cell population followed by cell differentiation, epithelial thickening, hyperkeratosis, and exophytosis.

Progression to malignant lesions occurs rarely in both dogs and cats, with some species differences in histologic features and clinical presentation. Disregulation and overexpression of two early genes, E6 and E7, have been linked to the development of PV‐associated squamous cell carcinomas (SCCs). In most animals, mild inflammation and a cell‐mediated immune response lead to spontaneous regression of papil­ lomas within 6–8 weeks. A humoral immune response produces antibodies that protect against reinfection with closely related types of PV. Immunocompromised animals may develop persistent and generalized papillomatosis with an increased risk of progression to malignant neo­ plastic transformation.

­Epidemiology Papillomaviruses infect domestic and wild canids and felids throughout the world. Although only a few canine and feline PV have been sequenced to date, almost 150 human PV types have been reported. In general, PVs are highly host specific; however, bovine papillomavirus type 1 infection causes equine sarcoid tumors in horses, and ruminant‐like PVs have been linked to the develop­ ment of feline sarcoids. Papillomaviruses are transmitted through epidermal microabrasions or transepidermal inoculation of virus with contaminated fomites. Because the virus lacks an envelope, it is relatively resistant to adverse environmen­ tal conditions and difficult to kill with disinfectants. These facts, along with the ubiquity and diversity of PVs, make it likely that most animals will be infected at some point in their lives, but there is little information on the prevalence of PV infection in canine and feline populations.

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­Signalment Dogs and cats of all ages and breeds can be infected with PVs. Young dogs are most likely to be affected by canine oral papillomatosis, while older animals with chronic persistent infections are more likely to develop bowe­ noid in situ carcinoma (BISC) and invasive SCC. Young pugs and miniature schnauzers seem to be dispropor­ tionately affected by canine pigmented plaques, suggest­ ing that host genetic factors play a role in the development of this disease. Oral papillomas in dogs rarely progress to SCC but there is some indication that malignant trans­ formation of canine oral papillomas may be increasing. Oral and cutaneous papillomatoses are not observed as often in cats as in dogs; viral plaques are also uncom­ mon but have been more clearly associated with PV infections in cats. Feline BISCs develop in middle‐aged to older cats, occur disproportionately in cats with feline retrovirus infections, and are most severe in hairless breeds. Cutaneous and oral SCCs are common malig­ nancies of older cats, with ultraviolet (UV) exposure being an important risk factor for some of these lesions. While feline PV DNA has been identified frequently in cutaneous SCCs found in UV‐protected areas of the body, a strong association of oral SCC with PV infection in domestic cats has not been established.

­Clinical Signs Papillomatosis and viral plaque lesions are typically lim­ ited to the cutaneous or mucosal epidermis, sometimes occurring singly but more often in multiples. Lesions can range from papular or nodular lesions to pedunculated or cauliflower‐like, hyperkeratotic masses. Infection rarely generates more than a mild inflammatory response, so systemic signs are not seen. Bowenoid in situ carcinomas develop as multicentric irregular regions of epidermal and follicular hyperplasia primarily on the face, shoulders, and limbs. These premalignant lesions are hyperkeratotic, pigmented plaques that may ulcerate. If progression to SCC occurs, it is usually as a single non­ metastatic but potentially highly invasive tumor. A sum­ mary of diseases associated with canine and feline PV infections is provided in Table 83.1.

­Diagnosis Diagnosis of oral papillomatosis is based on clinical ­presentation for most typical cases in young dogs. Other types of lesions require full‐thickness excisional biopsy with histologic examination for a morphologic diagn­ osis. The etiologic diagnosis of papillomatosis can be

Table 83.1  Papillomavirus‐associated disease in domestic dogs and cats

Disease

Species affected

Oral papillomatosis

Canine Feline – rare

Multiple, raised, smooth nodular to pedunculated, hyperkeratotic masses on oral mucosa and lips (dogs), or ventral tongue (cats); spontaneous regression is common

Exophytic cutaneous papillomatosis

Canine Feline – rare

Single to multiple pedunculated, hyperkeratotic masses often on head or feet; spontaneous regression is common

Endophytic cutaneous papillomatosis

Canine

Unpigmented, raised firm masses with central pores found on ventral abdomen or feet; do not usually regress

Pigmented plaques

Canine

Small to medium‐sized flat to slightly raised, darkly pigmented, hyperkeratotic masses found in clusters on the limbs, axilla, ventral trunk or abdomen; do not usually regress; may progress to in situ carcinoma

Viral plaques

Feline

Multiple variably sized, slightly raised, hyperkeratotic masses that may or may not be pigmented; some may spontaneously regress while others progress to in situ carcinoma

Clinical presentation

Feline Multiple scaly, crusting to Bowenoid in situ carcinoma Canine – rare ulcerating plaques frequently on face, neck and limbs; some (BISC) may be slowly progressive and others become highly invasive or progress to squamous cell carcinoma Squamous cell carcinoma (SCC)

Feline Usually a solitary flattened Canine – rare ulcerated mass found in UV‐ protected areas of the body (as opposed to SCCs caused by UV exposure); rarely metastasize but can be highly invasive and damaging to surrounding tissue

Feline sarcoid

Feline

UV, ultraviolet.

Slow‐growing, solitary to multiple, smooth dense fibroblastic nodules that may ulcerate, located on head, neck and extremities; do not metastasize and are not highly invasive

83  Viral Papillomatosis

c­onfirmed by immunohistochemical staining for PV group‐specific antigens, in situ hybridization for PV nucleic acid detection, or electron microscopy to visual­ ize viral particles. Polymerase chain reaction is a highly sensitive and rapid method for detection of PV DNA in biopsy specimens, but interpretation is difficult because PV DNA is frequently amplified out of normal canine and feline skin.

­Therapy and Prognosis Spontaneous regression of many nonmalignant PV‐ associated lesions makes treatment unnecessary in most cases. Immunosuppressive drugs like glucocorti­ coids and ciclosporin should be avoided, if possible, in infected animals. Provision of a high‐quality diet and management of stress may help support an animal’s immune response to the virus. Interferons and other immunomodulators have been tried in severe cases of papillomatosis, but solid evidence of their efficacy is not yet available. Azithromycin (10 mg/kg once daily for 10 days) was used in one small trial to treat canine oral and cutaneous warts with some apparent efficacy. The mechanism of action is unclear but it may be associated with immunomodulatory properties of the drug. Surgical removal or cryotherapy may be effective if few lesions are present although new lesions may develop.

Removal of premalignant and malignant lesions like BISCs and SCCs should include histologically confirmed normal tissue margins on all borders.

­Prevention Currently, there are no commercial vaccines available for canine and feline papillomatosis. Because most PV infec­ tions are subclinical or produce benign self‐limiting lesions, immunization has not been a priority. The diver­ sity of PVs makes it difficult to stimulate protective immunity against this family of viruses. If specific types of canine and feline PVs are demonstrated to be associ­ ated with the development of SCC, then targeting those viruses with vaccines would make sense. The use of vac­ cines against some types of human PVs associated with genital cancer suggests that vaccination could be effec­ tive against PVs in other species also.

­Public Health Implications Canine and feline papillomaviruses have not been dem­ onstrated to be infectious to humans. However, human PV‐9 was amplified from a cutaneous papilloma in a cat, and bovine PV causes equine (and possibly feline) sar­ coids, proving that cross‐species infection can occur under some circumstances.

­Further Reading Egberink H, Thiry E, Möstl K, et al. Feline viral papillomatosis: ABCD guidelines on prevention and management. J Feline Med Surg 2013; 15: 560–2. Lange CE, Favrot C. Canine papillomatosis. Vet Clin North Am Small Anim Pract 2011; 41: 1183–95. Munday JS. Papillomaviruses in felids. Vet J 2014; 199: 340–7. Munday JS, Thomson NA, Luff JA. Papillomavirus in dogs and cats. Vet J 2017; 225: 23–31.

Nagata M, Rosenkrantz W. Cutaneous viral dermatoses in dogs and cats. Compend Contin Educ Vet 2013; 35: E1–E10. Thaiwong T, Sledge DG, Wise AG, et al. Malignant transformation of canine oral papillomavirus (CPV1)‐ associated papillomas in dogs: an emerging concern? Papillomavirus Res 2018; 6: 83–9.

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84 Canine Influenza Virus Ellen Collisson, MS, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology

­Epidemiology

While influenza A viruses had for many years not been considered pathogens of concern in dogs, they are today a serious differential for canine respiratory problems. In 2004, a previously known equine strain of H3N8 influenza A virus emerged as a canine respiratory concern during an outbreak in greyhound racing dogs in Florida. Twenty‐ two dogs suffered from fever, followed by coughing and either recovery or, in the case of eight dogs, peracute death occurred associated with extensive hemorrhaging of the lungs, mediastinum, and pleural cavity. Severe respiratory illness with mortality was reported in dogs in 2008 in Korea and later in China and Thailand due to an H3N2 strain of influenza that has been shown to originate from the avian virus and to also transmit from dog to dog. In March 2015, canine influenza virus (CIV) H3N2 was identified in the United States in the Chicago area and during the same year identified in dogs in at least 30 states throughout southern, eastern and midwestern United States. In 2015, shelter cats in Indiana were also diagnosed with an H3N2 strain. Typical of other influenza A viruses, CIV viral particles capable of cell‐to‐cell or host‐to‐host transmission carry H (hemagglutinin) and N (neuraminidase) proteins that are anchored in the bilipid envelope coating the viral particle. These glycosylated proteins, used to define subtypes, are prone to mutations that result in the evolution of new influenza strains. A hallmark of influenza viruses is the eight‐segmented RNA genome which provides a mechanism of rapid genetic change through segment reassortment. Thus, infection with more than one type of influenza virus can result in progeny that are a combination of RNA segments from more than one parent strain. In addition, the RNA polymerase which generates the viral RNA segments for progeny virus is error prone, creating mutations that may contribute to altered viral tropism, pathogenesis, and antigenicity.

Influenza viruses are highly contagious and easily transmitted from dog to dog by aerosol or contact exposure, but the H3N2 also was found to infect cats, resulting in respiratory illness. The H3N8 virus has been shown experimentally to also infect cats. Animals in high‐density environments, such as shelters, competition events, and breeding facilities, are at greatest risk of infection. The original canine H3N8 outbreak in greyhounds, first recognized in January of 2004, had spread in a few months to tracks in six states, and as of April 2013 had been reported in 40 states (dogflu.com). Similar to influenza in other animals, a dog may be infected and shed contagious CIV for up to four days before clinical signs occur. Currently, infection of dogs is estimated to result in 80% morbidity and 1–5% mortality although higher rates of mortality have been reported. Humans have been reported to also carry the virus from an infected animal to a naive animal although the Centers for Disease Control (CDC) states that human infections with the CIV strains are not a health concern. This may be a matter of adaptation and infection with or without clinical illness. The virus isolated from the lungs of dogs in Florida during the first cases of CIV was found to be a canine‐ adapted H3N8 virus originating from an equine H3N8 subtype, recognized as respiratory pathogens of horses since 1963. Unlike previous reports of influenza infections in dogs, which were self‐limiting and not transmitted from dog to dog, the greyhound H3N8 virus of 2004 originating from an equine H3N8 virus and the H3N2 originating from an avian influenza virus were easily transmitted from dog to dog and, at least the latter, also from cat to cat. The sequencing of genomes of CIV isolates identified signature mutations that indicate stable changes in the canine‐adapted viruses.

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Cross‐species infections frequently occur with influenza A viruses but adaptation to a new species is not common. Dogs are susceptible to infection with human strains, normally without clinical illness or transmission to other dogs. It is not known why the H3 strains seem to have adapted to the dog as a stable host, but it is possible that other strains could, with the appropriate mutations, evolve into true canine pathogens. In 2012, a novel H3N1 human H1N1 and H3N2 reassortant virus was reported to infect dogs subclinically in Korea but was not reported to transmit from dog to dog.

­Signalment Any dog that has not developed immunity specific for the H3N8 virus can be infected regardless of age, sex or breed. Being a highly contagious respiratory virus, the length of time that a dog may remain in a shelter has been associated with increased risk of infection with CIV. With available vaccines and greater likelihood of natural exposure, more animals will develop immunity and morbidity will likely decrease in all ages and breeds, and both sexes.

­History and Clinical Signs Clinical history is indistinguishable from other respiratory diseases of dogs and resembles influenza infections in other animals. Morbidity may be expected to be at least 60%, but it has been reported that as many as 80% of infected animals may develop clinical signs of infection. Between two and four days following infection, dogs may present with clinical illness, such as fever, sneezing, dyspnea, anorexia, depression and coughing, and ocular and nasal discharge that can become mucopurulent. Illness, especially coughing, may continue for 10–30 days post infection. The mortality is generally low. A minority of infected dogs will develop pneumonia with tracheitis and bronchitis. Complications of secondary infections from bacteria or mycoplasma may exacerbate lung involvement, such as hemorrhage in the lungs, mediastinum and pleural cavity, and greater mortality.

­Diagnosis Because clinical illness is similar to that of other respiratory illnesses in dogs, CIV involvement must be diagnosed by detection of viral antigen, viral RNA or the presence of specific antibody. Because of the short duration of virus infection, serology may be a more reliable

means of diagnosing infection. Serologic evaluations usually require comparisons of paired sera to document seroconversion to confirm infection. CIV‐specific antibodies can be detected as early as seven days after infection. Hemagglutination inhibition assays are used to determine antibodies specific for the H serotypes. However, convalescent sera, in the absence of widespread exposure, can often be used to identify specific infection. Enzyme‐ linked immunosorbent assay (ELISA) can be used to detect group‐specific antibodies for influenza A viruses. The group‐specific (not type‐specific) ELISA is based on detecting specific antibody that binds to the nucleocapsid protein, which coats the viral genome segments and is highly conserved among all influenza A viruses. Virus may be detected in samples collected with cotton or Dacron nasal swabs within four days after clinical signs are observed; that is, within seven days after infection. Necropsy‐collected tissue from lungs or associated lymph nodes may be used to detect virus. Diagnosis of viral infection is often first determined using real‐time reverse transcriptase polymerase chain reaction (PCR) of the gene that encodes the highly conserved matrix protein in influenza virion. A positive PCR result may be followed by viral isolation (after up to three passages) using Madin–Darby canine kidney cells or embryonated chicken eggs in order to determine the H and N subtypes of the isolated virus. Subtyping the H and N of the viral isolates can also be determined by laboratories that have the tools for sequencing the viral genome. The hemagglutination inhibition assay is often used to determine the serotype of CIV strains in the US. Antibodies that react with virus inhibiting red blood cell agglutination of CIV serotype standards can be used to inhibit agglutination and thus identify the H subtype of the CIV isolate.

­Therapy Supportive therapy may be necessary to provide optimum conditions for the immune system to respond, eliminating infection and reducing the severity of clinical signs. Oxygen, intravenous fluids, and nebulization with coupage may be needed for dogs with pneumonia. Good nutrition and basic biosecurity measures that include disposable or dedicated protective clothing and the use of kennel‐approved disinfectants can help to prevent viral spread. Soap and water hand washing which destroys the bilipid envelope membrane of influenza viruses is highly effective in inactivating the virus. Isolation of infected or suspected infected animals may protect dogs not exposed to the virus or vaccine although exposure is difficult to assess since infection occurs prior to the appearance of clinical illness.

84  Canine Influenza Virus

Canine influenza virus survives in the environment for more than 24 hours. Because secondary bacterial and mycoplasma infections can cause complications, antimicrobials may be considered if clinical signs are severe or suggest co‐infection. Antiviral drugs exist for influenza, but such treatments have not been approved for use in dogs. Information is lacking regarding their efficacy, doses or negative side‐effects in dogs. The Nobivac® Canine Influenza H3N8 and H3N2 inactivated bivalent vaccine from Intervet/Merck is labelled as effective at reducing disease severity. However, the capacity for influenza A viruses to mutate provides the risk of eventual adaptation and evasion of immunity from administered vaccines.

­Prognosis From 60% to 80% of CIV‐infected dogs will present with clinical illness, but subclinically infected dogs will also

shed contagious virus. Dogs will completely recover clinically within 2–3 weeks, sometimes, however, with a lingering cough. Animals that are generally in good health and well nourished are expected to recover without complications. While most infections will be self‐limiting, mortality is considered to be between 1% and 5%.

­Public Health Implications With the reported transmissions of influenza from birds and other mammals to humans, we should consider CIV strains a potential public health concern. Influenza viruses are commonly transmitted across species although usually with subclinical infection and without further transmission within the second species. However, the CIV strains have not been reported to infect humans, even owners with known infected dogs, and the Centers for Disease Control does not consider the canine influenza viruses to be a human threat.

­Further Reading Crawford PC, Dubovi EJ, Castleman WL, et al. Transmission of equine influenza virus to dogs. Science 2005; 310(5747): 482–5. He W, Li G, Zhu H, et al. Emergence and adaptation of H3N2 canine influenza virus from avian influenza virus: an overlooked role of dogs in interspecies transmission. Transbound Emerg Dis 2019; 66(2): 842–51. Ramírez‐Martínez LA, Contreras‐Luna M, De la Luz J, et al. Evidence of transmission and risk factors for influenza A virus in household dogs and their owners. Influenza Other Respir Viruses 2013; 7(6): 1292–6.

Song DS, An DJ, Moon HJ, et al. Interspecies transmission of the canine influenza H3N2 virus to domestic cats in South Korea. J Gen Virol 2011; 92(Pt 10): 2350–5. Song D, Moon HJ, An DF, et al. A novel reassortant canine H3N1 influenza virus between pandemic H1N1 and canine H3N2 influenza viruses in Korea. J Gen Virol 2012; 93(Pt 3): 551–4. Su S, Wang L, Fu X, et al. Equine influenza A (H3N8) virus infection in cats. Emerg Infect Dis 2014; 20(12): 2096–9. Voorhees IEH, Glaser AL, Toohey‐Kurth K, et al. Spread of canine influenza A(H3N2) virus, United States. Emerg Infect Dis 2017; 23(12): 1950–7.

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85 Feline Parvovirus Margaret C. Barr, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

Feline parvovirus (FPV) is the causative agent of feline panleukopenia, a disease that has also been called feline infectious enteritis, feline plague or feline distemper.

­Etiology/Pathophysiology Feline parvovirus is so closely related to canine parvovirus type 2 (CPV‐2), mink enteritis virus, and raccoon parvovirus that all these viruses are now classified as a single species, feline panleukopenia virus. The genome of this small single‐stranded DNA virus is packaged tightly in an icosohedral, nonenveloped capsid that is very resistant to environmental conditions. The feline transferrin receptor allows FPV access to a broad range of host cells, where it replicates in actively dividing cells including lymphocytes, enteric crypt epithelial cells, and bone marrow myeloid progenitor cells. Similar to the pathology seen with CPV‐2 infection in dogs, intestinal villi are severely blunted, absorptive capacity is impaired, and gastrointestinal integrity is compromised in FPV‐ infected cats. The interruption of myeloid cell proliferation along with lymphocytolysis and redistribution of circulating white blood cells to inflamed tissues results in the characteristic and severe panleukopenia seen with this disease. Perinatal infection (approximately three weeks before birth to three weeks after birth) of kittens results in infection of the central nervous system and impacts development of the cerebellum. The virus replicates in the dividing neuroblasts of the external granule cell layer, causing cerebellar hypoplasia in kittens that survive the infection. There is also evidence of FPV replication in Purkinje cells of the cerebellum but it is unclear how the virus replicates in these presumably postmitotic cells.

­Epidemiology Domestic and wild felids are susceptible to FPV infection, along with some nonfelid species like mink, raccoons, and foxes. Although canids are resistant to FPV infection, only a few mutations in the VP2 capsid protein gene led to the emergence of CPV‐2. Feline parvovirus‐associated disease is most often seen in multicat housing situations like animal shelters and breeding catteries, especially if rigorous sanitation, isolation, and vaccination protocols are not followed. Parvoviruses are shed at high titer in the feces of infected animals for several days after infection. Transmission is through the fecal–oral route, either directly or indirectly on fomites such as shoes, clothing, or feeding dishes. Extensive prevalence data are lacking. The prevalence of anti‐FPV antibodies in cats entering a Florida animal shelter, indicating either past exposure or vaccination, was just under 40%. If other cat populations in the US are similar, about 60% of cats are at risk for FPV infection. Anecdotal reports also suggest a recent resurgence of feline panleukopenia outbreaks, possibly associated with antigenic variation in FPV variants and the associated decreased efficacy of commercial vaccines.

­Signalment Susceptible cats of all ages can be infected, but young kittens experience a higher mortality rate (up to 90%) than older animals if not treated. There are no breed or sex predispositions reported for FPV.

­History and Clinical Signs The typical FPV patient will have a history of exposure to other cats in a high‐density multicat environment and

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little or no evidence of vaccination for FPV after 12 weeks of age. About two‐thirds of FPV‐infected cats will present with vomiting or diarrhea (occasionally bloody), but these signs may be absent or only transiently observed and reported as part of the animal’s history. Anorexia, lethargy, and dehydration are also common clinical signs; fever is a less consistent finding. Sudden death with no apparent clinical signs may result from peracute FPV infection, especially in young kittens. Leukopenia, due to neutropenia and/or lymphopenia, is present in most affected cats, but some cats are able to maintain normal leukocyte counts or may even have leukocytosis in response to secondary bacterial infections. Anemia, hypoalbuminemia, and electrolyte imbalances often occur in these cats. Sepsis can lead to the development of disseminated intravascular coagulation.

­Diagnosis Diagnosis of FPV infection is based on history (potential exposure, poor vaccination history) and clinical signs, including characteristic hematology changes. Confirmation of the diagnosis can be made through detection of fecal parvovirus antigens using point‐of‐care assays that are marketed for detection of CPV‐2. Fecal samples or rectal swab specimens may also be submitted for PCR analysis of viral DNA.

­Treatment The same general principles of supportive care used for managing CPV‐2‐infected dogs can be applied to FPV patients (see Chapter 82). Rehydration and maintenance of electrolyte and acid–base balances, and blood glucose are of primary importance. Parenteral administration of broad‐spectrum antimicrobial agents is also indicated to combat sepsis due to loss of integrity of the gastrointestinal tract lining and neutropenia. Extrapolation from canine research suggests that early reintroduction of enteral nutrition as soon as vomiting is controlled may be beneficial. Cats with severe anemia or hypoproteinemia may require blood or plasma transfusions, and intravenous nutrition should be considered for these cats if enteral feeding is not possible. Ancillary treatments may include immunomodulatory therapies such as passive immunoglobulin transfer using feline serum containing anti‐FPV antibodies and active immune stimulation with feline recombinant interferon‐omega, although one study showed no difference in survival for cats treated with the latter agent compared to controls.

­Prognosis Without treatment, a cat with severe enteric disease and leukopenia due to FPV infection has a poor prognosis. It is not clear just how effective good nursing care is at improving the prognosis for recovery. One study noted a 51% recovery rate of ill cats with a median hospitalization time of seven days, with a poor prognosis associated with leukopenia (specifically due to neutropenia), thrombocytopenia, hypoalbuminemia, and hypokalemia. Age was not associated with outcome in this group of treated cats. However, a recent retrospective evaluation of outcome predictors and survival time for FPV‐infected shelter cats determined that only about 20% of hospitalized cats survived to discharge, with a median survival time of three days. In this population of cats, animals with signs of lethargy, low body weight, rectal temperature lower than 100.2 oF, and leukopenia during hospitalization had the poorest prognosis. Predictors of survival included history of antiparasitic therapy, and treatment with amoxicillin‐clavulanic acid and maropitant.

­Prevention As with any virus infection, prevention of exposure to FPV would be the ideal method for preventing infection; however, the sturdy nature of the viral particles and high potential for environmental contamination decrease the feasibility of this approach. Good sanitation practices, strict isolation procedures, and prevention of overcrowding are all important methods for preventing outbreaks of FPV in multicat situations. Vaccination of cats with commercial modified‐live or inactivated FPV vaccines is considered to be a safe and effective method of FPV prevention, but recent studies showing a high degree of interference with vaccination by maternally derived antibodies have led to a change in vaccination protocols. Kittens should be vaccinated beginning at 6–8 weeks of age, continuing every 3–4 weeks until 16–20 weeks of age. Cats older than 16 weeks should be vaccinated twice, four weeks apart. Revaccination is at one year after the primary series and then every three years. If preexisting FPV antibody titers are relatively high, revaccination may have little or no benefit.

­Public Health Implications Feline parvovirus is not infectious to humans and does not pose a public health risk.

85  Feline Parvovirus

­Further Reading Bergmann M, Schwertler S, Reese S, et al. Antibody response to feline panleukopenia virus vaccination in healthy adult cats. J Feline Med Surg 2018; 20: 1087–93. DiGangi BA, Levy JK, Griffin B, et al. Prevalence of serum antibody titers against feline panleukopenia virus, feline herpesvirus 1, and feline calicivirus in cats entering a Florida animal shelter. J Am Vet Med Assoc 2012; 241: 1320–5. Jakel V, Cussler K, Hanschmann KM, et al. Vaccination against feline panleukopenia: implications from a field study in kittens. BMC Vet Res 2012; 8:62. Kruse BD, Unterer S, Horlacher K, Sauter‐Louis C, Hartmann K. Prognostic factors in cats with feline panleukopenia. J Vet Intern Med 2010; 24: 1271–6. Poncelot L, Héraud C, Springinsfeld M, et al. Identification of feline panleukopenia virus proteins expressed in

Purkinje cell nuclei of cats with cerebellar hypoplasia. Vet J 2013; 196: 381–7. Porporato F, Horzinek MC, Hofmann‐Lehmann R, et al. Survival estimates and outcome predictors for shelter cats with feline panleukopenia virus infection. J Am Vet Med Assoc 2018; 253: 188–95. Truyen U, Parrish C. Feline panleukopenia virus: its interesting evolution and current problems in immunoprophylaxis against a serious pathogen. Vet Microbiol 2013; 165: 29–32. Truyen U, Addie D, Belák S, et al. Feline panleukopenia: ABCD guidelines on prevention and management. J Feline Med Surg 2009; 11: 538–46. (Updated most recently December 2017; www.abcdcatsvets.org/ abcd‐guidelines‐on‐feline‐panleukopenia‐2012‐ edition/.)

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86 Feline Coronavirus Yvonne Drechsler, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology Feline coronavirus (FCoV) is a highly contagious virus that exists as two bioforms. Feline enteric coronavirus (FECV) is ubiquitous in multicat environments. This virus commonly causes an asymptomatic infection, which can persist in certain individuals. Sporadically and unpredictably, the infection turns pathogenic, in which case the virus is referred to as feline infectious peritonitis virus (FIPV), causing the highly fatal, systemic immune‐ mediated disease feline infectious peritonitis (FIP). The current hypothesis is that FIPV arises from the FECV by mutation, replicates in monocytes and disseminates, resulting in vasculitis and pyogranulomatous inflammation. However, our understanding of the pathophysiology of FIPV is very limited due to difficulties in replicating the virus in laboratory settings.

­Epidemiology Feline coronavirus is distributed worldwide and is ubiquitous in virtually all cat populations. There is great variability in prevalence among different cat populations. Two serotypes exist, I and II, with most epidemiologic studies indicating that serotype I is more common. However, most studies have been conducted in countries outside the United States so further study on serotype prevalence in the United States is needed. The enteric virus, FECV, is readily transmitted via the fecal–oral route herefore, the prevalence of FCoV infection is generally associated with the number and density of cats housed together. The length of time spent in multicat environments also increases the risk of exposure. For example, the risk of exposure was estimated to be five times higher for cats living in shelters for longer than 60 days than for cats staying for shorter periods of time. Housing and

husbandry practices that reduce exposure to feces and contaminated environments have a tremendous influence on the number of cats exposed to the virus. Although transmission of the pathogenic FIP virus occurs is controversial outbreaks in shelters have been reported. Therefore, quarantine of suspected FIP‐infected cats is advisable.

­Signalment Although typically a disease of young cats and kittens, the age distribution of affected cats appears to be bimodal. Cats younger than 2 years of age or geriatric cats are most commonly diagnosed with FIP. In addition, certain pure breeds, most notably Persians and Burmese, appear to be more susceptible. Male and female cats are affected equally in some studies while others show a male predisposition. Immune‐compromised animals, such as cats infected with feline leukemia virus (FeLV) or feline immunodeficiency virus (FIV), are also at increased risk.

­History and Clinical Signs Typically, cats from shelters, catteries, and multicat households are at higher risk of exposure and development of FIP due to the ubiquitous nature of the enteric virus in these settings. Stress due to overcrowding may also contribute to immune compromise and increased susceptibility to infection. Genetic predisposition has also been hypothesized to play a role in increased susceptibility in certain breeds. Clinical signs associated with the benign enteric virus are typically gastrointestinal, manifesting as mild vomiting and/or persistent or intermittent diarrhea. In cases of FIP, cats may be asymptomatic or present with different levels of lethargy and anorexia. Other common findings include

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weight loss, pale mucous membranes, fever of unknown origin, and uveitis. Commonly, a distinction is made between the wet (effusive) and dry (noneffusive) forms of FIP, but they are not mutually exclusive. In the wet form, abdominal distension due to the accumulation of exudative fluid is the most common presentation. A fluid wave on physical examination may be evident, but some cases will have less fluid accumulation, only detectable by abdominal ultrasound. Pleural effusion with secondary dyspnea, tachypnea, and muffled heart sounds may be present, whereas pericardial effusion is uncommon. Common signs of the dry form are mild and intermittent fever, decreased appetite, weight loss, stunted growth, depression, pale or yellow mucous membranes, and palpable abdominal organ enlargement. Clinical signs may also reflect dysfunction of affected organs. Pyogranulomatous lesions may develop in one or more abdominal organs. If granuloma formation involves the intestine, constipation, diarrhea, or vomiting may be the major clinical signs observed. Enlarged mesenteric lymph nodes and palpable nodular irregularities on the surface of kidneys and liver may also be present on physical examination. Uveitis is the most common ocular abnormality documented in FIP cases, but other ocular lesions such as conjunctivitis, corneal precipitates, iritis, cuffing of the retinal vasculature and retinal detachment may be present. Neurologic signs can also be seen with FIP, the most common being behavioral changes, ataxia, central vestibular signs, hyperesthesia, nystagmus, and seizures. Any part of the CNS can become affected in this disease.

­Diagnosis With the exception of histopathology and immunostaining, no single laboratory test can definitely diagnose FIP. Therefore, the diagnosis of FIP continues to be based primarily on the combination of history of risk factors, signalment, clinical abnormalities, and laboratory findings. The diagnostic process starts with a good history and comprehensive physical examination. Complete Blood Test and Biochemical Profile Abnormalities may include anemia, which is usually nonregenerative. Evidence of hemolysis, with or without evidence of immune‐mediated destruction, may be present. Lymphopenia, neutrophilia, sometimes with a left shift and toxic change, and thrombocytopenia also occur. Hyperbilirubinemia and elevated aspartate aminotransferase (AST) and alanine aminotransferase (ALT) may also occur. Hyperproteinemia (>8.0 mg/dL) due to hyperglobulinemia is a consistent finding, present in approxi-

mately 60% of cats with FIP. The hyperglobulinemia is usually polyclonal, although monoclonal gammopathy may occur. Hypoalbuminemia can be present, possibly due to hepatic insufficiency, vasculitis, protein‐losing nephropathy or enteropathy, or due to decreased production associated with the acute phase reaction. A normal or elevated serum albumin:globulin ratio can help rule out FIP as it is often decreased in affected cats. Acute Phase Proteins Acute phase proteins are a class of proteins whose plasma concentrations increase or decrease in response to inflammatory disorders. Levels of alpha‐1‐acid glycoprotein, an acute phase protein, of >1.5 g/L in plasma or effusions are suggestive of FIP in cats with compatible clinical signs. Effusion Fluid Effusions from the abdomen or pleural space are typically clear, straw‐colored, or viscous due to the high protein content. Rivalta’s test can differentiate transudate from exudate (www.abcdcatsvets.org/). A test tube is filled with 7–8 mL of distilled water and one drop of acetic acid (98%) and mixed thoroughly. One drop of the effusion fluid is layered onto the surface of this solution. Diffusion of the drop indicates a negative result, and therefore most likely indicates a FIP‐negative animal. If the drop floats slowly down in a drop or jellyfish‐like shape or stays at the surface, the Rivalta’s test is positive and the fluid may be considered an exudate. Fluid analysis is usually high protein (>3.5 g/dL) with a low nucleated cell count consisting primarily of neutrophils and macrophages, although findings may be variable. An effusion albumin:globulin ratio below 0.4 is suggestive of FIP. Serology Although different assays that detect antibodies directed against FCoV are commercially available, it is not possible to distinguish between exposure to FECV and FIPV infection. High antibody titer in serum does not always correlate with occurrence of FIP, and titers may be negative in cats with FIP. It has been suggested that many cats without FIP and who would never develop FIP have been needlessly euthanized due to a positive FCoV titer. Reverse Transcriptase Polymerase Chain Reaction Reverse transcriptase polymerase chain reaction (RT‐ PCR) assays can detect FCoV in a variety of samples

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(feces, blood, effusion, cerebrospinal fluid [CSF], tissue, and saliva) with high sensitivity. However, as with antibody tests, distinction of FECV from FIPV is not possible at this time. The finding of virus in blood or tissue also does not confirm FIP because FECV may be amplified from these tissues in cats without FIP. A quantitative PCR test for blood, effusion, and tissue is available. The test is based on the hypothesis that replicating virus in peripheral blood and tissues suggests FIPV rather than FECV. However, because viremia also occurs with FECV infection, healthy cats may also test positive using this assay. Histopathology and Immunostaining Relatively distinctive inflammatory infiltrates associated with FIP are characterized by variable degrees of severity and contain a combination of macrophages, lymphocytes, and plasma cells, mixed with lesser numbers of neutrophils. The hallmark of the histopathologic lesions is a granulomatous to pyogranulomatous reaction with vascular orientation and vasculitis. Detection of intracellular FCoV antigen in macrophages in effusions by immunofluorescence, or in tissue by immunohistochemistry, can confirm the diagnosis, and is considered the gold standard if properly performed. Immunostaining may be falsely negative if virus is not present in a given sample or if antibodies do not cross‐react with a particular strain.

­Therapy Supportive care and reducing stress are important. Specific treatment has been primarily focused on reducing the inflammatory and hyperimmune response to the virus. Unfortunately, increased survival has not been proven with any of these interventions. Of the immunosuppressants, prednisolone at immunosuppressive doses is currently the most consistently recommended. Although prednisolone may induce remission in some cats, treatment is not curative and may only slow the progression of the disease. Other immunosuppressive drugs such as chlorambucil have been used in combination with prednisolone but the risk:benefit ratio and efficacy are unknown. Feline interferon‐omega treatment is an immunomodulatory therapy that may be considered to treat FIPV‐infected cats, though studies show differences in efficacy. Similarly, a recent study showed that the immunostimulant polyprenyl extended the life of

three cats with mild dry FIP, with two cats still alive after two years of the diagnosis. However, follow‐up studies suggested no benefit in cats with more severe forms of the disease and its use is not recommended by some authors. Recent studies by Pedersen et al. have focused on antiviral therapy. Initially, the investigators performed a study using a 3C protease inhibitor (GC376) which showed some success in treating cats with FIP. Although 19 of 20 cats recovered, disease relapse occurred and 18 months after infection only six cats remained disease free. In another study, a small molecule nucleoside analog, GS441524, was used in 10 infected cats who were successfully treated and healthy eight months after treatment, with two cats requiring a second treatment after initial relapse. This nucleoside analog works by acting as an alternative substrate for the viral RNA polymerase terminating replication. These studies are promising developments for future clinical trials and treatments.

­Prognosis With the development of FIP, prognosis is poor to grave, with a reported median survival time of 49 days. Euthanasia should be considered especially when quality of life is poor and there is no response to therapy within a short period of time. However, newer research indicates that antiviral therapy might be successful in cats with complete reversal of clinical signs (see earlier).

­Prevention The formation of antibody enhances disease, so development of an effective vaccine is difficult. At the time of writing, the available vaccine is not recommended by the American Association of Feline Practitioners. Reducing stress and overcrowding, cleaning litterboxes daily, avoiding fecal–oral contact by keeping litterboxes away from food and water dishes, and separating newly acquired cats and cats suspected of being infected are recommended husbandry practices.

­Public Health Implications Feline coronaviruses are not thought to infect humans.

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­Further Reading Drechsler Y, Alcaraz A, Bossong FJ, Collisson EW, Diniz PPVP. Feline coronavirus in multicat environments. Vet Clin North Am Small Anim Pract 2011; 41(6): 1133–69. Murphy BG, Perron M, Murakami E, et al. The nucleoside analog GS‐441524 strongly inhibits feline infectious peritonitis (FIP) virus in tissue culture and experimental cat infection studies. Vet Microbiol 2018; 219: 226–33. Pederson NC. An update on feline infectious peritonitis: virology and immunopathogenesis. Vet J 2014; 201: 123–32.

Pederson NC. An update on feline infectious peritonitis: diagnostics and therapeutics. Vet J 2014; 201: 133–41. Pedersen NC, Kim Y, Liu H, et al. Efficacy of a 3C‐like protease inhibitor in treating various forms of acquired feline infectious peritonitis. J Feline Med Surg 2017; 20: 378–92.

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87 Feline Leukemia Virus David S. Bruyette, DVM, DACVIM (SAIM) Anivive Lifesciences, Long Beach, CA, USA

­Etiology/Pathophysiology Like all retroviruses, feline leukemia virus (FeLV) is an enveloped RNA virus. It carries an enzyme (reverse transcriptase) that transcribes the viral RNA genome into DNA (the FeLV provirus), which is then integrated into the host’s cell genome. Exogenous FeLV isolates are grouped into four major subcategories: FeLV‐A, ‐B, ‐C, and ‐T. The dominant FeLV‐A subgroup is horizontally transmitted and is found in all FeLV‐infected cats; it is the most infectious but low in pathogenicity. FeLV‐B and particularly FeLV‐C are less prevalent. They arise within the individual cat after FeLV‐A infection: FeLV‐B develops via recombination of FeLV‐A with endogenous FeLV‐related sequences and FeLV‐C and FeLV‐T evolve from FeLV‐A via mutations or insertions in the surface glycoprotein gene. Feline leukemia virus is shed in large quantities via saliva, but it can also be found in feces, urine, and milk. FeLV is unstable in the environment and transmission is thought to require intimate friendly (or aggressive) contact between infected and uninfected cats. Contact with infectious saliva or, less probable, infectious feces may also be sufficient to transmit the infection (sharing of food bowls or litterboxes). In addition, FeLV can be transmitted vertically from the queen to her kittens. Feline leukemia virus infection usually starts at the mucosa of the oropharynx. Subsequently, viral replication takes place in the adjacent tonsils and local lymph nodes. The virus is spread in the lymphoid tissues throughout the body via lymphocytes and monocytes. Replication in the bone marrow and infection of neutrophil and platelet precursors favor the initiation of secondary viremia and systemic infection. Cats that undergo bone marrow infection usually develop progressive infection. In cats that do not undergo bone marrow infection, only lymphocytes are provirus positive and

usually no viral RNA is detectable in peripheral blood cells. In addition, some cats seem to abort infection prior to provirus integration and an antibody response may be the only sign of previous FeLV exposure. After FeLV exposure, some, mostly young, cats develop progressive FeLV infection, characterized by persistent antigenemia/viremia. Cats with progressive infection shed high copy numbers of FeLV and pose an infection risk to other cats. Progressive infection is usually confirmed by repeated testing for antigenemia of the cat several weeks or a few months apart; only repeated positive antigen testing verifies the presence of a progressive infection, which is linked to a poorer prognosis than regressive FeLV infection. Cats with progressive FeLV infection are at high risk to die within a few months to years with FeLV‐associated diseases (see Prognosis). If a cat tests positive for FeLV p27 antigen, but upon later retesting becomes FeLV antigen negative, it has developed a regressive FeLV infection. Cats with regressive infection have mounted an effective immune response and recovered from FeLV viremia. Fortunately, most adult cats are found to develop regressive infections. Cats with regressive infection usually do not develop FeLV‐associated disease. Clearance of antigenemia is observed mostly within a few (2–8) weeks, but in rare cases it can also occur after many months although the chance for recovery decreases with increasing time. Following recovery from antigenemia (regressive infection), latent infection can be identified for a few months to years by culturing bone marrow cells in the presence of corticosteroids. Provirus polymerase chain reaction (PCR) from peripheral blood or bone marrow is positive in these cats and viral plasma RNA detected by reverse transcriptase (RT)‐PCR may be positive. Infection may be reactivated in latently FeLV‐infected cats. The potential for reactivation seems to be associated with the FeLV isolate and generally decreases with increasing timespan.

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­Epidemiology The prevalence of FeLV infection documented in cross‐ sectional surveys in North America has been declining throughout the past three decades and is attributed to testing and vaccination efforts. In the United States, 3.1% of cats in a large, nationwide dataset tested positive for FeLV in 2010, with increased risk among outdoor cats, unneutered males, and cats with other disease conditions (particularly respiratory disease, oral disease, and abscessation). Prevalence was highest in the midwestern and western regions of the United States and lowest in the northeast. Seroprevalence surveys of varying statistical power have found rates of positive test results to range from 3.6% in Germany and Canada to 4.6% in Egypt and 24.5% in Thailand. In a national (United States) study, FeLV infection was diagnosed in 9% of cats undergoing treatment for bite wounds, approximately three times the rate for cats in general. FeLV is considered to be an age‐dependent disease; young kittens are at higher risk of progressive infection and more rapid disease progression, whereas adults display some degree of age resistance. However, transmission can occur at any age, and factors affecting clinical course of disease are complex and incompletely understood.

­Signalment The susceptibility of domestic cats for FeLV infection varies and is mainly age dependent. Kittens are more prone to develop progressive FeLV infection than adult cats.

­History and Clinical Signs Feline leukemia virus‐related disorders are numerous and include anemia, neoplasia, immunosuppression, immune‐mediated diseases, reproductive problems, enteritis, neurologic dysfunction, and stomatitis. Some FeLV‐B‐infected cats develop lymphoid malignancies, while FeLV‐C infection is associated with the development of aplastic anemia. FeLV‐T is a T cell tropic cytopathic virus that causes lymphoid depletion and immunodeficiency in infected cats. The anemia caused by FeLV is typically nonregenerative and normochromic. Less commonly, macrocytosis or regenerative hemolytic anemia is seen in 10% of FeLV‐induced anemia cases. The cause of nonregenerative anemia is usually bone marrow suppression due to viral infection of the hematopoietic stem cells and the

s­ upporting stromal cells. Platelet dysfunction, thrombocytopenia, and neutropenia are all possible sequelae. Lymphoma is the most frequently diagnosed malignancy of cats. Tumors such as lymphoma and lymphoid leukemia develop in as many as 30% of cats with progressive FeLV infections. Regressive infections are also implicated in the occurrence of these tumors in the absence of viremia, but cats with progressive infections may face an increased risk of lymphoma development as high as 60‐fold. Most American cats with mediastinal, multicentric, or spinal forms of lymphoma are FeLV positive. However, these forms of lymphoma are becoming less common as the prevalence of FeLV decreases. Diffuse gastrointestinal (GI) lymphoma (small cell; T cell origin) is now more likely to be found in FeLV‐negative cats of middle or older age and can be difficult to differentiate from inflammatory bowel disease. Fibrosarcomas and quasi‐neoplastic disorders such as multiple cartilaginous exostoses (osteochondromatosis) can be associated with FeLV. Leukemia is characterized by the neoplastic proliferation of hematopoietic cells originating in the bone marrow, including neutrophils, basophils, eosinophils, monocytes, lymphocytes, megakaryocytes, and erythrocytes. Feline leukemias are strongly associated with FeLV infection and typically involve neoplastic cells circulating in the blood. Lymphoid leukemias are further classified as acute and chronic. Acute lymphocytic leukemia is characterized by lymphoblasts circulating in the blood, whereas chronic lymphocytic leukemias have an increased number of circulating lymphocytes with mature morphology. The immunosuppression caused by FeLV creates increased susceptibility to bacterial, fungal, protozoal, and viral infections. Numbers of neutrophils and lymphocytes in the peripheral blood of affected cats may be reduced, and those cells that are present may be dysfunctional. Many FeLV‐positive cats have low concentrations of complement which may contribute to FeLV‐associated immunodeficiency and oncogenicity, as complement is vital for antibody‐mediated tumor cell lysis. Immune complexes formed in the presence of moderate antigen excess can cause systemic vasculitis, glomerulonephritis, polyarthritis, and a variety of other immune disorders. In FeLV‐infected cats, immune complexes form under conditions in which FeLV antigens are abundant and anti‐FeLV IgG antibodies are sparse, a situation ideal for the development of immune‐mediated disease. Reproductive problems are commonly associated with FeLV infection. Fetal death, resorption, and placental involution may occur in the middle trimester of pregnancy, presumably as a result of in utero infection of fetuses by virus transported across the placenta in maternal leukocytes. Abortion typically occurs in late g­ estation.

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Transmission during birth and nursing constitutes the greatest risk of producing live, viremic kittens. There is some evidence that regressively infected queens may pass virus on to their kittens either in utero or in milk. Neonatal kittens are at risk of rapidly progressive infection with clinical manifestations of hypothermia, dehydration, failure to nurse, and early mortality, collectively termed “fading kitten syndrome.” It is likely that transmission from infected queens to their kittens is the single greatest source of FeLV infections. Although neurologic disorders associated with FeLV are most often caused by compression of the brain and spinal cord by lymphoma tumor tissue, a mechanism for neuropathology is also suspected to result in peripheral neuropathies, urinary incontinence, and ocular pathology, including anisocoria, mydriasis, Horner syndrome, and central blindness even in the absence of visible compressive lesions on diagnostic imaging. If antineoplastic therapy is planned, it is important to distinguish neoplasia from neuropathy. Stomatitis is more classically associated with feline immunodeficiency virus (FIV) infection, but FeLV infection can also predispose cats to chronic ulcerative proliferative gingivostomatitis. Clinical sequelae include pain, anorexia, and tooth loss. An immune‐mediated mechanism is likely, particularly in combination with co‐infections such as feline calicivirus.

found to be FeLV provirus positive and FeLV ­antigen negative. Provirus PCR testing is positive sooner (1–2 weeks) after FeLV exposure than antigen detection in blood. Some specialized laboratories offer RT‐PCR for the detection of viral RNA. The detection of viral RNA in the saliva correlates well with the detection of antigen in the blood of infected cats. However, FeLV viral RNA in saliva and blood of experimentally infected cats was found as early as one week after FeLV exposure, at least two weeks before antigen detection in the blood. Testing Cats for FeLV Infection ●●

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­Diagnosis The outcome of a FeLV infection can be characterized using the results from virus isolation, immunofluorescence assays, detection of FeLV p27 capsid antigen in serum or plasma, and detection of FeLV provirus or viral RNA in the blood or some tissues by molecular assays. Serologic assays for the detection of FeLV‐specific antibodies are currently not routinely used. Antigenemia (presence of p27 capsid antigen) is a marker of infection and in most, but not all cats, a parameter for viremia (presence of replication‐competent virus). Detection of viremia by virus isolation is not widely available, laborious, and time consuming. In contrast, antigenemia can be easily detected using enzyme‐linked immunosorbent assay (ELISA) in a specialized laboratory or by rapid point‐of‐care tests. Most cats will test antigen positive within 3–6 weeks after FeLV exposure. The detection of FeLV provirus in the peripheral blood or certain tissues like bone marrow using RT‐PCR is more sensitive than the detection of antigenemia or viremia to demonstrate previous FeLV exposure. Thus, a certain proportion of pet cats can be found to be PCR positive but antigen negative. In an early study in Switzerland, 10% of pet cats were

●●

Test sick cats for the presence of FeLV antigenemia/ viremia (using antigen tests, virus isolation, RT‐PCR from saliva). –– Any cat that tests FeLV positive: retest the cat after 1–2 months to determine whether progressive or regressive infection prevails. Quarantine the cat during this period since the cat is a FeLV infection source for other cats. This procedure may be repeated if circumstances permit and/or the testing interval may be discussed with the cat owner. Test healthy cats for the presence of FeLV antigenemia/viremia (using antigen tests, virus isolation, RT‐PCR from saliva). –– Prior to the first FeLV vaccination, testing is recommended for every cat; prior to the second vaccination (booster of basic immunization), testing is recommended if the cat may have been exposed to FeLV in the last 4–8 weeks. –– If the cat is at high risk for FeLV (outdoor access, multicat environment, any husbandry with changing cats, etc.). –– If the cat has moved to a new environment (new home, shelter, cattery, etc.): if the cat tests negative, quarantine (keep from other cats and any potential FeLV exposure) and retest after 4–6 weeks for absence of FeLV antigen prior to introduction into the new environment. –– Any cat that tests FeLV positive: retest the cat after 1–2 months to determine whether progressive or regressive infection prevails. Quarantine the cat during this period since the cat is a FeLV infection source for other cats. This procedure may be repeated if circumstances permit and/or the testing interval may be discussed with the cat owner. Test healthy cats for the presence of FeLV provirus in the blood (PCR). Confirmatory of positive or questionable antigen tests; true antigen‐positive samples are highly provirus positive. –– Early infection: provirus PCR from blood tests ­earlier positive than antigen tests.

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–– If prior exposure to FeLV needs to be known: ­analysis of the FeLV situation in a cat colony with FeLV problems. –– If the cat is used as blood or organ donor: FeLV can be transmitted by transfusion of FeLV provirus‐positive blood. Antigen testing is not sufficient. Test sick cats for the presence of FeLV provirus in the blood (PCR). Confirmatory of positive or questionable antigen tests; true antigen‐positive samples are highly provirus positive. –– Early infection: provirus PCR from blood tests ­earlier positive than antigen tests. –– If prior exposure to FeLV needs to be known: analysis of obscure clinical cases, analysis of the FeLV situation in a cat colony with FeLV problems, etc. Test any cat for the presence of FeLV viral RNA in the blood (RT‐PCR). Early infection: RT‐PCR tests earlier positive than provirus PCR and antigen tests. Not frequently used.

Kittens can be tested any time for antigenemia – maternal antibodies do not interfere with the p27 testing. A confirmed positive result is conclusive, while a negative result is not always; in some cases the kittens became positive only after weeks to months. Each kitten should be tested individually.

­Treatment Unfortunately, no curative treatment currently exists to cure retroviral infection. In vitro studies have yielded promising results suggesting virus‐suppressing activity of human FDA‐approved drugs used to treat HIV and other myelodysplastic syndromes against FeLV virus (e.g., raltegravir, tenofovir, gemcitabine, decitabine). Further research is needed to demonstrate efficacy and safety in vivo and in field trials, as well as to address affordability of these drugs for most cat owners. Feline interferon‐omega and human interferon‐alpha have been associated with improved survival, but concerns surrounding availability, cost, and absence of strong evidence in controlled field studies have limited their widespread integration into standard treatment protocols for FeLV. Anecdotal reports of various antiviral and immunotherapeutic agents to reverse viremia, improve clinical signs, and prolong survival are abundant. Controlled studies using naturally infected cats have either not been performed or have not confirmed anecdotal observations. Treatment efficacy must be demonstrated in ­controlled clinical trials, because spontaneous reversion to seronegative status or prolonged survival is not

uncommon, even in the absence of medical treatment. Some FeLV‐positive cats can live without major disease complications for years with routine prophylactic care, good husbandry, minimal stress, and avoidance of secondary infections. Infected cats should be kept strictly indoors to reduce the risk of exposure to infectious agents and prevent transmission of the virus to other cats. Physical examinations focusing on external parasites, skin infections, dental disease, lymph node size, and body weight should be performed semiannually, along with a routine program for parasite control and annual fecal, complete blood count, chemistry panel, and urinalysis testing. All infected cats should be neutered. Owners should be advised to watch for signs of FeLV‐ related disease, particularly secondary infections. Although FeLV‐positive cats often respond well to treatment for secondary infections, therapy for such infections or other illnesses should be early and aggressive because of immunocompromise. Because FeLV is historically associated with rapid and grave disease, the modern prognosis varies considerably depending on husbandry, veterinary care, and individual immune system variation.

­Prognosis Large‐scale studies have demonstrated an average survival of 2.4 years after diagnosis among positive cats (versus six years after testing for negative control cats), with 50% mortality in two years and 80% mortality by three years after diagnosis. Progression of disease is much more rapid in kittens, whereas some adult cats remain healthy for many years and may succumb to conditions unrelated to their retroviral status.

­Prevention and Control Feline leukemia virus is unstable in the environment and is susceptible to all common detergents and disinfectants. In a hospital or boarding setting, infected cats may be kept in the general population as long as they are housed in separate cages. Medical and surgical equipment contaminated with body fluids, even when dried, can be fomites for infection. Thorough cleaning and sterilization of equipment, strict attention to washing contaminated hands, and avoiding reuse and sharing of single‐use and consumable supplies between patients are critical practices to prevent iatrogenic transmission. Feline leukemia virus vaccines are noncore and are intended to protect cats against FeLV infection or reduce the likelihood of persistent viremia. Types of vaccines include killed whole virus, subunit, and genetically

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e­ ngineered. Vaccines are indicated only for uninfected cats, because there is no benefit in vaccinating an FeLV‐ positive cat. Vaccination is the most effective way to prevent FeLV‐associated disease. Currently, FeLV vaccines are very safe and have up to 98% efficacy. The American Association of Feline Practitioners (AAFP) Feline Retrovirus Management Guidelines include the recommendation that all kittens should receive the two‐dose FeLV vaccination as a component of the routine initial vaccination series and should also receive a booster vaccination one year later. This is prudent, because rehoming and lifestyle changes such as outdoor access frequently occur as cats mature. Annual revaccination after maturity would depend on the cat’s risk of FeLV exposure. The adult cat’s risk of exposure to FeLV‐positive cats should be assessed, and vaccines used only for those cats at risk. FeLV vaccines have been associated with development of sarcomas at the vaccination site, although the risk of tumor development is very low. Uninfected cats in a household with infected cats should be vaccinated; however, vaccination is not universally protective, and other means of reducing transmission to uninfected cats, such as physical separation, should also be used.

While testing of cats in an animal shelter environment is considered optional for individual housing, FeLV ­status should be determined before placement in group housing and is recommended at the time of adoption or foster home placement. Because tests are not 100% accurate, shelter cats placed in group housing should be vaccinated against FeLV, especially in long‐term conditions such as sanctuaries. Because of the equivalent prevalence of FeLV among feral and free‐roaming pet cats and the role of neutering in decreasing the spread of infection, expending resources on FeLV testing is not considered a mandatory component of community trap‐neuter‐ return programs.

­Public Health Implications Some strains of FeLV can be experimentally grown in human tissue cultures, leading to concerns of potential for transmission to people. Studies addressing this concern have shown no evidence that any zoonotic risk exists, and there are no known cases of zoonotic transmission.

­Further Reading Hartmann K. Clinical aspects of feline retroviruses: a review. Viruses 2012; 4(11): 2684–710. Levy J, Crawford C, Hartmann K. American Association of Feline Practitioners feline retrovirus management guidelines. J Feline Med Surg 2008; 10: 300–16. Levy JK, Crawford PC, Tucker SJ. Performance of 4 point‐of‐care screening tests for feline leukemia virus and feline immunodeficiency virus. J Vet Intern Med 2017; 31: 521–6.

Nesina S, Helfer‐Hungerbuehler K, Riond A, et al. Retroviral DNA – the silent winner: blood transfusion containing latent feline leukemia provirus causes infection and disease in naïve recipient cats. Retrovirology 2015; 12(1): 105. Westman ME, Malik R, Hall E, et al. Comparison of three feline leukaemia virus (FeLV) point‐of‐care antigen test kits using blood and saliva. Comp Immunol Microbiol Infect Dis 2017; 50: 88–96.

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88 Feline Immunodeficiency Virus Tom Phillips, DVM, MS, PhD, † College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA, Deceased

­Etiology/Pathophysiology Feline immunodeficiency virus (FIV) is a member of the Lentivirus genus in the Retroviridae virus family. As such, it is an enveloped virus, has an icosahedral capsid, and a genome composed of two (nearly identical) linear, positive sense, single‐stranded RNA molecules. Much has been learned about FIV since it was first described in 1987, particularly in regard to its applica­ tion as a model to study the closely related lentivirus human immunodeficiency virus (HIV). In particular, FIV and HIV share remarkable structure and sequence organization, utilize parallel modes of receptor‐medi­ ated entry, and result in a similar spectrum of immuno­ deficiency‐related diseases due to analogous modes of immune dysfunction. The disease course for FIV is variable, likely dependent upon the route of infection, strain of virus, age of the ani­ mal at time of infection, and the robustness of the immune response generated to the virus. Transmission is primarily through the bite of an infected cat, though sexual transmission has been shown to occur in experi­ mentally infected cats. Infection by blood transfusion may also potentially occur. Initially, FIV is transported by tissue macrophages to lymphoid tissues, where it repli­ cates in lymph nodes, thymus, spleen, bone marrow, brain, lung, gastrointestinal tract, and kidney. FIV gains entrance into the cell primarily through receptor media­ tion. The main cellular receptors for FIV are the chemokine receptor, CXCR4, and CD134, a receptor expressed by T lymphocytes and activated macrophages. Other mechanisms of virus entry may also be important, such as the use of other receptors and antibody‐medi­ ated entry through FC receptors. Experimentally, virus is detected in the blood of animals usually by two weeks post infection, and remains easily

detectable for up to six weeks post infection. Generally, FIV induces a strong but only partially ­effective immune response. However, there are reports of detectable levels of FIV DNA in cats without a detectable antibody response. Generally, antibodies to FIV are detected by two weeks post infection, and most cats develop antibody by 60 days post infection, although it may be delayed further in some cats. Over the next few weeks after antibody development, the virus levels in the blood are gradually reduced to undetectable quantities and become clinically latent. However, the virus is not truly latent, as it contin­ ues to replicate at low levels. During this clinical latent phase, the virus infection gradually is eroding normal immune function of the infected cat. FIV causes the T helper cell numbers to decrease with an inversion of the CD4:CD8 ratio. The effect of FIV on feline lymphocyte numbers is likely due to several mechanisms operating simultaneously: virus‐induced lysis of infected cells, virus suppression of the ability of bone marrow and other lym­ phoid tissues to produce new lymphocytes, immune‐ mediated lysis of infected cells, and apoptosis. The ability of lymphocytes from FIV‐infected cats to proliferate in response to both B and T cell mitogens is impaired, as is the expression of lymphocyte cell surface receptors that are important in control and proliferation of the immune response, such as cytokine receptors and MHC class II antigens. The pathophysiology of FIV neurologic dysfunction is not completely understood as it occurs in the apparent absence of virus replication in neurons, with minimal inflammatory lesions in infected cat brains, and only low levels of replication in astrocyte and microglia cells. Infected astrocytes have impaired glutamate uptake function, which may lead to an increased susceptibility of glutamate toxicity, oxidative stress, and ultimately neuron loss. Additionally, FIV envelope protein is toxic

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to rat neurons in culture and may play a role in neuro­ logic disease pathogenesis. The neurologic disease is more prevalent with certain isolates of the virus, which may point to certain envelope protein sequences being more toxic and/or relate to the ability of virus to replicate in the feline nervous system.

­Epidemiology Feline immunodeficiency virus infection is common around the world. The incidence of FIV varies (from 4% to 47%) geographically, correlating with the percentage of free‐roaming and feral cats in the population.

­Signalment Although it is possible for any cat to be infected with FIV, free‐roaming, nonpure‐bred, sexually intact, mature, male cats are more likely to contract an FIV infection. This is largely due to cat bites being the primary mode of virus transmission.

­History and Clinical Signs Acute clinical signs of the infection are generally mild and often consist of the nonspecific signs of lethargy, anorexia, pyrexia, diarrhea, and lymphadenopathy. Generally, these clinical signs coincide with the viremia. Once the viremia subsides, the virus appears to go into a clinical latent phase, with virus not detected in the blood and the apparent absence of most clinical signs. This clinical latent phase is of variable length but often lasts for years, as the virus slowly suppresses the immune system. The last phase of the infection is feline acquired immune deficiency syndrome (FAIDS). Clinical signs of FAIDS are the result of opportunistic infections, various neoplasias, and/or neurologic dis­ ease. Thus, clinical signs are variable depending upon how the immune suppression is manifested. Stomatitis is very common and can occur during any phase of the infection. Persistent diarrhea is also fairly common, as is respiratory disease. Neoplasia is common in FAIDS, with B cell lymphoma being the most frequent. Neurologic signs such as behavioral changes, disrupted sleep, impaired learning, paresis, and seizures have been reported. A wasting condition has also been described late in the FAIDS phase of the disease. Increased circulating immune complexes can result in immune complex disease manifested as uveitis and glomerulonephritis. ­

­Diagnosis Complete blood count (CBC) abnormalities may include anemia, lymphopenia, and neutropenia. Thrombocy­ topenia may sometimes occur. Hyperglobulinemia is the most common biochemical abnormality. Proteinuria may be seen in cats with glomerulonephritis. Antibodies to FIV infection persist for life and thus patient‐side ELISA antibody testing is recommended as a screen for FIV infection. However, false‐positive and ‐ negative results may occur. If a kitten under 6 months of age is antibody positive, it should be retested after 60 days, as maternal antibody could be the cause of the pos­ itive assay. If an animal is negative after a recent possible exposure, it should be retested after 60 days, allowing sufficient time for an active immune response to develop. Tests used to detect antibody cannot distinguish between infection and vaccination. Positive tests in healthy cats should be confirmed using ELISA from another manu­ facturer, western blotting, or PCR. In cases where the vaccination history is not known, real‐time quantitative PCR should be considered, as this assay can detect a low‐ grade viremia, indicative of an active infection. However, sensitivity and specificity of PCR are variable. PCR may be negative due to low levels of viremia or mutations, and false‐positive results have been reported in vacci­ nated cats.

­Management The best therapy for FIV is prevention. Keeping cats indoors and only allowing exposure to known FIV‐nega­ tive cats is the best way to prevent the infection. The vac­ cine for FIV is controversial, as efficacy is variable. Additionally, using standard FIV screening assays, a vac­ cinated cat currently cannot be distinguished from an infected cat. Since shelters may not perform a confirma­ tory assay on an animal that is found FIV positive by a screening assay, it is advised to microchip the cat at the time of vaccination with a chip whose data are linked to vaccination status to assist shelter personnel in the inter­ pretation of a positive result. The use of the routine feline vaccines in FIV‐positive cats is controversial due to the suppressed immune system of the FIV‐infected cat. Since it is best to keep infected cats indoors to reduce the risk of FIV exposure to other cats, the exposure of the infected cat to other common patho­ gens is also reduced, thereby decreasing the need for vac­ cination. Additionally, the duration of immunity to most of the vaccinatable feline pathogens likely lasts for years. If the cat has been vaccinated earlier in life, protective immu­ nity likely still remains. If, however, vaccines are to be used,

88  Feline Immunodeficiency Virus

killed vaccines are preferred. Even if they are kept indoors, housing cats in overcrowded conditions has been signifi­ cantly associated with disease progression in FIV‐positive cats. Therefore, reducing stress and limiting exposure to potential infectious agents are important husbandry prac­ tices that can decrease morbidity and mortality. For cases of prolonged severe stomatitis, removal of all the teeth is the treatment of choice. Zidovudine (AZT) at a dose of 5–10 mg/kg PO or SC q12h has also been shown to be effective in some cases of stomatitis. AZT is also used, to some effect, in cases with neurologic signs of dis­ ease. AZT can cause bone marrow suppression. CBCs should be monitored closely. AZT should not be used in cats with myelosuppression. Alfaferone (natural human alpha‐interferon) has been reported to increase the sur­ vival of FIV‐infected cats. Recombinant feline IFN‐omega may improve clinical signs and reduce inflammatory cytokine production but does not appear to affect viral load. A recent publication demonstrated that an analog of tenofovir, a widely prescribed anti‐HIV drug in human medicine, may also be effective as a therapy for FIV, though further investigation is needed. Griseofulvin treatment

of fungal disease should be avoided as bone marrow sup­ pression has been reported in FIV‐infected cats.

­Prognosis Though the virus can never be cleared from an infected cat, most infected cats have long life spans. While many die from conditions that are unrelated to the infection, many ultimately succumb to an FIV‐related disease.

­Public Health Implications Feline immunodeficiency virus infection of humans has never been documented, even in veterinarians or indi­ viduals who have been bitten multiple times by FIV‐ infected cats. However, FIV has been shown to replicate in human cells in vitro and cause disease in monkeys experimentally infected with FIV‐infected autologous cells. Thus, caution is still advised.

­Further Reading de Oliveira Medeiros S, Abreu CM, Delvecchio R, et al. Follow‐up on long‐term antiretroviral therapy for cats infected with feline immunodeficiency virus. J Feline Med Surg 2016; 18(4): 264–72. Hartmann K, Wooding A, Bergmann M. Efficacy of antiviral drugs against feline immunodeficiency virus. Vet Sci 2015; 2(4): 456–76. Levy J, Crawford C, Hartmann K, et al. American Association of Feline Practitioners’ feline retrovirus

management guidelines. J Feline Med Surg 2008; 10(3): 300–16. Miller C, Abdo Z, Ericsson A, et al. Applications of the FIV model to study HIV pathogenesis. Viruses 2018; 10(4): ii. Sahay B, Yamamoto JK. Lessons learned in developing a commercial FIV vaccine: the immunity required for an effective HIV‐1 vaccine. Viruses 2018; 10(5): ii.

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89 Feline Viral Upper Respiratory Tract Disease Yvonne Drechsler, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology

­Epidemiology

Feline viral infectious upper respiratory tract disease is primarily caused by two viruses: feline herpesvirus (FHV‐1) and feline calicivirus (FCV). Infection is common in multicat environments, and most cats have been exposed. FHV‐1 is an enveloped single‐stranded DNA virus belonging to the alpha‐herpesvirus family. As such, the virus exists in a latent form in the trigeminal ganglia after initial infection and can be reactivated by stressors. This leads to shedding via secretions from oral, nasal, and conjunctival membranes. Viral replication is usually restricted to the upper respiratory tract, due to viral preference for lower temperatures. However, in young kittens and possibly immunosuppressed cats, more severe systemic disease can occur. Feline calicivirus belongs to the Caliciviridae family, and consists of a single‐stranded positive RNA genome. This genome structure has a high mutation rate and therefore the isolates are highly variable. Strains can vary in their pathogenicity. Like FHV‐1, transmission occurs via secretions from oral, nasal, and conjunctival membranes. FCV persists in tonsillar and other oropharyngeal tissues, leading to a carrier state, but the mechanism of persistence is not well understood. In addition to causing upper respiratory disease, FCV can be associated with lameness due to acute synovitis, possibly due to antigen‐activated macrophages residing in the synovial membrane. Rarely, FCV can cause virulent systemic disease (VSD). Lesions associated with VSD are severe and include oral and dermal ulcerations, bronchointerstitial pneumonia, necrosis of the liver, spleen and pancreas, and death. The pathogenesis of VSD is not fully understood.

Infection with both FHV‐1 and FCV is widespread in the general cat population. The prevalence of infection and clinical disease is higher in multicat households, colonies, catteries or shelters. Viral reactivation may occur in latently infected cats when they are stressed or if their immune system is compromised. FHV‐1 is short‐lived in the environment, and transmission occurs mainly by direct contact with an infected cat. However, transmission through fomites in crowded environments is also important. Cats infected with FCV may be carriers who shed virus while appearing clinically healthy. Virus may be shed in the urine and feces in addition to respiratory secretions. FCV survives in the environment much longer than FHV‐1 and is resistant to quaternary ammonium compounds. Therefore, in addition to direct and aerosol contact with respiratory secretions, fomites are a very important means of transmission for FCV.

­Signalment Cats of any sex and age can develop respiratory disease caused by FHV‐1 or FCV, with kittens being more susceptible.

­History and Clinical Signs Cats from multicat households, colonies, catteries or shelters are commonly infected. A history of stress or immune suppression may be noted. FHV‐1 may cause  severe respiratory disease, especially in

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i­mmune‐­compromised animals. Nonspecific signs of FHV‐1 infection may include depression, anorexia, and pyrexia. Respiratory signs include sneezing and conjunctivitis associated with serous ocular and nasal discharge. Nasal turbinate destruction can lead to recurrent rhinitis and sinusitis. Coughing and dyspnea can develop in more severe cases. Ocular and nasal discharge that becomes mucoid or purulent may signal secondary bacterial infection. In addition to conjunctivitis, keratitis and corneal ulcerations also occur with FHV‐1 infection. Dendritic corneal ulcerations are considered pathognomonic for infection with FHV‐1. FHV‐1 might also play a role in other forms of ocular disease. Skin ulceration and herpetic dermatitis may also be caused by FHV‐1. Feline calicivirus causes a range of clinical signs due to strain variability. Similar to FHV‐1, upper respiratory signs predominate. Sneezing and ocular discharge associated with conjunctivitis are common. In contrast to FHV‐1 infection, corneal ulceration does not typically occur with FCV infection. However, oral ulceration is common, making FCV a top differential diagnosis for oral ulcerations in cats. Stomatitis may be severe, and lead to excessive ptyalism. Other signs of FCV infection include depression, pyrexia, anorexia, enlarged lymph nodes, and acute lameness. Infection with highly virulent strains of FCV causing VSD results in severe signs such as high fever, cutaneous edema, jaundice, other signs of multiorgan failure, and death.

­Diagnosis History, clinical signs, and evidence of possible immunosuppression (due to retroviral infection, stress or drug administration, for example) should increase the index of suspicion for infection with one or both viruses. For cats with signs of upper respiratory disease, diagnosis is usually made based on clinical signs alone. Specific laboratory tests documenting infection or exposure may be used to help confirm infection, particularly in patients with more severe or atypical clinical signs, although each has limitations. Complete blood count (CBC) and serum chemistry findings are not specific in cats with upper respiratory disease caused by FHV‐1 or FCV, although cats with stomatitis caused by FCV may be hyperglobulinemic. Cats with VSD may have anemia, thrombocytopenia, neutrophilia, and lymphopenia. Hypoalbuminemia, elevated alanine aminotransferase (ALT) and aspartate aminotransferase (AST), elevated bilirubin, and increased creatine kinase (CK) also occur. Direct testing for the presence of virus may include culture, polymerase chain reaction (PCR), histopatho-

logic examination, and immunofluorescence staining. False‐negative results may occur with any of these assays as virus may not be present in specimens. Laboratories should be contacted for appropriate sample acquisition and transport techniques for culture. The sensitivity and specificity of PCR may vary depending on the laboratory and design of the test. It should also be kept in mind that PCR may not detect all strains, particularly for calicivirus, due to variability in nucleic acid target sequences. Positive PCR results may occur in healthy cats, so a positive result does not necessarily mean that clinical signs may be attributed to infection. In addition, vaccination with attenuated virus may cause false‐positive PCR results. Serologic testing is also available, but it is not easily interpreted as vaccination and previous exposure are common for both viruses and the presence of antibody may not indicate active infection.

­Therapy In cases of acute respiratory disease, supportive care including fluid therapy and/or nutritional support may be indicated. Clinical signs resolve within 2–3 weeks. Judicious administration of appropriate antibiotics is indicated if evidence of secondary bacterial infection or co‐infection with Chlamydia, Mycoplasma or Bordetella is present. For more severe or recurrent disease, or if keratoconjunctivitis or stomatitis is present, more aggressive specific therapy may be indicated, including antiviral or immunomodulating agents, and for stomatitis, dental extractions, antimicrobial treatment, and antiseptic mouthwashes. Most antiviral therapies developed for human herpesviruses are too toxic for oral administration in cats, although the use of famciclovir appears promising. Topical use of idoxuridine, vidarabine, and other select nucleoside analogs appears beneficial for treatment for ocular disease. The therapeutic potential of feline interferon‐omega on FHV‐1 replication is under investigation. The use of oral L‐lysine to decrease clinical signs or infection with FHV‐1 infection has shown conflicting results.

­Prognosis In general, cats will recover but both viruses persist, with reactivation of FHV‐1 occurring due to stressors. FCV carriers can shed virus continuously, increasing the likelihood of infecting other cats. FCV infection with strains associated with VSD has been associated with greater than 50% mortality.

89  Feline Viral Upper Respiratory Tract Disease

­Prevention Vaccination information and guidelines are available for free online from the American Association of Feline Practitioners (https://catvets.com/guidelines/practiceguidelines/feline-vaccination). Vaccination, stress reduction, avoiding overcrowding and implementing strict disease prevention techniques in shelters and kennels are

important to prevent infection and associated clinical signs (see Chapter 117).

­Public Health Significance These viruses are not known to infect people.

­Further Reading Dawson S, Bennett D, Carter SD, et al. Acute arthritis of cats associated with feline calicivirus infection. Res Vet Sci 1994; 56(2): 133–43. Gould D. Feline herpesvirus‐1: ocular manifestations, diagnosis and treatment options. J Feline Med Surg 2011; 13(5): 333–46. Holland JL, Outerbridge CA, Affolter VK, Maggs DJ. Detection of feline herpesvirus 1 DNA in skin biopsy

specimens from cats with or without dermatitis. J Am Vet Med Assoc 2006; 229(9): 1442–6. Sykes JE. Feline respiratory viral infections. In: Canine and Feline Infectious Diseases. St Louis, MO: Elsevier, 2014, pp. 239–51.

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90 Rabies in Dogs and Cats Emily Beeler, DVM, MPH and Karen Ehnert, DVM, MPVM, DACVPM Veterinary Public Health Program, Los Angeles County Department of Public Health, Los Angeles, CA, USA

­Etiology and Pathophysiology The rabies virus is an enveloped, single‐stranded RNA virus in the family Rhabdoviridae and the genus Lyssavirus. There are 16 known lyssaviruses worldwide capable of causing rabies‐like symptoms in mammals. Bats are the reservoir for most of them. Rabies virus itself is by far the most prevalent lyssavirus, and the only one known to exist in the Americas. Rabies has the highest mortality rate of all known infectious diseases. Rabies is usually transmitted by the bite of a rabid animal. It may also be transmitted when virus‐laden saliva is introduced into mucous membranes. Theoretically, scratches may transmit the virus, if the claws are contaminated with fresh infectious saliva, but this mode of spread is very rare in people. Because the virus may be present in tissues throughout a rabid animal, exposure may occur during necropsy. Rabies has occasionally been spread by corneal and internal organ transplantation,  transplacentally, through aerosolization in closed spaces such as caves, and by ingestion of infected tissues. Because it is readily destroyed by UV light, desiccation, and most disinfectants, the rabies virus is not effectively transmitted by fomites. When the rabies virus first enters the body, it multiplies at a very low rate in a small amount of muscle cells and fibrocytes, and curtails the local immune reaction by inhibiting type I interferons. The majority of the incubation period, which can last for 3–12 weeks or more in dogs and cats, likely passes with the virus localized at the site of entry. The virus then enters the nervous system through a peripheral motor nerve, and is drawn up the axon at a rate of about 100 mm per day. Once the virus reaches the neuronal cell body, it multiplies and disseminates rapidly, although unevenly, throughout the central nervous system (CNS). Histologic analysis of brains from rabies victims typically reveals little tissue damage.

When increased inflammation and damage do occur in the CNS, they are usually found in patients in which paralytic clinical signs predominated. The total amount of virus in the brain tends to be significantly higher in cases of encephalitic, “furious” rabies than in cases of paralytic, “dumb” rabies. Some details of what happens at the molecular level of rabies pathogenesis remain a mystery. The rabies virus has an uncanny ability to evade early recognition and clearance by the immune system, and to preserve the health of cells of the nervous system throughout most of the infection. In laboratory studies, rabies virus strains that stimulate a significant immune response are less pathogenic than those that suppress it.

­Epidemiology Over 60 000 people die worldwide from rabies each year. The vast majority of rabies victims are children who were infected by bites from rabid dogs. Most cases occur in countries where rabies vaccination rates for dogs are below 50%, primarily in Asia and Africa. The human death toll from rabies is much lower in countries where animal control policies are well established and vaccination of pets is widespread. Over the past two decades in the United States, an average of three people per year died from rabies. Typically, the cases occurred after a person failed to obtain timely rabies postexposure prophylaxis, often due to lack of awareness about the risk of rabies. About 70%  of the cases were caused by bat bites, and most others  were caused by a dog bite inflicted while the person was visiting another country. The primary reservoirs for rabies are bats, domestic dogs, raccoons, skunks, foxes, mongooses, and jackals. In the past decade, 6000–7000 rabid animals per year

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were detected in the US. Over 90% were wild animals, with raccoons, bats, and skunks constituting over 80% of the total. Rabid foxes add another 5%. The island of Puerto Rico has consistently reported about 30–40 rabid mongooses each year. Rabid cats (4% of rabid animals) have outnumbered rabid dogs (1%) for many years, most likely because of better vaccination coverage in dogs. In North America, rabies cases are more frequent in spring and summer. The number of rabid animals reported typically reflects only those confirmed by laboratory testing; it is thought that the number of undiagnosed rabies cases in wild animals is far higher. There are multiple rabies virus substrains (aka variants), that are uniquely adapted to their reservoir hosts. Rabies variants tend to be found in identifiable geographic locations. This phenomenon is very useful for investigating the source of individual rabies cases. As an example, if a stray dog in the US was found to be rabid, analysis of the variant could reveal the source of the infection. Finding a North American bat, skunk, or raccoon variant in the dog would reveal that it caught the virus from indigenous wildlife. Finding a Thai dog variant would suggest the dog had recently been imported from Thailand, or had been bitten by a rabid dog from Thailand. In the Americas, insectivorous bat rabies variants are widely distributed. Vampire bat variants are found in certain areas in South America, Central America, and southern Mexico. The canine rabies virus variant has been eradicated from the US but is still present in parts of Central and South America. So far, no feline variant has been reported. The pathogenicity and virulence of the variant, the behavior of the reservoir species (natural behavior and behavior when rabid), and the degree of overlap in habitat between the reservoir species and other mammals all affect the risk of rabies transmission to pets and people. The risk of rabies exposure for domestic animals is higher in areas where variants associated with terrestrial animals (i.e., not just insectivorous bat variants) are prevalent. The risk is highest where the canine variant is enzootic (example: India), and much lower in areas where only insectivorous bat rabies variants are found (example: southern California). Many islands in the world, such as Hawaii, are considered completely rabies free. The raccoon variant is found throughout the eastern quarter of the US and in parts of eastern Canada. Multiple variants of skunk rabies are found in large areas in the center of North America, California, and parts of Mexico. Fox variants have been found in Alaska, Arizona, New Mexico, and Texas in the US, and in Ontario, Canada. One variant found periodically in southern Texas and northern Mexico appears to be adapted to both dogs and coyotes.

Humans have spread rabies variants in the US by moving infected animals. Before the late 1970s, the raccoon variant had been limited to the southeastern area of the US. It was translocated to the mid‐Atlantic states by people shipping raccoons from south to north for hunting purposes. The canine variant of rabies, which was considered officially eliminated from the US in 2007, could be reimported at any time from countries where canine rabies is still enzootic. Rabid dogs have been imported into the US many times but fortunately were quickly detected, isolated, and euthanized. If imported rabid dogs were released into populations of unvaccinated dogs, such as into a city with a significant stray dog population, canine rabies could become reestablished in the US. Mice, rats, and rabbits can be experimentally infected with rabies, but rabies is very rare in them in nature. Rabies has been detected in larger rodents in the US, such as woodchucks, groundhogs, and beavers, usually in areas where the raccoon variant is enzootic. It has also been rarely seen in rabbits attacked by raccoons. Opossums are partially resistant to experimental rabies infection. So far, no marsupial has been found to be a rabies reservoir. Nonetheless, opossums can become infected. In California (where skunk and bat rabies variants are enzootic), 11 rabid opossums were detected between 1983 and 2012 (data from Dr Curtis Fritz, California Department of Public Health). The omnivorous nature of opossums would make it natural for them to investigate a dying, rabid animal, and to potentially be bitten. Practitioners should have full understanding of the rabies variants that are common in their area. However, since rabies may be imported, a thorough travel history for all patients should also be taken, with special focus on the previous six months. The Centers for Disease Control and Prevention (CDC) maintains an online list of nations that present a higher risk for importation of rabies in animals.

­Signalment Any breed of dog or cat may present with rabies. Rabies is highly unlikely in animals less than 4 weeks of age.

­History Taking a thorough history is vital for clinical diagnosis. The risk of rabies depends on the geographic area where the pet lives or has spent time, especially in the previous six months. Knowledge of the clinical course of illness before presentation is also essential for diagnosis.

90  Rabies in Dogs and Cats

A history that increases the likelihood of rabies may include the following. ●●

●●

●●

●●

Travel. Pet imported within the last six months from a country where canine rabies is enzootic (includes many countries in Africa, Asia, and Central and South America). Imported strays or street animals present especially high risk. See the Public Health Implications section below for more information. Unvaccinated. No rabies vaccination in pet’s history. Dogs and cats that were vaccinated only once or twice against rabies may not be fully protected. Repeated vaccination (>2 times) significantly reduces the risk of rabies. Exposure to wild animal or suspected rabid pet. History of interaction with a wild animal (especially a known rabid animal or rabies reservoir species) 10 days to six months before presentation. Bite from the wild animal will likely be healed by the time rabies signs begin. Bites inflicted in thick fur or in the oral cavity, or bites from bats, may have gone unrecognized. Onset of clinical signs is gradual and steady. Once clinical signs begin, they worsen over days or hours. Animals are unlikely to be rabid if they have a peracute onset of neurologic signs, stabilization or improvement, or have survived through more than 10 days of overt illness.

Clinical signs in the prodromal phase are nonspecific. Dogs and cats may have fever, lethargy, dehydration, anorexia, vomiting or diarrhea. The animal may withdraw and hide, or may appear to become more sociable than usual. They may lick or chew at the site of the original bite wound. Activation of the autonomic nervous system may lead to piloerection, drooling, and dilated pupils. Furious (excitatory) phase clinical signs may include the following. ●●

●● ●●

●●

●●

­Clinical Signs Rabies is rarely recognized on physical exam alone. The initial clinical signs mimic other diseases and may not seem neurologic. The most consistent finding is a steadily worsening clinical course over hours or days, with an accumulation of signs that eventually reveal their neurologic origins. The pet will not respond to therapy, and will usually die within 10 days. Close monitoring, with the patient kept in isolation, is the most important premortem clinical tool. The incubation length is highly variable. Most dogs and cats develop clinical signs within six months after exposure, but some may become ill in as few as 10 days. The most common incubation period is 3–12 weeks. In general, the clinical signs of rabies may be divided into a prodromal phase, followed by a furious (excitatory or encephalitic) phase, then a dumb (paralytic) phase, then death. However, the clinical course can vary considerably. Phases may be absent or overlap. The majority of cases are associated with a prominent furious phase, followed by a short paralytic phase before respiratory arrest and death. Approximately 20% of cases lack a furious phase (i.e., dumb or paralytic rabies).

●● ●●

Restlessness. When confined, animals may pace or constantly readjust their body posture. When crouching, they may continuously shift the front paws. Unconfined animals may walk or run for miles. Hyperreactivity to sounds, movements, and touch. Bizarre behavior. The animal may bite or strike at unseen things, or stare at things in an unsettling manner. Dogs may lick their own urine or eat unusual objects (pica), and may present with gastrointestinal foreign bodies. Vocalization. Dogs or cats may repeatedly howl, with a noticeable change in voice because of alterations in laryngeal function. Cats may make aggressive screeching sounds, especially in response to stimuli. Aggression. Bites from rabid dogs usually occur without warning or behavioral cues (i.e., without snarling, growling or posturing). Cats may combine striking and biting. Confined rabid animals may repeatedly bite and gnaw at cage doors and walls. Rabies has sometimes been associated with spectacular levels of aggression, with the rabid animal repeatedly attacking even while receiving grave injuries from the bite victim. Bites are often firmly inflicted, with the rabid animal latching onto the object or individual. Grand mal seizures. Fine muscle tremors, especially in cats.

Dumb (paralytic) phase clinical signs may include the following. ●● ●●

●● ●● ●●

Persistent lethargy. Cranial nerve deficits. Pharyngeal paresis may cause a change in voice and dysphagia, leading to accumulation of saliva in the mouth and drooling. The animal may repeatedly lap at water in unsuccessful attempts to drink. Dogs often develop a dropped jaw and hanging tongue, and may gag and cough, causing them to appear to be choking on an object. Ataxia. A stiff or hunched gait. Paresis, usually beginning in the body part where the original bite was inflicted. May be unilateral at first, and the animal may present with an apparent limp. Progresses as an ascending flaccid paralysis.

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Rabid animals may exhibit a combination of excitatory and paralytic signs, in which disorientation and anxiety may be coupled with restlessness and weakness. Some animals die acutely. Most rabid animals eventually progress to generalized paralysis, respiratory arrest, and death.

effective against rabies. Secure isolation and observation, or euthanasia, of the suspect rabid animal is the primary clinical intervention.

­Diagnosis

Grave – rabies is fatal.

There is no validated test available for diagnosing rabies in a clinically ill, living animal. The direct fluorescent antibody (DFA) test is the gold standard for diagnosing rabies, and is performed on brain tissue. This test is almost always performed by a public health laboratory, and is the test that must be used when human rabies exposure is possible. For most species, the DFA test requires submission of the head or whole brain. When decapitation or debraining is required, only trained, rabies‐vaccinated staff should perform the procedure. Keep the specimen fresh and refrigerated; fixation with formalin renders it untestable by DFA. Freezing the sample will delay testing and complicate performance of the test. For bats, the entire body may often be submitted. The laboratory should be contacted in advance for requirements on specimen preparation, shipping, and submission. Serology is not useful in the diagnosis of clinical rabies in animals. Patients previously vaccinated against rabies before the illness may have positive titers. Young, unvaccinated animals may have measurable maternal antibodies against rabies, clouding interpretation. Fixed brain tissue may be examined for rabies through immunohistochemistry (IHC) by certain laboratories. Histopathology may reveal Negri bodies in rabies‐ infected neurons, but they are not consistently present. Polymerase chain reaction (PCR) testing is used as an antemortem test for rabies in people. The test is usually performed on saliva and on skin from the back of the neck of the human patient. The sensitivity of PCR testing on tissue outside the CNS is too low to be used to make decisions in managing a potential rabid animal, and is not routinely used in animals because of the risk to public safety. PCR testing may be used to supplement DFA testing on brain matter. A direct, rapid immunohistochemical test (DRIT) is used during surveys of rabies in wildlife. There is no test available for evaluating healthy animals for exposure to rabies, primarily because the virus usually remains localized and undetected by the immune system during much of the incubation period.

­Therapy If an animal is rabid, all clinical interventions will fail and the animal will die. No antivirals have been shown to be

­Prognosis

­Prevention A robust immune response is the only effective defense against rabies. Standard rabies vaccination, when administered through proper protocols, affords protection against all variants. Vaccine failure does occur, but is very rare. Numerous vaccines are licensed for use in the US. Vaccines are either inactivated (killed) or recombinant, in which a live canarypox virus expresses rabies proteins. Most rabies vaccines are first administered at age 3 months, boostered a year later, and then every one or three years thereafter. The vaccine label should be reviewed carefully for the exact protocol required. Adult dogs or cats being vaccinated for the first time should have a booster a year later. An animal is considered immunized about four weeks after its first rabies vaccination, or immediately after a booster. Indoor‐only cats should be vaccinated against rabies. Insectivorous bats, an important rabies reservoir in North America, are sometimes found inside buildings. In Los Angeles County (where the authors work), at least one rabid bat is identified indoors every year. The risk of injection‐site fibrosarcoma exists when vaccinating cats. Recombinant vaccinations appear to be associated with a lower risk than killed vaccines. All states require rabies vaccination for dogs, and several require vaccination of cats. Individual cities or counties may require vaccination of cats even when their state does not require it. Some states allow exemption from rabies vaccination in cases where it may present a severe risk to the health of the animal. The criteria for granting exemption, and the protocol for obtaining one, vary by state and locality. Note that rabies serologic testing is not accepted in lieu of vaccination in most areas of the US. Vaccination is not the only way to reduce the risk of rabies exposure. Encourage clients to keep dogs leashed when off property, and to keep cats indoors. Clients can reduce the attractions for wildlife on their property by keeping all pet food (and other consumables) and water indoors, by bat‐proofing their home, and by keeping crawl spaces and the underside of decks sealed off. Clients who rescue stray, feral or wild animals, and especially those who may participate in international rescue efforts, may face a higher risk of rabies exposure.

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They should be given detailed information about rabies, and be encouraged to budget for thorough veterinary examination as part of the processing of each animal. Rescue groups that import animals should be educated about associated legal requirements (see later).

­Public Health Implications Veterinarians routinely face rabies‐related circumstances in which they are called upon to protect not only their patient but also people and other animals. National guidelines on rabies control are routinely published by  the National Association of State Public Health Veterinarians (http://nasphv.org/documentsCompendia. html). Local animal control and public health agencies usually play the most prominent role in rabies control. Contact your local agencies in advance for information on rabies‐relating reporting, testing, and other protocols. Ask for the phone number to call in case of an emergency. Place the information in an easy‐to‐find location and review it with clinic staff. Four rabies‐related circumstances occur routinely in small animal practice. ●●

●●

Dog or cat bit a person. Legally reportable. Rabid dogs and cats can potentially shed rabies virus in their saliva for a few days before clinical signs of rabies begin. Quarantine and/or testing are typically required regardless of the rabies vaccination status of the animal. The quarantine protocol may vary depending on the severity of the bite and the requirements of the jurisdiction. If the dog or cat remains healthy for 10 days or longer, the risk of rabies for the bite victim is negligible. If the animal dies or is euthanized during this 10‐day period, it should be tested for rabies. The pet should not be vaccinated for rabies during this 10‐day quarantine. Dog or cat vs wild mammal. Legally reportable in many areas. (Note that certain wild mammal species may be exempt  –  for example, in California, bites from wild rabbits and small rodents are not reportable.) Bite wounds from the wild animal exposure may or may not be obvious. Pets found unattended near a wild mammal may have wounds hidden in the fur or mouth. Bite wounds from bats are especially hard to see, even on human skin. Pets with unexplained bite wounds may also have been exposed to a rabid animal. Ask the pet owner about the location of the biting wild mammal. If it is deceased, its body should be retrieved for testing  –  a negative rabies test on the wild mammal eliminates concerns about rabies. If the wild mammal is not available for testing, or tests positive for the virus, the pet is ­considered exposed to rabies. Regardless, the bite wound should be thoroughly

●●

●●

cleaned to remove virus particles. The next steps depend on the pet’s rabies vaccination status. –– Pet’s rabies vaccination was current before incident. Revaccinate the pet immediately. The owner should be advised that there is a risk of the pet becoming rabid. The postexposure quarantine and observation period in this case is usually 30–45 days, depending on the jurisdiction. The quarantine may potentially be performed at the pet owner’s home or may require a more secure setting, depending on the risk of rabies and local requirements. –– Pet was not vaccinated at all before incident. Tell the owner that there is an elevated risk of the pet becoming rabid, presenting risk to them and their family members. National guidelines call for the pet to be euthanized to prevent it from becoming rabid. Alternatively, the pet may be immediately vaccinated and then quarantined. In most states the quarantine period is 4–6 months. Texas requires only 90 days of quarantine, but also requires unvaccinated pets to be vaccinated three times while in quarantine: the first week, then three and eight weeks afterward. This three‐vaccine postexposure “Texas Protocol” may provide some protection against the development of rabies, and is sometimes recommended in other localities. –– Pet overdue for rabies vaccine at time of incident. These cases are often managed similarly to unvaccinated pets with a rabies vaccination being given immediately followed by a 4–6‐month quarantine period (see earlier). However, the risk of rabies transmission is lower, especially if the pet has a lifetime history of multiple rabies vaccinations, and some jurisdictions may allow for a shorter quarantine of 45 days if there is proof of the earlier vaccination. If there is no proof of earlier vaccination, national guidelines suggest a procedure called prospective serologic monitoring to determine if the animal was likely vaccinated in the past. It involves documenting a very rapid rise in rabies antibody titers following boostering, by measuring antibody levels in serum samples drawn just before the booster and again 5–7 days later. This procedure may or may not be accepted as proof of earlier vaccination, depending on the state and local law. Dog or cat suspected of being rabid. Legally reportable. Isolate the animal. Create a list of all people who may have been bitten or exposed to the saliva of the animal, and refer them to physicians for consultation. After the animal dies or is euthanized, submit the head or brain to your local rabies control agency following its protocols. Inform all the exposed individuals of the rabies test result once available. Pet imported from another country. Over the past decade, the globalization of the puppy trade and animal

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rescue efforts has led to dramatic increases in animal translocation. Rabid dogs have been imported into the US several times. As a result of the increasing number of imported dogs and an increase in noncompliance with import regulations, dog import requirements in the US were modified in 2014 to reduce the risk of importing rabies. CDC Requirements As of 2019, a health certificate is not required to import a dog or cat, but the imported pet must appear healthy on arrival. Importers of dogs must provide proof of rabies vaccination when the dog is imported from countries designated as high‐risk for rabies importation (see www.cdc.gov/importation/bringing‐ an‐animal‐into‐the‐united‐states/rabies‐vaccine. html). The vaccination must have been given 28 days or more before entry, when the dog was at least 3 months of age. Unvaccinated dogs may only be imported if they have lived in a country with a very low risk for rabies exposure for the past six months or longer, are being imported for research purposes, or on a rare case‐by‐ case basis. See www.cdc.gov/importation/bringing‐ an‐animal‐into‐the‐united‐states/index.html. USDA Requirements The US Department of Agriculture (USDA) requires dogs and cats to be at least 8 weeks old when shipped without their mothers. Dogs imported specifically for resale or adoption (i.e., importation by international dog rescue groups) must be healthy, at least 6 months old and be accompanied by a health certificate, rabies certificate, and proof of vaccination against distemper, leptospirosis, parvovirus, and parainfluenza (DHLPP). See www.aphis.usda.gov/aphis/pet‐travel/ bring‐pet‐into‐the‐united‐states. Federal, state, and local regulations regarding pet importation may vary over time, and can include additional requirements depending on the animal’s country of origin. Veterinarians are recommended to check the latest information on the websites for both CDC and USDA‐APHIS as well as any additional requirements from US Customs and Border Protection (https://help.cbp.gov/app/answers/ detail/a_id/3695/kw/3695) and from the state where the animal is arriving. Unfortunately, many imported dogs and cats are not physically inspected on arrival, so it is still possible that sick animals, or those incubating rabies, will be imported. Private practice veterinarians play a crucial role in detecting imported cases of rabies, since they may the first person to examine the pet after arrival. If a clinician suspects an imported pet has rabies or any other unusual disease, they should immediately contact their local authorities.

Bats and Public Health Bats represent a special case when it comes to rabies. Rabid insectivorous bats may transmit rabies via very shallow bites, even bites that do not bleed. In fact, two bat variants appear to have greater ability to infect through shallow bites than the canine variant. Clinic staff may hear about clients or staff who have handled bats. They should urge such people to seek public health or medical consultation. Bats found near pets, children or sleeping people, or people with lowered levels of consciousness, may have bitten a person or pet, and rabies testing of the bat should be prioritized. If the bat is not tested, it should be assumed to have been rabid for the sake of safety, and the exposed person needs to be referred for medical care. Oral Rabies Vaccination and Public Health Clients and their pets may encounter oral rabies vaccines (ORVs) that are intended for wildlife. ORVs distributed in wildlife habitats in North America and Europe have been very successful in controlling the spread of rabies in raccoons, coyotes, and foxes. Government agencies typically work to alert the local population as to when and where the baits are being distributed. These vaccines contain a live, attenuated Vaccinia virus‐vectored rabies recombinant liquid vaccine in a packet surrounded by a flavored bait. There is no live rabies virus in the vaccine. The Vaccinia virus may cause illness in immunocompromised people if the liquid vaccine comes into contact with abraded skin or mucous membranes. Human exposure to ORVs is rare. During 10 years of distribution of over 86 million ORVs in 18 states, only 296 incidents of human exposure were reported, with six illnesses. Most people were exposed when trying to remove the bait from their dog’s mouth. Clients should be advised to keep their dogs leashed when in areas that are part of an ORV baiting program. If a client may have had direct contact with the liquid vaccine inside the bait, they should wash their hands, and collect the vaccine without touching it directly, such as by wearing gloves or covering their hand with a bag. There is a toll‐free phone number printed on the side of the ORV bait that the person should call to report the incident and seek medical consultation. Rabies Preexposure Prophylaxis for Humans The CDC recommends preexposure vaccination for all people who routinely handle animals. Clinical staff, such as veterinarians and veterinary technicians, should be prevaccinated against rabies. The preexposure vaccine series consists of three intramuscular human rabies vaccinations given on days 0, 7, and either 21 or 28. Antibody titer levels should be checked every two years to insure

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that the body is still producing antibodies against the virus. Preexposure vaccination alone is not considered protective in the event of a rabies exposure. Rabies Postexposure Prophylaxis For people with a known, or suspected, exposure to  rabies, postexposure treatment is recommended. In  healthy, ­previously unvaccinated people, treatment

includes a dose of human anti‐rabies immune globulin (HRIG) injected into the bite wound (with the excess given intramuscularly) plus a series of four rabies vaccines over two weeks (on days 0, 3, 7, and 14). A fifth vaccine dose may be needed for immunosuppressed individuals on day 28. The HRIG should not be injected in the same muscle as the rabies vaccine. Previously vaccinated people only need a series of two vaccinations, on days 0 and 3, after an exposure.

­Further Reading Ehnert K, Galland GG. Border health: who’s guarding the gate? Vet Clin North Am Small Anim Pract 2009; 39(2): 359–72. Greene CE. Rabies and other Lyssavirus infections. In: Greene CE, ed. Infectious Diseases of the Dog and Cat, 4th edn. St Louis, MO: WB Saunders, 2012, pp. 179–97. National Association of State Public Health Veterinarians. Compendium of animal rabies prevention and control,

2016. www.nasphv.org/documentsCompendiaRabies. html (accessed July 2, 2019). Ugolini G, Hemachudha T. Rabies: changing prophylaxis and new insights in pathophysiology. Curr Opin Infect Dis 2018; 31: 93–101. Wilson PJ, Oertli EH, Hunt PR, Sidwa TJ. Evaluation of a postexposure rabies prophylaxis protocol for domestic animals in Texas: 2000–2009. J Am Vet Med Assoc 2010; 12: 1395–401.

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91 West Nile Virus Tracey McNamara, DVM, DACVP College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology West Nile virus (WNV) is a mosquito‐borne viral disease that causes disease primarily in birds, humans, and horses. Sporadic disease has also been reported in a number of other species including squirrels, chipmunks, bats, dogs, cats, white‐tailed deer, reindeer, sheep, alpacas, alligators, and harbor seals during periods of intense local viral activity. WNV is a zoonotic mosquito‐ transmitted arbovirus belonging to the genus Flavivirus in the family Flaviviridae. The virus can cause inapparent infection, mild febrile illness, meningitis, encephalitis or death. Dogs and cats are susceptible to WNV infection and may become infected through the bite of a mosquito, but they are significantly more resistant to disease than horses, humans, and some species of birds. It is unlikely that healthy dogs or cats will become ill. Limited information is available on WNV infections in dogs and cats but experimental studies have shown that the virus targets the meninges, dorsal horn, brainstem, cerebellar Purkinje cells, and cerebral cortex. In two naturally occurring cases in a dog and a wolf cub, myocarditis was also a finding.

­Epidemiology Like other arboviruses, WNV is maintained in nature by cycling through birds and mosquitoes. Migratory birds are thought to be primarily responsible for virus dispersal. Serologic studies of dogs and cats have shown high seroprevalence but little evidence of disease, suggesting that dogs and cats suffer little to no ill effect. In a 2002 study in Louisiana, the incidence of seropositivity was 19 times higher in outdoor dogs than in indoor dogs. Another study suggested that stray dogs and cats may

serve as better sentinels than chickens. There is a single report of WNV being isolated from a sick dog in Botswana in 1982 and in a 3‐month‐old wolf cub and older dog with immune‐mediated disease. There are three reports of naturally occurring WNV illness in cats, including one cat in New Jersey in 1999 and two cats in New York City in 2000. Experimentally infected dogs did not develop clinical signs of illness or infectious levels of viremia. Experimentally infected cats did develop brief episodes of febrile illness, lethargy or decreased appetite of short duration. Cats develop slightly higher levels of virus in their bloodstream but it is unclear if it is sufficient to infect mosquitoes. Cats fed infected mice developed viremias similar to those caused by a mosquito bite, suggesting that prey animals may serve as an important source of infection for carnivores and that outdoor cats are at greater risk of WNV infection. All dogs and cats cleared WNV from the bloodstream within days and mounted an antibody response. No evidence of prolonged or persistent infection was found. Neither dogs or cats are likely to serve as an important amplifying host epidemiologically.

­Signalment Clinical signs of WNV infection are nonspecific and diagnosis cannot be made on clinical presentation alone. Based on a small number of naturally occurring cases, dogs may present with lethargy, depression, ataxia, difficulty rising, weakness or paresis, muscle tremors or twitching, abnormal head posture, fever, and nonlocalizable pain. Other differential diagnoses for these signs would include bacterial meningitis, distemper, rabies, fungal or rickettsial infections, and primary neurologic disease. Cats develop nonspecific mild febrile illness.

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­Diagnosis

­Public Health Implications

Diagnosis of WNV in the live patient requires paired acute and convalescent serum samples for neutralizing antibody. Due to the widespread seropositivity, single serum samples are nondiagnostic. If a necropsy is performed (with appropriate biocontainment gear or facilities), the following tissues should be collected and submitted chilled, not frozen, for WNV polymerase chain reaction (PCR), immunohistochemistry and/or virus isolation: heart, kidney, brain/brainstem, a section of spinal cord.

As WNV is a biologic safety level (BSL) 3 pathogen, anyone performing a necropsy should wear protective apparel and, if possible, work in a laminar flow hood. In most cases, it would be preferable to send the body to a diagnostic laboratory with appropriate facilities for testing. There is no documented evidence of dog‐ or cat‐to‐ person transmission of WNV. WNV has not been detected in the saliva of dogs so dog bites would not appear to be a risk factor for human infection. Neither dogs or cats are considered epidemiologically important amplifying hosts, although peak viremias in cats may be high enough to infect mosquitoes at low efficiency. Suspect cases should be reported to your state veterinarian, state diagnostic laboratory or reference laboratory. Positive cases should be promptly reported to your local Department of Health. Clients may ask about methods of mosquito protection in dogs and cats. It is important to emphasize that DEET‐containing products are not approved for use in pets and should not be used. The same holds for citrus‐based oils that have been found to cause contact dermatitis. Owners should be advised to eliminate mosquito habitat and avoid outdoor exposure of pets during peak mosquito feeding activity (early morning and dusk).

­Therapy There is no specific therapy for WNV infection. Supportive care should be provided. An experimental recombinant canarypox vectored WNV vaccine protected dogs and cats against mosquito challenge but is not commercially available.

­Prognosis Full recovery is likely.

­Further Reading Austgen LE, Bowen RA, Bunning ML, Davis BS, Mitchell CJ, Chang G‐JJ. Experimental infection of dogs and cats with West Nile virus. Emerg Infect Dis 2004; 10(1): 82–88. Karaca K, Bowen R, Austgen LE, et al. Recombinant canarypox vectored West Nile virus (WNV) vaccine protects dogs and cats against mosquito challenge. Vaccine 2005; 23: 3808–13. Kile JC, Panella NA, Komar N, et al. Serologic survey of cats and dogs during an epidemic of West Nile virus

infection in humans. J Am Vet Med Assoc 2005; 226(8): 1349–53. Lichtensteiger CA, Heinz‐Taheny K, Osborne TS, Novak RJ, Lewis BA, Firth ML. West Nile virus encephalitis and myocarditis in wolf and dog. Emerg Infect Dis 2003; 9(10): 1303–6. www.cdc.gov/ncidod/dvbid/westnile/birds&mammals.htm (accessed June 24, 2019). www.oie.int/international‐standard‐setting/terrestrial‐ manual (accessed June 24, 2019).

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92 Ebola Virus Linda Kidd, DVM, PhD, DACVIM College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology

­Epidemiology

Ebola virus is an enveloped, negative‐sense RNA virus belonging to the genus Ebolavirus in the family Filoviridae. Filoviruses cause severe lethal hemorrhagic disease in people. The Ebola viral genome encodes a nucleoprotein (NP), a glycoprotein (GP), an RNA polymerase (L), and four structural proteins (VP 24, VP30, VP 35, and VP 40). A soluble form of GP thought to be important in evading humoral immunity is produced through receptor editing. The primary target cells of Ebola virus are dendritic cells (DC) and macrophages. Other cells, including hepatocytes, endothelial cells, fibroblasts, and cells of the adrenal gland, can be infected. Ebola virus can both inhibit and evade innate and adaptive immunity. For example, the virus inhibits interferon (IFN)‐alpha, IFN‐beta, and IFN‐gamma production. Infection also inhibits DC expression of CD 40, CD 80/86, and MHC II, co‐stimulatory molecules that are important in the induction of adaptive immunity. Examples of evasion tactics include shielding of GP epitopes that may be recognized by the immune system, and releasing soluble GPs that act as “decoys” that bind protective antibodies. Despite being able to avoid immune recognition, infection is associated with massive production of proinflammatory cytokines. Thus, additional cells are recruited and available for the virus to infect. The ­systemic effects of cytokines cause systemic inflammatory response syndrome (SIRS). Increased vascular permeability and endothelial cell activation associated with infection are in part mediated by GP. Disordered hemostasis may arise from endothelial cell dysfunction and damage. Tissue factor expression induced by virus or cytokines appears to play a major role in activation of coagulation and induction of disseminated intravascular coagulation (DIC).

Ebola was first discovered in 1976 near the Ebola River in what is now the Democratic Republic of Congo. Several epidemics have occurred in Africa. The largest outbreak began in 2014 and involved several West African countries. It was considered ended in 2016. Infection has been reported in healthcare workers returning from Africa in a few countries, including the United States, Spain, and the United Kingdom. Further spread in these countries has not occurred. Handling of dead infected animals or consumption of infected primates was thought to be the mechanism of entry into the human population. Human‐to‐human spread is the primary means of infection in people. Infection occurs by exposure to bodily fluids through mucous membranes or broken skin, or inoculation via needle sticks. Importantly, fomites may also play a role in transmission. The incubation period is thought to be 2–21 days. Bats are believed to be the reservoir host in Africa but this has not been confirmed. A summary of surveillance studies of naturally infected species in outbreak areas detected virus in nonhuman primates and bats and an ungulate. Recovered human patients can still shed the virus in semen and virus has been amplified from bodily fluids in patients with uveitis and meningoencephalitis after recovery. Antibodies have been detected in additional species, including dogs; 27.2% of dogs had formed antibodies to the virus in a survey of a 2001–2002 outbreak of Ebola in people in Gabon. Dogs in the study were not ill. The dogs were thought to be exposed to Ebola virus by eating carcases of dead infected wild animals (bush meat) and vomit of infected people. No virus was amplified from the blood samples. A recent in vitro study suggested that both canine and feline cells are less susceptible to infection than primate and human cells.

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At the time of this writing, whether Ebola can cause viremia and be shed by or cause clinical signs in dogs and cats is not known. As a precautionary measure, during the 2014 Ebola outbreak, a healthy dog owned by an infected Spanish nurse was euthanized by officials without testing. Global public outcry was significant given that it had not been shown that dogs can be infected with or shed the virus and that the risk was unknown. A call for testing and not automatic euthanasia was made by the World Small Animal Veterinary Association. Shortly thereafter, a dog owned by an infected nurse in Texas was quarantined and tested rather than euthanized. Reverse transcriptase polymerase chain reaction (RT PCR) testing of blood, urine, and feces was negative, and the dog showed no evidence of illness during the 21‐day quarantine.

­Clinical Signs Ebola causes severe disease in humans and nonhuman primates. Whether the virus causes clinical signs in dogs and cats is not yet known. In people, flu‐like symptoms occur, including fever, headache, and muscle pain. Gastrointestinal signs also occur. Dramatic signs of ­disordered hemostasis such as maculopapular rash, petechiae, conjunctival hemorrhage, epistaxis, and hematemesis are common. Encephalopathy also occurs.

­Diagnosis Diagnosis in people depends on the phase of infection. Direct detection of the virus is attempted if clinical signs have been present for a short period of time. Detection techniques include virus isolation, antigen‐detecting

ELISA, immunohistochemistry, and RT PCR. RT PCR is used in the emergent clinical setting. Detection of IgM antibody or demonstration of seroconversion can facilitate diagnosis of acute infection.

­Therapy Treatment of infection in people is primarily supportive. Experimental therapies that appear to hold promise were utilized in some cases in the 2014 outbreak. Vaccines are in development.

­Prognosis The mortality rate in people and nonhuman primates is high. Prompt and adequate supportive care appears to be important for survival.

­Public Health Implications In the fall of 2014, the AVMA Ebola Companion Animal Response Plan Working Group developed recommendations regarding dog and cat quarantine after exposure to a human with confirmed Ebola virus disease. These recommendations are available at www.cdc.gov/vhf/ebola/ pdf/dog‐cat‐quarantine.pdf. Public health officials and state veterinarians should be contacted immediately on an emergency basis if a pet is exposed to a person with Ebola virus infection. Regardless of whether infection occurs in dogs and cats, recommendations for people with exposure to Ebola virus include avoiding contact with their pets due to the potential risk of fur acting as a fomite.

­Further Reading Allela L, Boury O, Pouillot R, et al. Ebola virus antibody prevalence in dogs and human risk. Emerg Infect Dis 2005; 11(3): 385–90. Ansari A. Clinical features and pathobiology of Ebola virus infection. J Autoimmun 2014; 55: 1–9. Han Z, Bart SM, Ruthel G, et al. Ebola virus mediated infectivity is restricted in canine and feline cells. Vet Microbiol 2016; 182: 102–7. Martinez O, Leung L, Basler C. The role of antigen‐ presenting cells in filoviral hemorrhagic fever: gaps in current knowledge. Antiviral Res 2012; 93(3): 416–28.

Mérens A, Bigaillon C, Delaune D. Ebola virus disease: biological and diagnostic evolution from 2014 to 2017. Med Mal Infect 2018; 48(2): 83–94. Olson S, Reed P, Cameron K, et al. Dead or alive: animal sampling during Ebola hemorrhagic fever outbreaks in humans. Emerg Health Threats J 2012; 5: 1–9. Spengler JR, Stonecipher S, McManus C, et al. Management of a pet dog after exposure to a human patient with Ebola virus disease. J Am Vet Med Assoc 2015; 247(5): 531–8.

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93 Ehrlichiosis and Anaplasmosis Pedro P. Vissotto de Paiva Diniz, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology Ehrlichiosis and anaplasmosis are tick‐borne diseases caused by a group of gram‐negative intracellular bacteria belonging to the family Anaplasmataceae, order Rickettsiales (Figure 93.1). These organisms are capable of causing an array of clinical signs and even death in dogs. Some bacterial species from this family are capable of infecting cats, horses, other animals, and humans. Cats are generally subclinically infected so this chapter will focus on infection in dogs. Ehrlichia and Anaplasma preferentially parasitize leukocytes or platelets and form intracytoplasmic clusters called morulae (Figure  93.2). Understanding the differences in the target cell type, clinical signs, primary vectors, and geographic distribution can help clinicians differentiate the likely infecting organism, which has therapeutic and other clinically relevant implications. For dogs, Ehrlichia canis is the most pathogenic organism (see Figure 93.2a,b), targeting monocytes and rarely lymphocytes. Ehrlichia canis is transmitted by the brown dog tick, Rhipicephalus sanguineus, which has a ubiquitous distribution (Figure 93.3). Like E. canis, E. chaffeensis also targets monocytes but causes comparatively mild to moderate signs, and is transmitted in the United States by Amblyomma americanum (the Lone Star tick), which has a more limited geographic distribution (see Figure  93.3). Ehrlichia ewingii and Anaplasma phagocytophilum target granulocytes (see Figure 93.2c,e,f ) and often cause acute lameness or ­ polyarthritis in dogs. These organisms are transmitted by different ticks (A. americanum and Ixodes spp., respectively) with distinct but overlapping areas of distribution in the United States (see Figure  93.3). ­ Anaplasma platys is the only organism from this family to target platelets (see Figure  93.2d), causing cyclic thrombocytopenia and bleeding tendencies. A. platys is

probably transmitted by the brown dog tick so co‐­ infections with E. canis are not uncommon and result in more severe clinical disease than either agent alone. A  summary of the species of importance in dogs and cats, their associated vectors, target cell, and geographic distribution is given in Table 93.1. With advances in molecular methods of diagnosis, a growing number of species of Ehrlichia that infect humans have been documented in dogs, including E. muris and Panola Mountain Ehrlichia sp. After transmission via tick saliva, the incubation period for E. canis and A. phagocytophilum ranges from one to three weeks. The acute phase of the disease can last 1–4 weeks, when the organism rapidly multiplies within the targeted blood leukocytes and, in the case of E. canis, tissue macrophages (spleen, liver, lungs, and lymph nodes). Intracellular multiplication is achieved by impairing the antimicrobial activity of host monocytes or granulocytes by inhibiting lysosome fusion, decreasing the formation of reactive oxygen species, and inhibiting phagocytosis. The organisms also increase host cell survival by inhibiting apoptosis and leukocyte migration to tissues. In addition, Ehrlichia and Anaplasma spp. avoid the immune system by repressing the early innate immune response, and by antigenic variation of outer membrane proteins. Consequently, infected monocytes or granulocytes remain in circulation longer, facilitating subsequent transfer to ticks. Genetic analyses have demonstrated that multiple strains of E. canis and ­ A. phagocytophilum exist, which may be associated with differences in pathogenicity. In addition, host susceptibility, size of the inoculum, and concurrent infection with other tick‐borne diseases transmitted by the same tick affect the clinical presentation. An exacerbated host immune response against the infection is likely an important mechanism of the disease. Dogs infected with E. canis may develop polyclonal

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

Figure 93.1  Phylogeny of selected organisms of veterinary importance from Anaplasmataceae and Bartonellaceae families. The scale bar indicates 0.02 substitutions per nucleotide position of the 16S rRNA gene.

Ehrlichia canis Ehrlichia chaffeensis Ehrlichia ewingii Anaplasma phagocytophilum Anaplasma platys

Anaplasmataceae Family

Wolbachia pipientis Neorickettsia helminthoeca Bartonella clarridgeiae Bartonella vinsonii berkhoffii Bartonella henselae 0.02

Bartonellaceae Family

Bartonella koehlerae

(a)

(b)

10 μm

(c)

10 μm (d)

10 μm (e)

10 μm (f)

2 μm

1 μm

Figure 93.2  Ehrlichial morulae (black arrows) in canine leukocytes as seen in blood smears (a–d) and by electron microscopy (e,f ). (a) E. canis morula in a monocyte. (b) E. canis morulae in a lymphocyte. (c) Morula in a neutrophil compatible with A. phagocytophilum or E. ewingii. (d) A. platys morula in a platelet. (e) Electron microscopy of a morula in a granulocyte. (f ) Magnification of the morula seen in (e). Source: (a–d) from Allison and Little (2013). Reproduced with permission from John Wiley & Sons. (e,f ) Courtesy of Kristin Nunez, Purdue University, USA.

93  Ehrlichiosis and Anaplasmosis

or monoclonal hypergammaglobulinemia. Immune‐ mediated consumption of platelets is thought to contribute to thrombocytopenia in dogs infected with these organisms. Other mechanisms of thrombocytopenia such as splenic and vascular sequestration may also be involved. Thrombocytopathia may also occur due to antiplatelet antibodies. Deposition of immune complexes and infected cells in the vascular endothelium of kidneys, joints, and eyes may predispose to glomerulonephritis, polyarthritis, and uveitis. Systemic inflammation associated with E. canis infection may also contribute to myocarditis, meningitis, thrombosis, hyperviscosity, and vasculitis. The development of high circulating levels of antibodies against Ehrlichia or Anaplasma spp. does not promote protective immunity against reinfection. The progression of the disease appears to vary according to the cell type targeted by the parasite. Granulocytic anaplasmosis caused by A. phagocytophilum appears to be an acute illness in naturally infected dogs that can be self‐limiting. However, chronic persistent infection has been shown in experimentally infected dogs, and has been suspected in some naturally infected dogs. Similarly, granulocytic ehrlichiosis caused by E. ewingii appears to be self‐limiting, but persistent infection was detected in experimentally infected dogs. In contrast, monocytic ehrlichiosis associated with E. canis infection causes an acute illness, which may vary from mild to severe depending upon the pathogenicity of the parasite, followed by subclinical phase of months to years in duration. It is unclear if dogs are able to eliminate E. canis or if the organism remains sequestered in the spleen, bone marrow or other primary niche indefinitely. Some dogs develop a chronic phase characterized by hyperglobulinemia and bone marrow suppression, which is more difficult to treat. E. chaffeensis also appears to cause chronic infection in dogs, but dogs are generally asymptomatic.

­Epidemiology

Figure 93.3  Distribution of Amblyomma americanum, Dermacentor variabilis, Ixodes pacificus, Ixodes scapularis, and Rhipicephalus sanguineus ticks in the USA. Source: Centers for Disease Control and Prevention, www.cdc.gov/ticks/geographic_distribution.html.

The presence of Ehrlichia and Anaplasma spp. primarily follows the geographic distribution of their vectors and mammalian reservoir hosts (see Figure 93.3). E. canis has worldwide distribution, being one of the most common tick‐borne pathogens in dogs in tropical and subtropical regions. The ubiquitously distributed brown dog tick (R. sanguineus) is well adapted to urban environments. Consequently, dogs with no access to wooded areas are still at risk of E. canis infection. In contrast, the Lone Star tick (A. americanum), vector for E. ewingii and E. chaffeensis, is most frequently found in woodland habitats, especially where a high number of white‐tailed deer are present. The geographically limited distribution of dogs

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Table 93.1  Species of Ehrlichia and Anaplasma of importance to dogs and cats

Granulocytic ehrlichiosis

Granulocytic anaplasmosis

Cyclic thrombocytopenia

Ehrlichia chaffeensis

Ehrlichia ewingii

Anaplasma phagocytophilum

Anaplasma platys

Monocyte

Granulocyte

Granulocyte

Platelet

Amblyomma americanum

A. americanum

Ixodes scapularis, I. pacificus, I ricinus, I persulcatus

R. sanguineus

Dermacentor variabilis

Dermacentor variabilis

Otobius megnini

I. spinipalpis, I. trianguliceps

Main animal host

Dog

Dog

Dog

Dog, horse

Other animal hosts

Cat

Coyote, lemur, raccoon

Main reservoir

Dog

White‐tailed deer

White‐tailed deer

White‐tailed deer, white‐footed mouse, dusky‐ footed woodrat, wood mouse

Dog

Other potential reservoir(s)

Jackal, fox, coyote

Zoonotic disease

Yes

Yes

Yes

Yes

No

Geographic distribution

Worldwide

USA (southeastern and south central US)

USA (southeastern and south central US)

Worldwide

Worldwide

Disease

Monocytic ehrlichiosis

Organism

Ehrlichia canis

Main targeted cell

Monocyte

Main vector

Rhipicephalus sanguineus

Other potential vector(s)

Dog

Cat, cattle, goat, sheep

93  Ehrlichiosis and Anaplasmosis

exposed to E. chaffeensis and E. ewingii in the United States has recently been described (Figure 93.4). A. phagocytophilum is transmitted by the blacklegged tick (Ixodes scapularis or I. pacificus) in the United States and by the castor bean tick (I. ricinus) and the taiga tick (I. persulcatus) in Europe and Asia. In the United States, A. phagocytophilum is more prevalent in the Northeast and upper Midwest regions, and in

­ orthern California. Updated prevalence maps of canine n ehrlichiosis (E. canis and E. chaffeensis combined) and anaplasmosis (A. phagocytophilum and A. platys combined) in the United States are available at the Companion Animal Para­ site Council website (https://capcvet. org/maps/#2019/all/anaplasmosis/dog/united‐states/). A detailed world map of vector‐borne diseases and vectors by country is available at the Canine Vector‐Borne

Seroprevalence: No Data < 50 Samples 0% 0.1% – 1.6% 1.7% – 5% > 5%

Seroprevalence: No Data < 50 Samples 0% 0.1% – 1.6% 1.7% – 5% > 5%

Figure 93.4  Seroprevalence by state of E. chaffeensis (green) and E. ewingii (purple) in dogs. Source: Beall MJ, Alleman AR, Breitschwerdt EB, et al. Seroprevalence of Ehrlichia canis, Ehrlichia chaffeensis and Ehrlichia ewingii in dogs in North America. Parasites Vectors 2012; 5(29): 1–11. Reproduced under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0).

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Disease World Forum website (www.cvbd.org/en/ occurrence‐maps/world‐map/). Recently, two new species of Ehrlichia have been described in dogs in the United States: E. muris from a sick dog in Minnesota, and Panola Mountain Ehrlichia (PME) from a sick dog in North Carolina. E. muris is believed to be transmitted by I. scapularis in the United States, with wild rodents as reservoir, whereas PME is transmitted by A. americanum, with white‐tailed deer as the probable vertebrate reservoir in the United States.

­Signalment Male or female dogs at any age can develop the disease. German shepherd dogs are predisposed to a more severe illness during E. canis infection. Despite the fact that cats are naturally exposed to Ehrlichia and Anaplasma spp. infection, clinical manifestations are rarely reported.

­History and Clinical Signs The classic presentation in dogs is characterized by depression, lethargy, mild weight loss, and anorexia.

Frequently, pet owners will not recall tick exposure, since infection may have occurred weeks to months prior to illness. Canine monocytic ehrlichiosis is more frequently associated with history of epistaxis and signs of pallor, petechiation, and uveitis (Table 93.2, Figure 93.5). Canine granulocytic ehrlichiosis and anaplasmosis are more frequently associated with history of reluctance to stand or walk and signs of joint effusion and lameness (see Table 93.2). Anaplasma platys infection rarely causes clinical signs in the United States, but moderate to severe manifestation has been described in Europe, the Middle East, and South America. Fever, lymphadenopathy, and splenomegaly are common findings in monocytic or granulocytic ehrlichiosis. Severe cases of monocytic ehrlichiosis caused by E. canis may present with central nervous system (CNS) signs, retinal hemorrhage and detachment, and/or cardiac arrhythmias. The hallmark of the chronic phase of E. canis infection is pancytopenia from hypoplasia of all bone marrow cell lines, but protein‐losing nephropathy, diffuse muscle wasting, and secondary infections, presumably due to immunosuppression, have been described. Chronic illness associated with granulocytic ehrlichiosis/anaplasmosis has not yet been well documented in naturally infected dogs.

Table 93.2  Main features of ehrlichiosis and anaplasmosis in dogs

Disease

Monocytic ehrlichiosis

Granulocytic ehrlichiosis/anaplasmosis

Thrombocytic anaplasmosis

Disease course

Acute, subclinical, chronic

Acute

Acute, subclinical

History

Lethargy, depression, inappetence, weight loss, epistaxis

Lethargy, depression, inappetence, weight Generally asymptomatic. Weight loss, epistaxis loss, weakness, reluctance to stand or walk, lameness, stiff or stilted gait

Common signs

Fever, lymphadenopathy, splenomegaly, petechiae, pallor, uveitis

Fever, lymphadenopathy, splenomegaly, joint effusion

Less common signs

Ecchymosis, bleeding gums, melena, ocular and Pallor, scleral congestion, uveitis, head tilt, tremors, anisocoria, vomiting, nasal discharge, scleral congestion, hyphema, retinal detachment, ataxia, seizures, vestibular diarrhea signs, peripheral edema, muscle atrophy, cardiac arrhythmias, vomiting, diarrhea, erythema multiforme

Common laboratory findings

Thrombocytopenia, neutropenia, nonregenerative anemia, lymphocytosis or lymphopenia, monocytosis, eosinophilia, hyperglobulinemia (polyclonal or monoclonal), increased ALT and ALP activities, proteinuria, pancytopenia

Thrombocytopenia, lymphopenia, eosinopenia, mild nonregenerative anemia,spherocytes, polyclonal hyperglobulinemia, increased ALT and ALP activities, proteinuria, neutrophilic polyarthritis

Thrombocytopenia, mild nonregenerative anemia, mild hypoalbuminemia

Main differential diagnoses

Rocky Mountain spotted fever, idiopathic immune‐mediated hemolytic anemia, multiple myeloma, lymphocytic leukemia, systemic lupus erythematosus

Lyme disease, idiopathic immune‐ mediated polyarthritis, idiopathic immune‐mediated hemolytic anemia, systemic lupus erythematosus

Idiopathic immune‐ mediated thrombocytopenia

ALP, alkaline phosphatase; ALT, alanine aminotransferase.

Fever, lymphadenopathy, petechiae Pallor, uveitis

93  Ehrlichiosis and Anaplasmosis

­Diagnosis No single laboratory test is capable of detecting Ehrlichia or Anaplasma infection in all cases, so clinicians should be aware of the limitations of each diagnostic test (Table 93.3). Diagnosis is generally based on clinical and hematologic abnormalities, response to therapy and presence of specific antibodies against Ehrlichia and Anaplasma spp. Common hematologic abnormalities are listed in Table 93.2. In addition, it is worth noting that E. canis is an important differential for lymphocytic leukemia or multiple myeloma, as infection has occasionally been associated with marked granular lymphocytosis and monoclonal, rather than polyclonal gammopathy. E. canis infection may also be associated with positive Antinuclear Antibody (ANA) tests, so it should be ruled out in suspected cases of systemic lupus erythematosus.

Figure 93.5  Corneal edema, opacity, and pigmentation in a dog with uveitis caused by ehrlichiosis. Source: Courtesy of José Luis Laus and Ivan Ricardo Martinez Padua, UNESP, Brazil.

Table 93.3  Features of diagnostic assays available for ehrlichiosis and anaplasmosis in dogs Assay

Advantages

Morulae detection

●● ●●

Fast and cost‐effective Can be performed in practice

Disadvantages ●●

●●

IFA serology

●● ●● ●●

●●

Gold standard, widely used Antibodies can be quantified (titer) E. canis antigens cross‐react with Panola Mountain Ehrlichia Can be used to monitor therapy (decrease in titers generally associated with successful therapy)

●●

●●

●● ●●

●●

●●

ELISA serology: SNAP 4Dx Plus

●● ●● ●● ●●

●●

●●

Fast and cost‐effective Can be performed in practice Procedure is standardized E. canis spot cross‐react with E. chaffeensis A. phagocytophilum spot cross‐react with A. platys Detects five Ehrlichia and Anaplasma spp., including E. ewingii

●●

●●

●● ●●

●●

●●

●●

Recommended testing time

Sensitivity and specificity are low Requires trained operator

Acute phase (first days of illness)

Cannot be performed in practice Procedure not standardized among veterinary laboratories Indicates exposure, not infection Four‐fold increase in titer is required for definitive diagnosis Assay not available for A. platys, E. ewingii and E. muris as they cannot be cultured Lack of antibody titer in the first weeks does not rule out exposure

After 2–3 weeks post infection. Low antibody titers may be detected as early as 7 days

After 2–3 weeks post infection Does not quantify antibody response ●● A. phagocytophilum and E. canis: after 7–21 days Intended to be a screening test, post infection not a diagnostic test ● ● A. platys: after 10–21 days Indicates exposure, not infection post infection Negative results in the first weeks do not rule out exposure ●● E. ewingii: 3–4 weeks post infection May not detect Panola Mountain Ehrlichia Unknown sensitivity for E. muris Limited use for monitoring therapy, as it remains positive for months after exposure (Continued)

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Table 93.3  (Continued) Assay

Spinning‐disc interferometry serology: Accuplex4 Bio‐CD

Advantages ●●

●●

Detects antibodies against E. canis and A. phagocytophilum May detect E. canis antibodies earlier than the ELISA

Disadvantages ●●

●● ●●

●●

●●

Western immunoblotting

●●

●●

Detects specific antibody response to ehrlichial species Less cross‐reactivity among species than other serology methods

●●

●● ●● ●●

Polymerase chain reaction

●● ●● ●●

●●

●● ●●

Early detection of active infection Highly sensitive for acute phase Identifies pathogens at the species level, and even new species or strains Can test a wide variety of samples (whole blood, buffy coat, cavitary effusions, synovial liquid, tissue aspirates or fragments, and ectoparasites) Can test frozen historical samples Widely available

●●

●●

●●

●●

●●

●●

●●

Recommended testing time

Currently unknown. Cannot be performed in Recommended after 3–4 practice Indicates exposure, not infection weeks post infection Does not quantify antibody response May have less accuracy and reproducibility than the ELISA for Anaplasma spp. Unclear if detects antibodies against A. platys, E. chaffeensis, E. ewingii, Panola Mountain Ehrlichia, and E. muris Cannot be performed in practice Technically difficult Not widely available Primarily used on research

Subclinical or chronic phases

Cannot be performed in practice Procedure not standardized among veterinary laboratories Sensitivity and specificity depend on assay design Risk of false‐positive results if laboratory quality control is not implemented Limited sensitivity for chronic infection Negative results do not rule out infection, even after therapy, as bacteria levels may be lower than limit of detection of assay Not recommended as a single screening test for blood donors

Acute phase. May detect carrier state and chronically infected, but sensitivity ranges from 25% to 68%

ELISA, enzyme‐linked immunosorbent assay; IFA, immunofluorescent antibody.

Visualization of morulae in blood smears using light microscopy can facilitate diagnosis, but it is insensitive. In addition, stain artifacts or basophilic precipitates can be confused with inclusions so other diagnostic techniques are recommended. Serology based on assays such as immunofluorescent antibody (IFA), enzyme‐ linked immunosorbent assay (ELISA), spinning‐disc interferometry or western blotting is the most common method used for diagnosis. Most of the serology methods cannot precisely identify the species of Ehrlichia or Anaplasma involved due to serologic cross‐reactivity. Serology only documents exposure to, not infection with, one or more organisms, so previous exposure or self‐limiting infection should be considered in a patient with a positive result. Results of serology should be interpreted in light of clinical signs, laboratory ­abnormalities,

and other potential differential diagnosis. A single positive serology result in a healthy dog with no clinical, hematologic or biochemical abnormalities or proteinuria may not require therapy. Because serology depends upon production of specific antibodies, false‐negative results may occur in the first 1–2 weeks post infection. Seronegative dogs with compatible clinical signs should be retested 2–3 weeks later to demonstrate seroconversion, and alternative methodologies, including polymerase chain reaction (PCR), should be utilized. Continuous decrease in antibody titers after antibiotic therapy may suggest clearance of infection; however, it may take several months to years for titers to decrease. In addition, dogs with chronic infection may be seronegative due to the ability of the pathogen to evade the immune system.

93  Ehrlichiosis and Anaplasmosis

Generally speaking, in contrast to serologic assays, PCR assays are very sensitive during the acute phase of infection when organisms are circulating in high numbers. PCR panels are widely available, and many can identify the species of the pathogen(s) involved. Some PCR panels can also detect and differentiate new species or strains. Bodily fluids, effusions, cytology aspirates, tissues, and ectoparasites may all be tested using PCR. However, a negative PCR result never rules out infection. PCR assays have low to moderate sensitivity for subclinical or chronic infections because the pathogen may be sequestered in the spleen or bone marrow, or be circulating in the bloodstream at very low levels, below the limit of assay detection. Since the length of time since infection is generally unknown for a given patient, it is recommended that serology and PCR assays are used in combination (see Table 93.3).

­Therapy Doxycycline remains the treatment of choice for ehrlichiosis and anaplasmosis. In this author’s experience, a dose of 5 mg/kg PO q12h is associated with fewer gastrointestinal (GI) adverse effects (vomiting, anorexia) but has the same clinical efficacy as 10 mg/kg PO q24h. For cats and small dogs, liquid formulations are preferred, because tablets may become stuck in the esophagus and cause esophageal damage and stricture. Alternatively, at least 6 mL of liquid should be given PO following each tablet. The optimum duration of therapy is unknown. Since tetracyclines are bacteriostatic, it may require several weeks of therapy. A minimum of 28 days is recommended for acute and subclinical monocytic ehrlichiosis, and a minimum of 14 days for granulocytic ehrlichiosis and anaplasmosis for dogs and cats. Chronic cases may require much longer treatments, and consequences of long‐term infection such as bone marrow suppression may not resolve despite appropriate antibiotic therapy. Because clinical improvement generally occurs as early as 24–48 hours after the beginning of therapy, clients should be reminded not to interrupt or terminate the therapy early. Aggressive tick control should be implemented. After appropriate treatment of a dog with acute or subclinical infections, the pathogen should no longer be detected in the bloodstream by PCR or culture, but dogs with chronic infections may remain intermittently positive. Decrease of antibody titers within 6–9 months post infection may indicate clearance of the pathogen, but titers can remain elevated for more than 30 months after infection. If the patient does not tolerate oral antibiotics, IV doxycycline, oxytetracycline (7.5–10 mg/kg IV q12h) or chloramphenicol (25–50 mg/kg IV or SC q8h) can be

used until GI signs subside. Of note, chloramphenicol should be avoided in dogs with severe anemia and pancytopenia. The use of enrofloxacin is associated with initial clinical improvement, but it is not recommended as single therapy due to antibiotic resistance by E. canis. Imidocarb dipropionate should be avoided due to limited efficacy. Supportive therapy may be crucial for survival in advanced cases. IV fluids and blood transfusions are frequently used in severe anemic and thrombocytopenic cases. Granulocyte colony‐stimulating factor and recombinant human erythropoietin were used with success in one case of bone marrow suppression. Desmopressin acetate (DDAVP), a synthetic vasopressin analog, has been successfully used to treat bleeding associated with ehrlichiosis when used at 1 μg/kg SC q24h for three days. In addition, low immunosuppressive doses of prednisone (1–2 mg/kg PO q24h for 2–7 days) are recommended when thrombocytopenia fails to resolve with antibiotic therapy, or when severe or life‐threatening thrombocytopenia is present. Treatment of seropositive dogs without clinical or laboratory abnormalities is currently not recommended.

­Prognosis Acute ehrlichiosis and anaplasmosis have very good prognosis if antibiotic therapy is initiated immediately. Dogs with severe anemia, severe leukopenia, hypokalemia, prolonged activated partial thrombin time (APTT) and protein‐losing nephropathy have a higher risk of mortality. Prognosis is guarded in cases of bone marrow hypoplasia or aplasia.

­Public Health Implications Several ehrlichial species have been detected in humans worldwide. Most human cases are presented with acute evolution of fever, malaise/fatigue, myalgia and arthralgia, and fully respond to doxycycline therapy. No direct transmission from dogs to humans has ever been documented, but dogs can serve as environmental sentinels, as well as natural reservoirs of some species. Blood from infected dogs should be handled with caution and needle sticks should be avoided. Humans get infected by the bite of an infected tick so measures to prevent exposure to ectoparasites are fundamental to avoid disease in humans. E. chaffeensis is the etiologic agent of human monocytic ehrlichiosis, while A. phagocytophilum is the etiologic agent of human granulocytic ehrlichiosis, and E. ewingii causes human

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monocytic ehrlichiosis. They are considered emerging diseases in the United States, with over 7400 human cases reported in the United States in 2017. Human ehrlichiosis is most frequently reported in the southeastern and south‐­central regions of the United States. Three states (Oklahoma, Missouri, and Arkansas) account for 30% of all reported cases. Conversely, human anaplasmosis is most frequently reported in the upper midwestern and northeastern US, with six states (New York, Connecticut, New Jersey, Rhode Island, Minnesota, and Wisconsin) accounting for 90% of all reported cases. Additionally, E. chaffeensis has been documented in

Africa, Asia, the Middle East, Central and South America, while A. phagocytophilum has been documented in Europe, Asia, and South America. The fatality rate of human ehrlichiosis and anaplasmosis ranges from 0.2% to 3.7%. Ehrlichia canis infection in humans has been documented by molecular methods, but only in Venezuela (named Venezuelan human ehrlichiosis). In 2008, Panola Mountain Ehrlichia was first detected from a sick human in Georgia. In addition, infection with E. muris eauclairensis, formerly  E. muris-like agent (EMLA), has been reported in 115 human cases in the upper Midwest. No deaths have been described.

­Further Reading Allison RW, Little SE. Diagnosis of rickettsial diseases in dogs and cats. Vet Clin Pathol 2013; 42(2): 127–44. Diniz PPVP, Breitschwerdt EB. Anaplasma phagocytophilum infection (canine granulocytotropic anaplasmosis). In: Greene CE, ed. Infectious Diseases of the Dog and Cat. St Louis, MO: Elsevier Saunders, 2012, pp. 244–54. Dumler JS, Barbet AF, Bekker CP, et al. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia. Descriptions of six new species combinations and designation of Ehrlichia equi and ‘HGE Agent’ as

subjective synonyms of Ehrlichia phagocytophila. Int J Syst Evol Microbiol 2001; 51(Pt 6): 2145–65. Harrus S, Waner T, Neer TM. Ehrlichia canis infection. In: Greene CE, ed. Infectious Diseases of the Dog and Cat. St Louis, MO: Elsevier Saunders, 2012, pp. 227–38. Neer TM, Breitschwerdt EB, Greene RT, Lappin MR. Consensus statement on ehrlichial disease of small animals from the Infectious Disease Study Group of the American College of Veterinary Internal Medicine. J Vet Intern Med 2002; 16(3): 309–15. Sykes JE. Ehrlichiosis. In: Canine and Feline Infectious Diseases. St Louis, MO: Elsevier Saunders, 2013, pp. 278–89.

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94 Salmon Poisoning Disease Pedro P. Vissotto de Paiva Diniz, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology Neorickettsia helminthoeca, also known as salmon poisoning disease (SPD), belongs to the family ­ Anaplasmataceae (Figure 94.1). The organism causes an acute febrile and often fatal disease in dogs. In contrast to other pathogens of this family, N. helminthoeca does not depend on blood‐sucking ectoparasites for transmission. Dogs acquire the infection by ingesting fish infected with the SPD vector, the trematode Nanophyetus salmincola. In order to complete its life cycle, this trematode requires a snail (Oxytrema silicula), a fish (salmon, trout, lamprey, goldfish, stickback, among others), and a ­mammal or a bird (human, dog, coyote, raccoon, fox, river otter, bobcat, rat, heron, merganser, among others). Once the trematode matures and attaches to the gastrointestinal (GI) tract of the dog, N. helminthoeca is transmitted, initially multiplying in the epithelial cells and intestinal lymphoid tissue and then spreading systemically to visceral and somatic lymph nodes, where proliferation within macrophages occurs. After 8–12 days post infection, the organisms circulate in high numbers in the bloodstream, spreading to brain, liver, lungs, and spleen, causing systemic signs. Death of the dog occurs around 18 days post infection. The disease also occurs in other canids (foxes, coyotes) and bears, but it has not been described in domestic cats.

­Epidemiology Salmon poisoning disease is endemic in the Northwest Pacific of the United States (coastal regions of northern California, Oregon, and Washington) and southern

British Columbia in Canada. A similar syndrome has been described in dogs from south Brazil.

­Signalment Dogs from any breed, age, and sex can be infected with N. helminthoeca and develop SPD. In one study, intact male dogs and Labrador retrievers were overrepresented, probably associated with the popularity of this breed.

­History and Clinical Signs History of consumption of raw or improperly cooked fish, obtained by fishing or purchased at the supermarket, is very common among sick dogs. However, dogs can also be infected by swimming in rivers and lakes where contaminated cercaria are present. Inappetence and depression are usually the first findings, followed by progressive weight loss. Fever and peripheral lymphadenopathy are frequently seen during initial physical exams. Because of the multiplication of the pathogen in mesenteric and ileocecal lymph nodes, with consequent inflammation and tissue edema, vomiting, diarrhea, and dehydration are present in over 70% of the cases. Melena, hematemesis, and abdominal pain may occur in one‐third of cases. The hemorrhagic gastroenteritis may be indistinguishable from canine parvoviral enteritis in some dogs. Less frequently, polyuria, polydipsia, serous to purulent ocular discharge, neurologic signs, tachypnea, peripheral edema, hyphema, splenomegaly, scleral injection, and epistaxis may be seen. If left untreated, the illness progresses quickly,

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Section 9  Infectious Disease

Figure 94.1  Fine-needle aspirate of mesenteric lymph node of a dog with salmon poisoning. Black arrowheads indicate the presence of intracytoplasmic inclusions (morulae) within macrophages. Left figure: close-up of a macrophage with three morulae. Wright’s-Giemsa stain, x100 objective. Source: Johns et al. (2006). Reproduced with permission of John Wiley & Sons.

which can cause severe hypotension, mucosal pallor, cardiac arrhythmias, and death.

­Diagnosis Thrombocytopenia, anemia, lymphopenia, and neutrophilia with a left shift are the most consistent hematologic abnormalities. Hypocalcemia, hypoalbuminemia, hyponatremia, and hypokalemia are frequent abnormalities in the serum biochemistry. Urinalysis may reveal bilirubinuria and proteinuria. Coagulation testing may indicate evidence of disseminated intravascular coagulation. Fecal microscopy for the detection of N. salmincola is a fast, cost‐effective, and specific diagnostic procedure that can be performed in practice. Both the sedimentation and the zinc sulfate centrifugal flotation tests should be performed for better sensitivity. Light brown operculated eggs are present in fecal samples 5–8 days after ingestion of infected fish. However, eggs may not be detected in the feces of every infected dog. Fine needle aspirates from peripheral and mesenteric lymph nodes or spleen may reveal intracytoplasmic neorickettsial bodies within macrophages and histiocytic inflammation. However, the absence of morulae does not rule out the disease. Currently, serologic assays for N. helminthoeca are not  available from commercial diagnostic laboratories. However, due to acute presentation of the disease and the time needed for production of specific antibody, serology tests may not help the diagnosis. Polymerase chain reaction (PCR) assays are available for the specific detection of N. helminthoeca DNA from blood, aspirates of lymph nodes, spleen or liver, tissue samples, and feces. Because PCR assays can be specific and highly sensitive, they may aid the diagnosis of SPD if the laboratory has a

short turnover time. Due to the high mortality of SPF, clinicians should not wait for laboratory confirmation before starting appropriate antibiotic therapy.

­Therapy Tetracyclines are the antibiotic class of choice, and clinical improvement is seen within 24–72 hours. If the patient can receive oral medication, doxycycline 5 mg/kg PO q12h for 1–2 weeks is recommended. Vomiting dogs should receive doxycycline (5 mg/kg IV q12h) or oxytetracycline (7–10 mg/kg IV q8h) until GI signs subside. Trematode infection should be treated with praziquantel (10–30 mg/kg PO q24h for 1–2 days). For complicated cases, hospitalization and intensive care may be required.

­Prognosis Survival is directly related to early diagnosis and introduction of appropriate antibiotic therapy and support care, achieving >85% success rate. Most untreated ­animals die within 6–10 days of the onset of clinical signs. In contrast to other pathogens in the family Anaplasmataceae, infection with N. helminthoeca generates protective immunity against the same strain; however, alternate strains can still cause illness.

­Public Health Implications No confirmed case of human infection has ever been reported, although humans can be naturally infected with the trematode N. salmincola, which may cause abdominal discomfort and GI signs.

94  Salmon Poisoning Disease

­Further Reading Dumler JS, Barbet AF, Bekker CP, et al. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia. Descriptions of six new species combinations and designation of Ehrlichia equi and ‘HGE Agent’ as subjective synonyms of Ehrlichia phagocytophila. Int J Syst Evol Microbiol 2001; 51(Pt 6): 2145–65. Gorham JR, Foreyt WJ, Sykes JE. Neorickettsia helminthoeca infection (salmon poisoning disease). In: Greene CE, ed. Infectious Diseases of the Dog and Cat. St Louis, MO: Elsevier Saunders, 2012, pp. 220–4.

Headley SA, Scorpio DG, Vidotto O, Dumler JS. Neorickettsia helminthoeca and salmon poisoning disease: a review. Vet J 2011; 187(2): 165–73. Johns JL, Strasser JL, Zinkl JG, et al. Lymph node aspirate from a California wine-country dog. Veterinary Clinical Pathology 2006; 35(2): 243–246. Sykes JE. Salmon poisoning disease. In: Canine and Feline Infectious Diseases. St Louis, MO: Elsevier Saunders, 2013, pp. 311–19. Sykes JE, Marks SL, Mapes S, et al. Salmon poisoning disease in dogs: 29 cases. J Vet Intern Med 2010; 24(3): 504–13.

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95 Wolbachia pipientis Infection Pedro P. Vissotto de Paiva Diniz, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology Wolbachia pipientis is a gram‐negative bacteria belonging to the family Anaplasmataceae. The organism is found in all developmental stages of Dirofilaria immitis, the filarial helminth that causes heartworm disease in dogs and cats. Wolbachia is an endosymbiont of D. immitis (Figure 95.1) and supports the nematode’s survival, maturation, and reproduction. It is transmitted by the mosquito vector along with the nematode, and is released into the bloodstream in large numbers during parasite molts, microfilaremia, and upon death of the parasite. It was originally believed that Wolbachia was not pathogenic to mammals. However, in recent years, studies have shown that components of their surface membranes stimulate production of inflammatory and vasoactive cytokines from neutrophils and endothelial cells, resulting

in inflammatory lesions and pulmonary artery vasoconstriction. Experimental infection with D. immitis free of Wolbachia results in a reduced inflammatory response after anthelminthic therapy. Furthermore, antibiotic therapy that targets Wolbachia results in a significant reduction in inflammatory reactions when anthelmintic drugs are used to kill D. immitis in infected dogs. It has been shown that cats naturally infected with D. immitis and seropositive for Wolbachia surface proteins (WSP) have a greater acute inflammatory response at bronchial, vascular, and parenchymal levels when compared to healthy cats. WSP have also been detected in glomerular capillaries and may contribute to the glomerulonephropathy seen in heartworm disease. It is currently unclear if Wolbachia can cause disease in animals without the presence of D. immitis, because the bacterium has never been cultured in vitro.

Figure 95.1  Ultrastructural location of Wolbachia within their filarial host (black asterisks), characterized by bacteria within host vacuoles. Scale bar = 5 μm (left) and 1 μm (right). Source: Taylor et al. (2013). Reproduced with permission of John Wiley & Sons.

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­Epidemiology

­Therapy

Wolbachia is ubiquitous in D. immitis, being present in 100% of the helminths. Therefore, the distribution of Wolbachia follows the presence of D. immitis, which is endemic in North, Central, and South America, the coastal regions of Africa, southern Europe, India, north, central‐ east and south China, Southeast Asia, and Australia. Updated prevalence maps for heartworm are available at the Canine Vector‐Borne Disease World Forum website (www.cvbd.org/en/occurrence‐maps/world‐map/).

Because of the symbiotic relationship between Wolbachia and D. immitis, decreased bacterial load affects embryogenesis, maturation, and survival of heartworm. Treating dogs infected with D. immitis with an antibiotic that targets Wolbachia results in a decrease in microfilaremia, and when combined with ivermectin, fewer pathologic pulmonary lesions are observed compared to adulticide therapy alone (melarsomine). Therefore, the current therapeutic recommendation from the American Heartworm Society is to administer doxycycline at 10 mg/kg PO BID for four weeks in combination with ivermectin prior to adulticide therapy. Doxycycline eliminates >90% of the Wolbachia organisms from the nematode for at least three months.

­Signalment Because of the life cycle of D. immitis, clinical signs are generally detected in animals older than 1 year. Dogs are more frequently affected than cats.

­History and Clinical Signs While Wolbachia may collaborate in the pathogenesis of D. immitis infection, clinical signs are indistinguishable from heartworm disease.

­Diagnosis Wolbachia DNA can be detected using polymerase chain reaction (PCR) on EDTA–anticoagulated blood samples. Interestingly, using PCR to detect Wolbachia has been proposed as a tool to detect early D. immitis infection. WSP can also be detected using serology, but because all D. immitis stages are infected with Wolbachia, the presence of the organism is assumed in a dog with a positive test for occult heartworm antigen. Interestingly, one study suggested that dogs infected with D. immitis and seropositive for WSP were more likely to have proteinuria than WSP‐seronegative dogs.

­Prognosis Because all D. immitis are infected with Wolbachia, the prognosis of Wolbachia infection follows the prognosis for treatment of heartworm infections. Patients with no abnormal physical examination findings or radiographic changes have an excellent prognosis. Those with mild clinical signs and mild thoracic radiograph abnormalities have a good prognosis if adequately treated. Patients with moderate to severe disease, including caval syndrome, have a poor prognosis.

­Public Health Implications Dirofilaria immitis is able to infect humans, where it generally causes granulomatous lesions in lungs and subcutaneous tissue. Similar to what has been documented in dogs and cats, inflammatory reactions to these parasites in humans may be associated with their co‐infecting Wolbachia, but it is unclear if Wolbachia can cause disease without the presence of filarial nematodes.

­Further Reading American Heartworm Association. Current canine guidelines for the diagnosis, prevention, and management of heartworm (Dirofilaria immitis) infection in dogs. www.heartwormsociety.org/ veterinary‐resources/canine‐guidelines.html (accessed June 24, 2019). Frank K, Heald RD. The emerging role of Wolbachia species in heartworm disease. Compendium 2010; 32: E4. Garcia‐Guasch L, Caro‐Vadillo A, Manubens‐Grau J, et al. Is Wolbachia participating in the bronchial reactivity of cats with heartworm associated respiratory disease? Vet Parasitol 2013; 196: 130–5.

Morchon R, Carreton E, Grandi G, et al. Anti‐Wolbachia surface protein antibodies are present in the urine of dogs naturally infected with Dirofilaria immitis with circulating microfilariae but not in dogs with occult infections. Vectorborne Zoonot Dis 2012; 12: 17–20. Nelson CT. Wolbachia pipientis infection. In: Greene CE, ed. Infectious Diseases of the Dog and Cat, 4th edn. St Louis, MO: Elsevier Saunders, 2012, pp. 225–6. Taylor MJ, Boronin D, Johnston K, et al. Wolbachia filarial interactions. Cellular Microbiology 2013; 15(4): 520–526.

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96 Bartonellosis Pedro P. Vissotto de Paiva Diniz, DVM, PhD College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

­Etiology/Pathophysiology Bartonellosis is a complex syndrome caused by gram‐ negative bacteria of the genus Bartonella, which are transmitted by blood‐sucking arthropods and have tropism for mammalian erythrocytes, endothelial cells, and bone marrow progenitor cells. There are at least 14 species, subspecies or species candidatus of Bartonella capable of infecting dogs, with at least six species capable of infecting cats (Table 96.1). Most species of Bartonella reported from dogs and cats can also infect humans. Few Bartonella spp. are host specific, such as B. bacilliformis in people, with the vast majority of species having a preferred host but being capable of infecting other terrestrial mammals and even some marine animals. Bartonella spp. cause absent to minimal clinical manifestations in their preferred hosts. However, when a Bartonella sp. infects a nonadapted incidental host, clinical abnormalities are commonly seen. Severe illness is more frequently seen in immunosuppressed hosts or when co‐infection with other vector‐borne disease agents occurs. Bartonella causes chronic infections in dogs, cats, and humans, many of which are misdiagnosed and misreported. Unlike some other vector‐borne disease agents, species of Bartonella that infect dogs and cats do not form intracytoplasmic morulae. Therefore, they cannot be seen on blood smears or cytologic specimens without the use of special staining. Based on genetic analysis and presence of virulent factors, Bartonella spp. are divided into four lineages, with B. bacilliformis being the ancestor of all other species. The vast majority of Bartonella spp. capable of infecting dogs and cats (lineages 3 and 4) have virulence factors that mediate erythrocyte adhesion in a host‐specific manner (Trw type IV secretion system [T4SS]) and transfer effector proteins into host cells (VirB T4SS) to promote chronic infection. Consequently, Bartonella can invade and multiply in erythrocytes and endothelial cells and evade the host’s

immune system. Other virulence factors also participate in the pathogenesis of the infection. These include adhesion molecules, regulatory systems and a factor capable of inhibiting proinflammatory responses mediated by Toll‐like receptor IV. Bartonella spp. also infect dendritic cells, microglial cells, monocytes, macrophages, and CD34+ progenitor cells in the bone marrow. From these primary niches, Bartonella spp. are periodically shed into the bloodstream. Like bacteria belonging to the family Anaplasmataceae, Bartonella spp. inhibit apoptosis of host cells. This allows infected cells to remain in circulation longer, and increases the likelihood of acquisition by a blood‐sucking arthropod vector. The pathogen often establishes subclinical bacteremia that can last from weeks to several months. It is still unclear why some dogs or cats may develop clinical manifestations while other animals infected with the same Bartonella species are asymptomatic. Factors such as host immune response, species virulence, predisposing factors (congenital valvular disease), infection chronicity, and concomitant infections may play a role in disease manifestation.

­Epidemiology Bartonella spp. can be transmitted by an increasing number of arthropod vectors. In dogs and cats, cat fleas (Ctenocephalides felis) have been well defined as the major mode of transmission. However, epidemiologic evidence also supports the role of ticks in transmission to companion animals. Other suspected vectors include Pulex flea species, lice, and biting flies. The prevalence of Bartonella infection differs significantly between dogs and cats, being generally low in dogs and high in cats. There are also differences in infecting species, with the most common species detected in dogs being B. henselae and B. vinsonii subsp. berkhoffii, and the most common species in cats being B. henselae,

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 96.1  Species of Bartonella detected in dogs, cats, and other hosts Bartonella spp.

Preferred host

Other hosts

Vector

B. bovis

Cattle

Dog, cat, deer, elk

Biting flies?

B. clarridgeiae

Cat

Dog, human

Cat flea

B. elizabethae

Rat

Dog, human

Rodent fleas

B. grahamii

Wild mouse, vole

Dog, human

Rodent fleas

B. henselae

Cat, dog

Human, other terrestrial and marine mammals

Cat flea

B. koehlerae

Cat

Dog, human

Cat flea

B. quintana

Human

Dog, cat, monkey

Body louse

B. rochalimae

Dog and other canids, raccoon

Human

Fleas

B. taylorii

Wild mouse

Dog

Rodent fleas

B. vinsonii subsp. arupensis

Wild mouse

Dog, human

Ticks?

B. vinsonii subsp. berkhoffii

Dog and other canids

Cat, human, sea turtle

Fleas? Ticks?

B. volans‐like

Squirrel

Dog, human

Unknown

B. washoensis

Squirrel

Dog, human, rat

Unknown

Candidatus B. merieuxii

Dogs and other canids

Unknown

Unknown

followed by B. clarridgeiae and B. koehlerae. It is believed that infection with these Bartonella species in dogs and cats has a worldwide distribution, but epidemiologic data are limited. Bartonella spp. are more frequently documented in temperate regions of the world, especially in warm and humid areas. Bacteremia documented by polymerase chain reaction (PCR) ranges from 1% to 17% in sick or healthy dogs, depending upon the geographic location evaluated. In the United States, up to 10% of sick dogs with clinical manifestations compatible with tick‐borne diseases are infected with Bartonella spp. Exposure to Bartonella spp. in dogs, documented by serology methods, is generally ≤10% of tested dogs, with higher prevalence detected in sick dogs and in some tropical countries. Conversely, bacteremia prevalence in cats ranges from 5% to 40%, with prevalence of ≥ 80% in some populations (feral cats, colonies with frequent flea infestation). Seroprevalence is also frequently high, with some colonies having over 90% of cats seroreactive to Bartonella species. Because of its ubiquitous presence in cats, establishing a causal relationship between Bartonella infection and illness is challenging in this species.

­Signalment Dogs from any breed, sex, and age can be infected but in one study, over 95% of dogs bacteremic for Bartonella spp. were older than 12 months. Outdoor access, residence in a rural environment, and tick exposure are considered risk factors for infection in dogs. Similarly, cats from any breed

and sex can become infected with Bartonella spp. Young cats (≤1 year) are more likely to be bacteremic, while older cats are more likely to be seropositive. Stray or feral cats are more likely to be infected than pet cats.

­History and Clinical Signs Bartonella spp. have been associated with a wide range of diseases in dogs and cats (Box  96.1). Endocarditis is the most commonly reported manifestation in dogs (Figure 96.1). In one study, Bartonella spp. infection was confirmed as the cause of culture‐negative endocarditis in 20% of canine cases presented at a US tertiary hospital. Aortic valve involvement and congestive heart failure are more frequent in dogs with endocarditis from Bartonella spp. infection despite most of these patients having no fever. Because of the cyclic shedding of the pathogen into the bloodstream, several other organs can be affected. Consequently, the most frequent signs in dogs are nonspecific, including fever (40% of cases), lethargy (40%), weight loss (34%), anorexia (32%), and lymphadenopathy (30%). In one study, weight loss was significantly associated with Bartonella when compared to other dogs suspected of vector‐ borne diseases. Although Bartonella spp. DNA has been detected in dogs and cats with diverse signs and syndromes (see Box 96.1), causal relationships have mainly been confirmed in dogs and cats with endocarditis, myocarditis, fever of unknown origin, and specifically in cats, fetal reabsorption and stillbirth. (see Box 96.1).

96 Bartonellosis

Box 96.1  Clinical signs and syndromes potentially associated with Bartonella infection in dogs and cats Reported from dogs and cats Endocarditis Fever of unknown origin Hepatitis Lameness Lymphadenopathy Myocarditis Skin lesions Uveitis Only reported from dogs Bacillary angiomatosis Cavitary effusion Chronic erosive polyarthritis Epistaxis Granulomatous hepatitis Hyperviscosity syndrome Meningoradiculoneuritis Peliosis hepatis Polyarthritis Systemic pyogranulomatous disease Only reported from cats Cholangitis Diaphragmatic myositis Gingivitis Hepatitis Interstitial nephritis Lower urinary tract disease Neurologic signs (nystagmus, focal motor seizures, behavior changes) Osteomyelitis Reproductive failure (fetal reabsorption, stillbirth) Stomatitis

(a)

Chronic infection in dogs is associated with pyogranulomatous disease, which can be located in several sites, including lymph nodes, liver, and the central nervous system (Figure  96.2). Consequently, clinical signs vary depending upon the extent of damage caused in one or more organs. Bartonella infection in humans is associated with vasculoproliferative disease (bacillary angiomatosis and peliosis hepatis), which has been also described in dogs (Figure 96.3). Despite the growing number of reports of sick dogs infected with Bartonella spp., two recent studies failed to reproduce clinical signs in dogs that were experimentally infected with B. henselae, B. vinsonii subsp. berkhoffii, and B. rochalimae. In addition, another study documented that 18% of asymptomatic dogs carry Bartonella DNA in the blood or lymph nodes. In one study, 8% of Bartonella‐infected dogs were seropositive for at least one other vector‐borne pathogen. Therefore, the pathogenesis of bartonellosis in dogs is still unclear, and clinicians should rule out other relevant differentials before considering this diagnosis. In naturally infected cats, clinical signs are rarely reported. Similarly, experimental infection often causes mild and transient clinical signs such as fever, lethargy, and lymphadenopathy in less than a third of infected cats. However, more severe manifestations have been reported from experimental studies, including mild neurologic signs (nystagmus, intermittent tremors, and focal motor seizures), epaxial muscle pain, reproductive failure, and severe myocarditis (respiratory distress, cardiac murmurs, and arrhythmias). As seen in dogs, Bartonella infection in cats can be associated with granulomatous lesions in several organs. Osteomyelitis (Figure  96.4), uveitis, hepatitis, gingivitis, and stomatitis, among other syndromes (Table 96.2), have been associated with natural Bartonella spp. infection in cats. (b)

Figure 96.1  Endocarditis caused by Bartonella vinsonii subsp. berkhoffii in a dog. (a) Heart of a dog with mitral valve perforation (black arrow). (b) Close‐up of the perforation in the supravalvular region of the mitral valve. Source: Chomel et al. (2009). Reproduced with permission of John Wiley & Sons.

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(a)

(b)

Figure 96.2  Computed tomography (CT) of spinal cord compression (a, black arrow) and vertebral bone reabsorption (b, black arrows) caused by an inflammatory granuloma (m, mass lesion) in a dog infected with B. vinsonii subsp. berkhoffii. Source: Cross et al. (2008). Reproduced with permission of John Wiley & Sons.

(a)

(b)

(c)

200 μm

30 μm

Figure 96.3  Bacillary angiomatosis in a dog. (a) Erythematous nodular skin lesion. (b) Proliferating capillaries (arrowheads) seen in skin nodules. (c) Bacteria present within the lesion, positively stained by Warthin–Starry. Source: Yager et al. (2010). Reproduced with permission of John Wiley & Sons.

­Diagnosis The most common laboratory abnormalities in dogs with endocarditis caused by Bartonella spp. are leukocytosis (78%), hypoalbuminemia (67%), thrombocytopenia (56%), and elevated liver enzymes (56%). Commonly, dogs infected with Bartonella spp. have laboratory abnormalities indistinguishable from other vector‐borne diseases. Interestingly, although hyperglobulinemia occurs in dogs with bartonellosis, in one study, the presence of hypoglobulinemia was more frequent in Bartonella‐ infected dogs than dogs suspected of other vector‐borne disease. In cats, laboratory abnormalities, when present, are limited to transient mild anemia, inflammatory leukogram, and/or eosinophilia in most cases. ­ Hyperglobulinemia, particularly polyclonal gammopathy, is associated with Bartonella spp. in cats. Because of the diversity of clinical presentations, the lack of an in‐clinic rapid diagnostic test, and the limited diagnostic value of serology, diagnosis of canine or feline

bartonellosis is a great challenge. For life‐threatening illnesses such as endocarditis, myocarditis, and neurologic disease, clinicians should promptly investigate the presence of Bartonella spp. and presumptively treat for this infection, because specific antibiotic therapy is required for adequate management (see therapy section). The diagnostic approach of nonlife‐threatening cases is based on ruling out other illnesses, including other vector‐borne pathogens present in the area, submitting samples for microbiologic tests, banking samples, and evaluating response to antibiotic treatment, since laboratory test results may take weeks to months to become available. This author strongly recommend clinicians to bank samples at –20 °C (EDTA‐blood, serum, plasma, tissues, etc.) collected prior to antibiotic therapy, which can be later tested by DNA and culture methods, because antibiotic therapy rapidly decreases the chances of detection of Bartonella spp. by these laboratory methods. The microbiologic diagnosis of Bartonella infection is based on detection of Bartonella from whole blood,

96 Bartonellosis

(a)

(b)

Figure 96.4  Radiograph images of the right forelimb of a cat with osteomyelitis due to Bartonella vinsonii subsp. berkhoffii. Dorsopalmar (a) and mediolateral (b) views show cortical bone destruction, irregular osteoproliferation and moderate thickening of the soft tissue (white arrows). Source: Varanat et al. (2009). Reproduced with permission of John Wiley & Sons. Table 96.2  Average bacteremia level found in immunocompetent cats, dogs, and humans Cats

Humans

Dogs

Host type

Reservoir

Accidental

Accidental

Genome copies/μL

105–106

1–10

1–10

Bacteria per red blood cell (RBC)

1/10–100 RBC

1/105–1/106 RBC

1/105–1/106 RBC

Source: Breitschwerdt et al. (2010). Reproduced with permission of John Wiley & Sons.

other body fluids (cavitary effusions, synovial fluid, cerebrospinal fluid, etc.) or tissue specimens by specific polymerase chain reaction (PCR) assays or isolation by blood culture. While isolation remains the microbiologic gold standard, standard blood culture has limited diagnostic use because Bartonella spp. are  fastidious bacteria that take months to grow in standard culture media. In addition, Bartonella bacteremia levels in dogs are very low (see Table 96.2) when compared to cats. Therefore, sensitivity of standard blood culture is very low in dogs. Consequently, preenrichment techniques are required in dog samples to increase the number of organisms to detection levels of PCR assays. Currently, the most sensitive diagnostic technique combines preenrichment culture in an insect‐based liquid media (BAPGM) coupled with

isolation in blood‐based solid media and specific PCR testing of samples pre‐ and postenrichment culture, with DNA sequencing to confirm the species of Bartonella involved (so‐called “enrichment PCR” platform). This approach is capable of detecting more than twice the number of infected dogs than the direct PCR from clinical specimens. In addition, using this platform allows initial results to be obtained as early as two weeks, with a number of new Bartonella species being detected in dogs. This assay is available at one commercial laboratory (www.galaxydx.com). The direct detection of Bartonella DNA by PCR from blood, other body fluids or tissues is also available through commercial laboratories. Depending upon the assay design, this test may have high specificity but sensitivity is always limited by the magnitude of bacteremia (see Table  96.2). Therefore, direct PCR is a good diagnostic tool for feline samples, but it has limited sensitivity for canine samples. In a recent study, direct PCR from clinical specimens only detected 44.7% of dogs infected with Bartonella spp. Therefore, negative PCR results should never be used to rule out Bartonella spp. infection in dogs. It is important that clinicians choose a diagnostic laboratory that offers PCR testing for the entire Bartonella genus, not only for selected species, because a growing number of new Bartonella species has been described in sick and healthy companion animals recently. Serology assays are available through veterinary laboratories, but have limited diagnostic use in dogs and cats because presence of antibodies does not correlate with infection. Only one‐third to one‐half of infected dogs and cats have detectable antibodies against Bartonella spp. Therefore, negative serologic testing does not rule out infection with Bartonella spp. Also, seropositivity does not indicate infection because only one‐third of seropositive animals are bacteremic. When seropositive dogs are detected, serology may be helpful to monitor therapy progression, because many dogs that experience resolution of disease manifestation have decrease in antibody titers within 3–6 months following treatment. Serology has better diagnostic use in Bartonella endocarditis, which is usually associated with high antibody titers. There is some level of cross‐reactivity among Bartonella spp. but multiple species‐specific assays should be performed, including specific assays for B. henselae, B. vinsonii subsp. berkhoffii, B. koehlerae, and B. clarridgeiae (which cross‐reacts with B. rochalimae).

­Therapy The optimal treatment protocol for Bartonella spp. infection in dogs and cats is unknown. Similar to other vector‐borne infections, elimination of Bartonella may

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not be achievable in all cases despite long‐term use of antibiotics. Clinicians should focus on resolution of clinical abnormalities and monitor therapeutic success with serologic testing, PCR, and culture. Antibiotic therapy is recommended for two weeks past clinical resolution, which may require more than three months in some cases. In dogs, while single therapy with doxycycline or azithromycin has been previously used with moderate success, rapid development of resistance has been documented. The current recommendation for clinically stable dogs is doxycycline at 5 mg/kg PO q12h for seven days, adding a second antibiotic such as a fluoroquinolone (enrofloxacin at 5 mg/kg PO q12h for 4–6 weeks) or rifampin (5 mg/kg PO p24h for 4–6 weeks). This stepwise approach is recommended in order to decrease the risk of Jarisch–Herxheimer reaction, which is caused by rapid death of bacteria and release of endotoxins with consequent increase of inflammatory cytokines in the bloodstream. Dogs that experience this reaction show lethargy, vomiting, and signs resembling bacterial sepsis that can last for a few days, and can be misinterpreted as an adverse drug event. Therefore, clinicians should not interrupt antibiotic therapy if a Jarisch–Herxheimer reaction occurs. Empirically, corticosteroids at antiinflammatory doses generally alleviate these signs. Life‐threatening diseases associated with Bartonella spp. infection in dogs (such as endocarditis, myocarditis, meningoencephalitis, etc.) should be treated with a combination of a fluoroquinolone or doxycycline with an aminoglycoside (amikacin at 15–20 mg/kg IV/IM/SC q24h), with constant monitoring of renal function. When discharged, these patients should receive doxycycline and enrofloxacin as described above. In cats, administration of antibiotics generally limits bacteremia but does not eradicate the infection in  all cats. The current recommendation from the American Association of Feline Practitioners (AAFP) is based on doxycycline at 5 mg/kg PO q12h or 10 mg/ kg PO q24h for a minimum of four weeks. If a poor clinical response is observed in the first seven days of therapy, enrofloxacin at 5 mg/kg PO q24h or pradofloxacin at 3–5 mg/kg PO q24h should be used. It is currently unknown if these fluoroquinolones should be used in addition to doxycycline or as monotherapy in cats, as is recommended in dogs. Pradofloxacin has been shown in vitro to have superior activity against feline and human Bartonella isolates than enrofloxacin. Therefore, it is currently recommended for bartonellosis in cats. However, no information is currently available about the efficacy of pradofloxacin for canine bartonellosis.

­Prognosis Prognosis of bartonellosis in dogs and cats varies with the organ or system affected. For nonlife‐threatening bartonellosis, anecdotal reports indicate moderate to good prognosis when adequate antibiotic therapy is instituted. Bartonella endocarditis has guarded prognosis, with dogs having shorter survival time than those with other causes of endocarditis.

­Public Health Implications Human bartonellosis ranges from benign self‐limiting manifestation of cat scratch disease (CSD) to devastating hemolytic anemia seen in Oroya fever. The most common manifestation of human bartonellosis is cat scratch disease, followed by bacillary angiomatosis in immunocompromised patients and blood culture‐negative endocarditis. Recent reports have associated Bartonella spp. infection with chronic neurologic disease, rheumatic signs, and small vessel syndrome. Direct transmission from infected dogs has not been documented, but they can be a reservoir for vector‐mediated transmission to humans. Conversely, since Bartonella spp. can survive in flea feces for several days, cats can transmit Bartonella spp. to humans through scratching when cat claws are contaminated with infected flea feces. The Centers for Disease Control and Prevention and the AAFP currently recommend the following steps to avoid human infection. ●●

●●

●●

●●

●●

●●

●●

●●

●●

Flea control should be initiated and maintained year round. If a family member is immunocompromised and a new cat is to be acquired, adopt a healthy cat >1 year of age and free from fleas. Immunocompromised individuals should avoid contact with cats of unknown health status. Be cautious about adding stray cats or cats from shelters to the household without controlling fleas. Cat claws should be trimmed regularly, but declawing of cats is generally not required. Scratches and bites should be avoided (including rough play with cats). Cat‐associated wounds should be washed promptly and thoroughly with soap and water and medical advice sought. While Bartonella spp. have not been shown to be transmitted by saliva, cats should not be allowed to lick open human wounds. Keep cats indoors to minimize hunting and exposure to fleas and other possible vectors.

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­Further Reading Breitschwerdt EB, Maggi RG, Chomel BB, Lappin MR. Bartonellosis: an emerging infectious disease of zoonotic importance to animals and human beings. J Vet Emerg Crit Care 2010; 20: 8–30. Brunt J, Guptill L, Kordick DL, Kudrak S, Lappin MR. American Association of Feline Practitioners 2006 Panel report on diagnosis, treatment, and prevention of Bartonella spp. infections. J Feline Med Surg 2006; 8: 213–26. Chomel BB, Kasten RW, Williams C, et al. Bartonella endocarditis: a pathology shared by animal reservoirs and patients. Ann N Y Acad Sci 2009: 1166: 120–6. Cross JR, Rossmeisl JH, Maggi RG, et al. Bartonella‐associated meningoradiculoneuritis and dermatitis or panniculitis in 3 dogs. J Vet Intern Med 2008; 22: 674–8.

Diniz PPVP. Canine bartonellosis. In: Bonagura JD, Twedt DC, eds. Kirk’s Current Veterinary Therapy, 15th edn. St Louis,MO, Elsevier Saunders, 2014, pp. 1261–7. Guptill L. Bartonella infections in cats: what is the significance? In Practice 2012; 34: 434–45. Harms A, Dehio C. Intruders below the radar: molecular pathogenesis of Bartonella spp. Clin Microbiol Rev 2012; 25: 42–78. Sykes JE, Chomel BB. Bartonellosis. In: Sykes JE, ed. Canine and Feline Infectious Diseases. St Louis, MO: Elsevier Saunders, 2013, pp. 498–511. Varanat M, Travis A, Lee W, et al. Recurrent osteomyelitis in a cat due to infection with Bartonella vinsonii subsp. berkhoffii genotype II. J Vet Intern Med 2009; 23(6): 1273–7. Yager JA, Best S, Maggi RG, et al. Bacillary angiomatosis in an immunosuppressed dog. Vet Dermatol 2010; 21: 420–8.

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97 Hemotropic Mycoplasma Séverine Tasker, BSc, BVSc (Hons), PhD, DSAM, DACVIM-CA, PGCertHE, MRCVS Bristol Veterinary School, University of Bristol, Bristol, UK

­Etiology/Pathophysiology The hemotropic mycoplasmas (hemoplasmas) are unculturable bacteria that parasitize red blood cells and can induce hemolysis, causing anemia. Commonly recognized feline and canine hemoplasma species are shown in Table 97.1. Mycoplasma haemofelis is the most pathogenic of the feline and canine species described, and can cause significant hemolytic disease in immunocompetent cats. In cats, Mycoplasma haemofelis appears to be most pathogenic during acute infection, whereas chronic infection is not usually associated with significant anemia. Candidatus Mycoplasma haemominutum and Candidatus Mycoplasma turicensis infections are not usually associated with clinical signs, unless concurrent disease or immunusuppression exist in the host cat, such as feline leukemia virus (FeLV) infection, neoplasia. However, anemia due to these agents has occasionally been reported in immunocompetent cats. Ca. M. haemominutum has also been associated with the development of myeloproliferative diseases in cats with concurrent FeLV infection. In dogs, M. haemocanis and Ca. M. haematoparvum infections are not often associated with hemolytic anemia unless infection occurs in a splenectomized host (particularly for M. haemocanis) or in association with concurrent disease or immunosuppression. Asymptomatic carrier status can exist with all of the hemoplasma species, and so the detection of hemoplasma infection is not always indicative of clinical disease associated with that species.

­Epidemiology The prevalence of the commonly recognized feline and canine hemoplasma species is shown in Table 97.1; these vary considerably due to the differing populations of cats

and dogs sampled in studies used to generate these data. Retroviral‐infected cats are also usually found to be at increased risk in studies, although results are variable for FeLV and feline immunodeficiency virus (FIV). A recent study in the United States reported a low prevalence of canine hemoplasma infection in dogs. The prevalence of feline hemoplasma infections is increased in cats that are male, non-pedigree and with access outdoors. The natural route of transmission of hemoplasma infection in the field has not yet been determined but the clustered geographic distribution of infection in some studies supports the role of an arthropod vector in hemoplasma transmission. The cat flea, Ctenocephalides felis, has been implicated in feline hemoplasma transmission, but this has not been definitively proven in experimental studies, and transmission of M. haemocanis by the brown dog tick, Rhipicephalus sanguineus, has been suggested. Studies have successfully transmitted feline hemoplasma infection via subcutaneous inoculation of blood containing low numbers of organisms, suggesting that transmission by aggressive interaction (e.g., fights) in the field is possible. Blood transfusion is another potential route of transmission, and blood donors should be screened for hemoplasma infection. Vertical transmission may also occur.

­Signalment Older cats are more likely to be infected with Ca. M. haemominutum whilst younger cats are more likely to show disease due to M. haemofelis infection. Canine hemoplasma infections appear to be more common in  kennel‐housed dogs, younger dogs, and dogs with mange.

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 97.1  Hemoplasma species, their prevalence and pathogenicity Hemoplasma

Prevalence

Pathogenicity summary

Mycoplasma haemofelis

0.4–46.6%

Can result in hemolytic anemia in immunocompetent cats

Candidatus Mycoplasma haemominutum

8.1–46.7%

Candidatus Mycoplasma turicensis

0.4–26%

Can result in a drop in erythrocyte parameters but not usually severe enough to cause anemia unless cat has concurrent disease or is immunocompromised, e.g., retrovirus infection

Mycoplasma haemocanis

0–45%

Candidatus Mycoplasma haematoparvum

0–33%

Hemolytic anemia primarily seen in splenectomized dogs, and occasionally in immunocompromised dogs (e.g., with neoplasia)

­History and Clinical Signs When anemia results from hemoplasma infection, historical features can include lethargy, inappetence, pallor, weakness, pica, and weight loss. In immunocompromised hosts, evidence of concurrent disease is present. Dogs may have a history of splenectomy. Clinical signs can include pyrexia, weakness, pallor, tachypnea, tachycardia (sometimes with a hemic murmur), bounding pulses, dehydration, cardiac murmurs, sometimes splenomegaly (in nonsplenectomized hosts) and, occasionally, jaundice.

­Diagnosis Acute hemoplasma infections are typically associated with a regenerative anemia with reticulocytosis, macrocytosis, polychromasia, and anisocytosis. However, the anemia may be nonregenerative if the animal is sampled early in the disease process (i.e., during the preregenerative phase) or if concurrent disease is present that inhibits regeneration (e.g., concurrent FeLV infection, neoplasia). Acute M. haemofelis and M. haemocanis infections can be associated with the presence of persistent saline autoagglutination or positive Coombs’ testing, indicating the presence of erythrocyte‐bound antibodies. Auto­ agglutination may be noted on blood smear examination but the contribution of such antibodies to the develop-

ment of hemoplasma‐associated anemia has not been confirmed. Serum biochemistry may reveal elevated liver ­parameters (ALT and AST) due to anemia‐associated hypoxia, mild to moderate hyperbilirubinemia due to hemolysis, and prerenal azotemia due to dehydration. Hyperproteinemia may occur. Hemoplasmas are ­currently unculturable in vitro. Cytologic detection of hemoplasma organisms on blood smears is occasionally possible but this is known to be very insensitive and shows poor specificity, and cytology cannot differentiate between hemoplasma species. When organisms are visible, they appear as small basophilic organisms on the surface of erythrocytes with Romanowsky‐based stains (e.g., Wrights’, Diff‐Quik™). M. haemocanis has a tendency to form chains on the surface of erythrocytes, and so may be easier to differentiate from stain artifacts and other erythrocyte inclusions with which hemoplasmas are confused. Polymerase chain reaction (PCR) assays are usually sensitive and specific, if designed appropriately, for the diagnosis of hemoplasma infection, and species‐specific assays exist for both dogs and cats. EDTA‐anticoagulated blood is  usually the appropriate sample type for PCR. Real‐time PCR (qPCR) assays are now increasingly used, and these allow for quantification of hemoplasma organism numbers (Figure 97.1), as well as detection of infection. Organism numbers in feline hemoplasma infections are often high during acute infection but it should be noted that large fluctuations in blood organism numbers can occur over time in some M. haemofelis‐infected cats, meaning that qPCR results may not be correlated with anemia. In addition, PCR results may be negative in an infected cat if a sample is obtained when the number of circulating organisms is very low, below the limit of detection of the PCR  assay. Furthermore, the existence of asymptomatic carrier cats for all hemoplasma species means that the PCR results should always be interpreted in conjunction with the patient’s clinical signs, the degree and nature of the anemia present, and any concurrent signs or diseases that could be contributing to the clinical presentation.

­Therapy Therapy for hemoplasmosis is required if infection is diagnosed in an animal with clinical signs and clinicopathologic changes consistent with hemoplasma infection. However, no treatment regime has yet been described that consistently eliminates hemoplasma infection  with all species and the aim of most treatment protocols is resolution of clinical signs associated with infection.

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­Further Reading Baumann J, Novacco M, Riond B, Boretti FS, Hofmann‐ Lehmann R. Establishment and characterization of a low‐dose Mycoplasma haemofelis infection model. Vet Microbiol 2013; 167(3–4): 410–16. Compton SM, Maggi RG, Breitschwerdt EB. Candidatus Mycoplasma haematoparvum and Mycoplasma haemocanis infections in dogs from the United States. Compar Immunol Microbiol Infect Dis 2012; 35(6): 557–62. Maggi RG, Compton SM, Trull CL, et al. Infection with hemotropic Mycoplasma species in patients with or without extensive arthropod or animal contact. J Clin Microbiol 2013; 51(10): 3237–41.

Novacco M, Meli ML, Gentilini F, et al. Prevalence and geographical distribution of canine hemotropic mycoplasma infections in Mediterranean countries and analysis of risk factors for infection. Vet Microbiol 2010; 142: 276–84. Novacco M, Sugiarto S, Willi B, et al. Consecutive antibiotic treatment with doxycycline and marbofloxacin clears bacteremia in Mycoplasma haemofelis-infected cats. Vet Microbiol 2018: 217:112–20. Steer JA, Tasker S, Barker EN, et al. A novel hemotropic Mycoplasma (hemoplasma) in a patient with hemolytic anemia and pyrexia. Clin Infect Dis 2011; 53(11): e147–51.

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98 Nonhemotropic Mycoplasma, Ureaplasma, and L‐Form Bacteria Joachim Spergser, Dipl.Tzt., Dr. Med. Vet., DECVM Institute of Microbiology, University of Veterinary Medicine, Vienna, Austria

­Etiology Mycoplasmas (general name for members of the class Mollicutes) are unusual bacteria representing the small­ est and simplest self‐replicating organisms. They are distinguished from ordinary bacteria by their complete lack of a cell, resulting in cellular pleomorphism and resistance to cell wall‐inhibiting antimicrobials. The lack of a protective cell wall also makes mycoplasmas fragile outside their hosts. Due to their extremely small genome size, mycoplasmas possess limited anabolic and metabolic capabilities and maintain intimate para­ sitic lifestyles, depending on nutrients from their host cell environment. To keep this parasitic mode of life, mycoplasmas have developed sophisticated mecha­ nisms to colonize their hosts and resist host defense. A highly dynamic, versatile membrane surface architec­ ture appears to be crucial for their survival, and for establishing and maintaining a subtle relationship with their host. Certain mycoplasmas are capable of entry into nonphagocytic cells, providing them with the abil­ ity to resist host defenses or antibiotic treatment, which may contribute to chronic infection. Nonhemotropic mycoplasmas usually exhibit a rather strict host and tissue specificity with predilection for mucous membranes of the respiratory and urogenital tract. Pathogenic mycoplasmas are not considered highly virulent and mostly cause mild, slowly progressive, chronic infections. Host cell damage and the resulting clinical manifestations appear to be mainly due to host immune reactions and inflammatory responses rather than to direct toxic effects of mycoplasma components. Nonhemotropic mycoplasmas that have been isolated from cats and dogs include species within the genera Mycoplasma, Ureaplasma, and Acholeplasma. Some canine and feline mycoplasmas are considered mere

commensals, but certain members are proven pathogens or play an etiologic role as opportunists in miscellaneous conditions (Table. 98.1). Little is known about the virulence factors of canine and feline mycoplasmas so far. However, the availability of the complete genome of two canine Mycoplasma spe­ cies (M. canis, M. cynos) will certainly increase under­ standing of their pathogenic properties in the future. Recently, a hemagglutinin that is probably involved in cytadherence to host cells has been identified and char­ acterized in M. cynos. Furthermore, a secreted sialidase that presumably promotes colonization and tissue inva­ sion has been proposed as a candidate virulence factor of M. canis and M. cynos. Its expression in canine myco­ plasmas varies significantly among strains which may contribute to the variable spectrum of clinical manifesta­ tions and disease outcomes. Intracellular localization of M. canis has also been demonstrated which may contrib­ ute to chronicity of infection or perturbation of cell func­ tion and integrity. L‐forms are cell wall‐deficient morphotypes of normal bacteria that resemble mycoplasmas. L‐forms can be generated from many bacterial species by treatment with lysozyme or by exposure to beta‐lactam antibiotics or host immune responses. Knowledge of the clinical sig­ nificance of L‐form bacteria is fragmentary and their role as a cause of disease is still under debate.

­Epidemiology Mycoplasmas are common inhabitants of the orophar­ ynx and upper respiratory tract of dogs and cats wherein they are thought to be part of the normal bacterial flora. Mycoplasmas have been isolated from lungs of dogs and cats with pneumonia, are shown to be absent in the lungs

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Table 98.1  Nonhemotropic mycoplasmas frequently isolated from domestic dogs and cats Primary colonization site

Diseases associated with or caused by mycoplasmas isolated from domestic dogs and cats

ONP, RT, UGT

CIRD, tracheobronchitis, bronchopneumonia, pyothorax

ONP, RT, UGT

Urethritis, epididymitis, orchitis in male dogs; endometritis, metritis, adverse pregnancy outcomes in female dogs

Mycoplasma edwardii

ONP, RT, UGT

Polyarthritisa, meningoencephalitisa

Mycoplasma molare

ONP, UGT

ND

Mycoplasma maculosum

ONP, RT, UGT

ND

Mycoplasma opalescens

ONP, UGT

ND

Mycoplasma mucosicanis

ONP, RT, UGT

ND

Mycoplasma spumans

ONP, RT, UGT

Polyarthritisa

Ureaplasma canigenitalium

UGT

Infertility in male dogsa

C, ONP, RT

Conjunctivitis, keratitisa, URTD, bronchopneumonia, pyothorax, polyarthritis, meningoencephalomyelitisa

Mycoplasma gateae

ONP, RT

Polyarthritis, bronchopneumoniaa

Mycoplasma feliminutum

ONP, RT

ND

Ureaplasma felinum

ONP

ND

ONP

ND

ONP, RT, UGT

ND

UGT

ND

Species

Primary host

Mycoplasma cynos

Domestic dogs (canine mycoplasmas)

Mycoplasma canis

Mycoplasma felis

Domestic cats (feline mycoplasmas)

Ureaplasma cati Mycoplasma arginini Acholeplasma laidlawii

Various (ubiquitous mycoplasmas)

C, conjunctiva; ND, not defined, likely to be a commensal; ONP, oro‐ and nasopharynx; RT, respiratory tract; UGT urogenital tract. a  Single reports.

of healthy cats but can be isolated from the lungs of healthy dogs. Younger, group‐living animals with syn­ demic viral/bacterial infections or underlying diseases that impair defense mechanisms are more likely to be infected by mycoplasmas. They are efficiently transmit­ ted through oronasal contact with aerosolized respira­ tory secretions or contact with freshly contaminated fomites. Mycoplasmas are among the recognized normal flora of the vagina and prepuce of dogs but are rarely isolated from the feline urogenital tract. They have been associ­ ated with canine urogenital diseases in subpopulations of colonized individuals. Reasons and risk factors for dis­ ease development and progression are unknown. Canine genital mycoplasmas may be transmitted via sexual intercourse, artificial insemination or orogenital contact. Mycoplasmas may also be transmitted vertically to fetus or offspring by ascending intrauterine infection, hema­ togenously acquired placental infection, or during pas­ sage through the birth canal.

­Signalment Younger, immunosuppressed dogs and cats are more likely to be affected by mycoplasmas and develop more severe diseases.

­History and Clinical Signs Ocular Mycoplasma Infections in Cats Mycoplasma felis is considered a major pathogen in con­ junctivitis of cats. The prevalence rate of the agent in ocular swabs taken from cats displaying conjunctivitis was 9.6% or 25% which is significantly higher than the respective rates of 2.3% or 0% present in clinically healthy cats. In contrast, M. arginini and M. gateae more often exist on the conjunctival surface of healthy cats. Experimental infections of M. felis have produced con­ junctivitis only in young cats or when a large inoculation

98 Nonhemotropic Mycoplasma, Ureaplasma, and L‐Form Bacteria

dose was used. Natural Mycoplasma conjunctivitis pre­ dominantly occurs when infected cats are housed in groups or in kittens soon after weaning. The clinical signs observed may vary in severity and have been described as blepharospasm, conjunctival hyperemia, chemosis, and serous discharge followed by mucoid to sticky exudate. The cornea is usually not involved; how­ ever, M. felis and M. gateae have been occasionally asso­ ciated with ulcerative keratitis and/or keratomalacia.

disease. Clinical findings were fever, productive cough, and leukocytosis with a left shift. Radiographs fea­ tured alveolar and bronchointerstitial pulmonary den­ sities  and mild pleural effusion. Infection was fatal for some pups and necropsy findings were mucopuru­ lent airway exudates and hemorrhagic fibrinous necro­ tizing bronchopneumonia. M. cynos was abundantly demonstrated at neutrophilic inflammatory sites using immunohistochemistry.

Respiratory Mycoplasma Infections in Cats

Urogenital Mycoplasma Infections in Dogs

Mycoplasmas have been associated with chronic and acute feline upper respiratory tract disease (URTD) by epidemiologic evaluation, but available data are more inconclusive hypotheses than definite proofs of causality. Studies have also suggested that approximately 22% of cats with lower airway disease may have a concurrent Mycoplasma infection. Experimental challenge with M. felis induced pneumo­ nia in kittens, emphasizing its role as a primary pathogen in the lower respiratory tract. Naturally occurring infec­ tions with M. felis have been increasingly reported in the feline host associated with suppurative bronchitis and/or pneumonia. Radiographic or computed tomography (CT) findings may include diffuse interstitial pulmonary disease similar to that observed in adult human patients with M. pneumoniae pneumonia. Cats with concurrent pleural effusion show minimal signs of dyspnea and a nonodorous fluid is expected with uncomplicated Mycoplasma pyothorax.

Mycoplasmas are commonly present in the lower uro­ genital tract of dogs. They have occasionally been asso­ ciated with canine reproductive diseases including poor conception rates, early embryonic death, fetal resorption, abortion, weak pups, and neonatal death. M. canis and U. canigenitalium are the species most consistently associated with canine genital mycoplas­ mosis. Mycooplasma canis has been isolated from dogs urogenital disease and infertility, despite pro­ with ­ longed antibiotic therapy. It has also been cultured from the prostate, epididymis, and chronically inflamed bladder wall. Experimental infection with Mycoplasma canis produced urethritis, prostatitis or epididymitis in males and endometritis or metritis in females. In a sin­ gle study, U. canigenitalium was associated with infer­ tility in male dogs. Nevertheless, as conclusive evidence is lacking, further studies are required to establish whether M. canis or U. canigenitalium is linked to geni­ tal tract infections and infertility.

Respiratory Mycoplasma Infections in Dogs

Arthritis in Cats and Dogs Caused by Mycoplasmas

Mycoplasmas have been isolated from 78% of tracheo­ bronchial lavages of younger dogs with pulmonary dis­ ease but may also be present in the lower respiratory tract of healthy individuals (20–25%). Consequently, the role of certain canine Mycoplasma species as patho­ gens in canine pulmonary disease is still a subject of controversy. Experimental infection with M. canis, M. gateae, and M. spumans failed to produce pulmonary disease. However, M. cynos has been consistently asso­ ciated with canine respiratory diseases, including ­tracheobronchitis and bronchopneumonia, conditions that have been reproduced experimentally. A more recent study noted a correlation of M. cynos with canine infectious respiratory disease (CIRD) in younger dogs (6 months of age), adult, and older dogs. Cats of any breed can be affected by this infection. Disease is most common in adult or older cats [13]. History and Clinical Signs The clinical manifestations of this infection are very broad and variable, mainly due to the various types of immune responses which may be elicited in the dog in response to the parasite, the different organs affected and the different types of pathogenic mechanisms of disease that can be activated. Infection with L. infantum causes chronic infection, which can sometimes occur in a subclinical or mild self‐limiting illness, or as a moderate or severe illness in dogs which can be fatal. Four

clinical stages of the disease have been described based on the severity of the disease (clinical and laboratory abnormalities) and serology results, establishing different therapeutic regimens and forecast for each stage of the disease. The main clinical findings found on physical examination in classic CaNL include skin lesions, local or generalized lymphadenomegaly, loss of body weight, exercise intolerance, decreased appetite, lethargy, splenomegaly, polyuria and polydypsia, ocular lesions, epistaxis, onychogryposis, lameness, vomiting, and diarrhea [12,13]. Skin lesions are the most frequent manifestation of CaNL in dogs brought for treatment due to suspicion of the disease. Several dermatologic lesion patterns have been described: exfoliative dermatitis with alopecia which can be generalized or localized over the face, ears and limbs; ulcerative dermatitis; nodular dermatitis; mucocutaneous proliferative dermatitis; and papular dermatitis. The most common ocular manifestations of CaNL are anterior uveitis, blepharitis (exfoliative, ulcerative, or nodular), and keratoconjunctivitis, either common or sicca. About 25% of dogs with clinical leishmaniosis have ocular and periocular lesions including keratoconjunctivitis and uveitis. Ocular consequences of systemic hypertension such as retinal detachment and/or hemorrhages, retinal arterial tortuosity, and hyphema are present in the disease but not diagnosed frequently. Some degree of renal pathology is present in most dogs with CaNL and subsequent renal disease due to immune‐complex glomerulonephritis eventually develops and is believed to be the main cause of death in dogs with CaNL. The most common clinical signs and clinical‐pathologic abnormalities compatible with feline leishmaniosis (FeL) include lymph node enlargement and skin lesions such as ulcerative, crusty or nodular dermatitis mainly on the head or distal limbs, ocular lesions mainly with uveitis, feline chronic gingivostomatitis (FCGS) and mucocutaneous ulcerative or nodular lesions. Clinical illness is frequently associated with impaired immune competence, such as retroviral infections or immunosuppressive therapy. Diagnosis The most common laboratory findings in CaNL are hyperglobulinemia mainly with polyclonal gammaglobulinemia, hypoalbuminemia, decreased albumin:globulin ratio, mild to severe proteinuria, mild to moderate nonregenerative anemia, renal azotemia, elevated liver enzyme activities, thrombocytopenia, and thrombocy­ ormocytic topathy. Hypergammaglobulinemia and mild n normochromic anemia are commonly reported in FeL.

110  Protozoal and Protozoa‐Like Infections

The pathologic findings observed by cytology or histology in CaNL and FeL are macrophagic, neutrophilic‐ macrophagic and lymphoplasmacytic inflammation in affected tissues and reactive hyperplasia of lymphoid organs. The clinical diagnosis of leishmaniosis could be complex because of the broad spectrum of clinical and clinicopathologic abnormalities which are often not specific. For this reason, it is important to separate the diagnosis of Leishmania infection from clinical disease and to apply different diagnostic techniques for each situation [14]. Accurate diagnosis of CaNL and FeL often requires an integrated approach consisting of a clinicopathologic assessment and specific laboratory tests. The detection of L. infantum infection in dogs and cats includes parasitologic (cytology, histology, immunochemistry, and culture), molecular (conventional, nested, and real‐time PCR) and serologic methods (qualitative and quantitative antibody tests). Detection of specific serum antibodies to L. infantum should preferably be based on quantitative serologic techniques, such as the IFA and ELISA. The challenges of serology include cross‐reactivity with other pathogens such as other Leishmania and Trypanosoma species, and with antibodies elicited by vaccination against CaNL. High antibody levels are associated with severe parasitism and moderate to severe disease. However, the presence of low antibody levels is not necessarily indicative of disease and further work‐up is necessary to confirm CaNL and FeL by other diagnostic methods such as cytology, histopathology, and PCR. Information provided by PCR should not be separated from the data obtained from clinicopathologic and serologic evaluations. These should all be combined for a comprehensive assessment. It is essential to know the terms and limitations of each diagnostic test and make an adequate clinical interpretation [12–14]. Therapy Treatment for CaNL depends on the severity of disease. Different therapeutic regimens have been established for each stage of CaNL. The drugs most frequently used in moderate to severe CaNL (stage 2 and 3) are a combination of meglumine antimoniate at 75–100 mg/kg q24h or 40–75 mg/kg BID for four weeks SC and allopurinol at 10 mg/kg/BID/PO for at least one year. Alternatively, a combination of miltefosine at 2 mg/kg/ PO q24h for four weeks and allopurinol at the same dose described above is also employed. Other drugs have been proposed for the treatment of CaNL but their efficacy or safety are limited. Clinical cure in CaNL is often obtained and is associated with reduction in the parasite load and infectiousness to sand flies, but infection may persist and clinical

relapse might occur and therefore life‐long follow‐up post therapy should be maintained. The most common drug used for FeL is allopurinol administered alone at a dose of 10 mg/kg/BID/PO for at least six months. Treatment with meglumine antimoniate alone has been reported in rare cases of FeL. Most cats recover clinically following prolonged treatment and as for dogs, follow‐up with routine laboratory tests, serology, and PCR is essential for the prevention of clinical relapses [12–14]. Prognosis The prognosis of dogs and cats with leishmaniosis depends mainly on the severity of disease. Different prognosis is expected for each stage of CaNL with generally poor prognosis observed in dogs with renal insufficiency or severe renal disease associated with profuse proteinuria. The prognosis in cats is generally fair and a poorer prognosis is associated mainly with the presence of concomitant diseases and immune suppression. Prevention Several preventive measures are available for CaNL. The use of individual topical pyrethroid insecticides in collars or spot‐on formulation has been shown to be effective in reducing L. infantum infection and, therefore, they are commonly applied to dogs in endemic areas to prevent infection. Vaccination is also considered a promising tool for controlling CaNL. Second‐generation vaccines for CaNL are commercially available in Europe and Brazil, with different efficacy rates in reducing the incidence of the clinical illness in field. Domperidone, a hyperprolactinemic immunomodulatory drug, has recently been licensed in some European countries for preventive use against CanL. However, limited studies exist regarding its efficacy in preventing the disease. Dog culling practiced in Brazil as a control measure has been reported to have little impact on the incidence of human and canine leishmaniosis [15]. To date, preventive measures for this infection in cats are not commercially available.

­Hepatozoonosis Etiology and Pathophysiology Hepatozoonosis is an arthropod‐borne infection caused by apicomplexan protozoa of the family Hepatozoidae which are phylogenetically related to the piroplasms (Babesia, Theileria, and Cytauxzoon). Two different ­species of Hepatozoon infect dogs: Hepatozoon canis globally and Hepatozoon americanum in some parts of

1011

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Section 9  Infectious Disease

the United States, mainly the southeastern states. Hepatozoon canis infects the hemolymphatic tissues and causes anemia and lethargy whereas H. americanum primarily infects muscular tissues and induces severe myositis and lameness [16,17]. Feline hepatozoonosis is caused by H. felis, which is prevalent in the Mediterranean basin, Asia, Africa, South, Central and North America. It infects muscle tissues including the myocardium and has not been associated with severe clinical disease [18]. Recently, H. canis and H. silvestris infections have been described in cats in Europe. In contrast to most tick‐borne pathogens which are transmitted via the tick salivary glands, Hepatozoon spp. infect vertebrates when they ingest arthropod hosts containing infective sporozoites. The main vectors of H. canis are the ticks Rhipicephalus sanguineus and R. turanicus worldwide and Amblyomma ovale in South America, whereas the Gulf Coast tick Amblyomma maculatum is the vector of H. americanum in North America. The vector of H. felis has not been identified to date but transplacental transmission in the feline host has been shown. Transplacental transmission from dam to pups has also been demonstrated for H. canis infection but not for H. americanum. Transmission by carnivorism and predation of the canine host on intermediate or wildlife transport hosts with ingestion of parasite cyst forms from their tissues has been shown for H. americanum and is considered as a main route of dog infection with this parasite [19]. The life cycle of H. canis revolves between the canine and tick hosts. When dogs ingest the vector tick or tick parts, H. canis sporozoites are released from sporocysts in the intestine and penetrate the gut wall. The sporozoites invade mononuclear cells and disseminate hematogenously or via the lymph to hemolymphatic target organs that include the bone marrow, spleen, and lymph nodes and to other internal organs such as the liver, kidney, and lungs. Meronts in which asexually dividing merozoites develop are formed in the dog’s tissues in the process of merogony and may cause cellular necrosis in tissues. Merozoites release from mature meronts, invade neutrophils and monocytes, and develop into gamonts. Alternatively, merozoites can produce secondary meronts in the target tissues. The tick, which serves as the definitive host, is infected by ingesting leukocytes containing gamonts when feeding on a parasitemic dog. Morphologically indistinguishable male and female H. canis gamonts are released from the dog leukocytes within the tick gut, associate in syzygy, and differentiate in the process of gametogenesis to distinct gametes. After fertilization, the zygote divides and sporogony takes place with the formation of oocysts that release into the tick’s hemocoel. The oocysts are large, spherical forms consisting of a membrane that envelops multiple sporocysts in which the infective sporozoites are found.

The life cycle of H. americanum differs from that of H. canis particularly with regard to the parasite’s target tissues in the dog, the formation of distinct large cysts and the inflammatory reaction formed in response to infection. Sporocysts of H. americanum sporozoites from the hemocoel of an infected vector tick ingested by dogs release infectious sporozoites following contact with bile in the gut. Sporozoites are thought to cross the gut wall, and then taken up by host macrophages and transmitted via the lymph system or hematogenously to the target organs consisting of skeletal and cardiac muscles. Hepatozoon americanum‐infected macrophages lodge between striated muscle fibers in these tissues and form muscle cysts. Concentric layers of mucopolysaccharide material are deposited by the infected cell forming a cystic structure referred to as “onion skin cyst.” The encysted zoite undergoes merogony and multiple merozoites are formed. Eventually, the cyst ruptures, releasing mature individual merozoites into the surrounding tissue and eliciting a strong local inflammatory response. The pyogranulomatous inflammatory response is associated with severe musculoskeletal pain. The merozoites are taken up by leukocytes and are thought to undergo additional merogonic cycles or enter the blood vasculature, develop into gamonts and circulate in monocytes. Epidemiology Hepatozoon canis infection is reported mainly from tropical, subtropical, and temperate climate regions where vector tick species are abundant. Infection with H. canis has also been reported in dogs in the southeastern United States, mainly in some regions where H. americanum is also prevalent, and co‐infections of both species have been reported. In Europe, autochthonous dog infection is found mostly close to the Mediterranean basin. It has also been described in dogs imported into nonendemic countries such as Germany, in foxes in European countries where autochthonous canine infection is rare or has not been reported, and in Slovakia and Poland where R. sanguineus, its tick vector, is not present. The prevalence of H. canis infection differs between regions. The rates of infection based on blood smear examination or PCR range from 1% to 64%. The presence of H. americanum infection in the United States has expanded north and east from Texas, where it was first detected in 1978. It is found in several southeastern states, and occasionally cases are reported in other locations including Nebraska, Vermont, Washington, and California. Hepatozoon americanum infects wild coyotes (Canis latrans) and in experimentally infected young coyotes, it causes similar disease to that seen in dogs. Surveys of free‐ranging coyotes in Oklahoma showed that 40–50% were infected naturally with H. americanum.

110  Protozoal and Protozoa‐Like Infections

Signalment Infection with Hepatozoon species is more common in dogs and cats with frequent outdoor access and those with contact to tick vectors. Hepatozoon americanum is often detected in hunting and guard dogs in contact with prey and potentially infected wild mammals. No age or breed predilections have been reported. The pathogenesis of H. canis infection is influenced by immunodeficient conditions including immunosuppressive treatment, an immature immune system in young pups, concurrent infectious diseases or debilitating concomitant diseases. Co‐infections with Toxoplasma, Leishmania, Babesia, Ehrlichia canis, or viral infections might predispose to clinical illness. History and Clinical Signs Hepatozoon canis infection varies from being subclinical to a severe disease in which dogs present with lethargy, fever, cachexia, and pale mucous membranes due to anemia. In contrast to the generally mild disease usually found in H. canis infection, dogs diagnosed with H. americanum infection are presented with fever, gait abnormalities, muscular pain induced by myositis, generalized muscular atrophy and mucopurulent ocular discharge with decreased tear production associated with inflammation of the extraocular muscles. Clinical signs may start 4–6 weeks after infection due to the pyogranulomatous inflammatory response that occurs when the  encysted parasite cyst ruptures in muscle tissue. American canine hepatozoonosis can have an acute presentation or a waxing and waning chronic pattern. Pain associated with disease may be generalized or localized in the lumbar and cervical spine, long bones, or joints. Gait abnormalities range from limb stiffness to complete recumbency and inability to rise. Chronic infections can be associated with polydipsia and polyuria due to renal amyloidosis or glomerulonephritis and an ophthalmic examination may reveal uveitis. Hepatozoon felis infection has not been associated directly with overt clinical manifestations but H. silvestris has been reported to cause severe disease in domestic cats. Diagnosis Subclinical infection or mild disease is the most common presentation of H. canis infection and it is usually found in dogs with a low level of parasitemia (1–5%) (Figure  110.4). Severe illness can be found in dogs and may be associated with white blood cell concentrations as high as 150 × 109 leukocytes/L. High parasitemia rates of up to 100% can accompany extreme neutrophilia. A case–control study of dogs with H. canis parasitemia

Figure 110.4  Hepatozoon canis gamont in a neutrophil from the blood smear of a naturally infected dog (May Grunwald–Giemsa stain 1000× magnification).

indicated that 15% had a high number of circulating ­ arasites (>800 gamonts/μL) accompanied by elevated p body temperature, lethargy, weight loss, anemia, and hyperglobulinemia. An association between the severity of clinical signs and the level of parasitemia was also found in a study of canine hepatozoonosis from Turkey. The most frequent hematologic abnormality reported in H. canis infection is mild to moderate normocytic normochromic nonregenerative anemia and, more rarely, regenerative anemia. Abnormalities in serum chemistry include hyperproteinemia with hyperglobulinemia due to polyclonal gammopathy, hypoalbuminemia and increased creatine kinase and alkaline phosphatase activities. Meronts of H. canis are commonly found in the spleen, bone marrow, and lymph nodes by cytology and histopathology. Although lesions of the periosteum are more frequently associated with H. americanum infection, periostitis has also been described as a rare finding in H. canis infection. A marked neutrophilia is also one of the consistent hematologic findings in H. americanum infection with leukocyte counts of 30–200 × 109/L blood, although the level of parasitaemia is typically very low. A mild to moderate normocytic normochromic nonregenerative anemia is frequent. Serum biochemical abnormalities include increased alkaline phosphatase activity and hypoalbuminemia. Histopathology of skeletal muscles from dogs with H. americanum infections reveals pyogranulomatous myositis and large round to oval “onion skin” cysts (250–500 micrometer diameter). Radiography of the long bones or pelvis demonstrating periosteal proliferation can be used for screening suspected animals. Hepatozoon canis infection is usually diagnosed by microscopic detection of intracellular H. canis gamonts in stained blood smears. Gamonts can be detected in the

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blood at 28 days post infection. The gamonts are found in the cytoplasm of neutrophils and monocytes, have an ellipsoidal shape and are about 11 ×4 micrometers. In contrast, H. americanum parasitaemia usually does not exceed 0.1% of the leukocytes and rare gamonts may be detected in the blood from 32 days post infection. The confirmation of H. americanum infection can be achieved by muscle biopsy and demonstration of typical parasite cysts and granulomas or by PCR. PCR of blood for H. canis or H. americanum is a sensitive and specific diagnostic option. Diagnostic real‐time PCR assays that can also quantify the parasitic load are able to detect both H. americanum and H. canis and distinguish between the two species. Hepatozoon felis infection is usually associated with a low parasitaemia and is best detected by blood PCR. Serologic testing for H. canis or H. americanum by an IFA test or ELISA is used in epidemiologic studies but not commonly employed for clinical diagnosis in practice. Therapy The current treatment for H. canis is the administration of imidocarb dipropionate at 5–6 mg/kg IM or SC every 14 days until gamonts are no longer present in blood smears. The decrease of parasitemia is slow and usually requires several imidocarb treatments. Studies with follow‐up by evaluation of buffy coat smears and sensitive PCR indicated that complete elimination of the parasite may frequently not be achieved with imidocarb dipropionate alone and also  with combinations of imidocarb dipropionate and toltrazuril/emodepside, or imidocarb dipripionate with clindamycin [20]. Hepatozoon americanum is treated with a combination oral therapy of trimethoprim‐sulfadiazine (15 mg/ kg q12h), pyrimethamine (0.25 mg/kg q24h), and clindamycin (10 mg/kg q8h) for 14 days. Ponazuril (10 mg/kg PO q12h for 14–28 days) has been suggested as an alternative to the combination antiprotozoal treatment. After initial improvement of clinical disease signs is obtained, remission can be prolonged with the oral administration of the coccidiostat decoquinate at 15 mg/kg mixed in food q12h for two years. Relapse of clinical disease is frequent following discontinuation of treatment. Supportive therapy with nonsteroidal antiinflammatory drugs is effective in relieving pain and fever in dogs with H. americanum infection and administered initially with the combination antiprotozoal therapy. No controlled trials have been published on the treatment of feline hepatozoonosis. Single cases have been treated with doxycycline, a single dose of 2 mg/kg primaquine PO or imidocarb at 2.5 m/kg IM with undetermined efficacy in eliminating the infection.

Prognosis The prognosis of treated dogs with a low H. canis parasitemia is generally good even if decrease of parasitemia is slow and requires several imidocarb treatments. The prognosis for dogs with high parasitemia is good to guarded and sometimes associated with the outcome of a concurrent illness. Hepatozoon americanum infection may be fatal without treatment within several months. Prognosis with antiprotozoal treatment followed up by long‐term decoquinate therapy is guarded. Prevention Prevention of both H. canis and H. americanum infections consists of the use of topical acaricides and environmental parasiticides. Furthermore, avoidance of tick ingestion while scavenging or grooming, and of eating raw prey meat potentially harboring infectious tissue stages of the parasite is recommended [16]. To date, hepatozoonosis has not been shown to be established in recipient dogs following transfusion. Transfusion of the blood gamont stage is not likely to result in establishment of infection in the recipient dog as the gamont stage found in blood has to continue to the sexual parts of the parasite’s life cycle in the tick before the parasite replicates.

­Feline Cytauxzoonosis Etiology and Pathophysiology Feline cytauxzoonosis is an emerging tick‐borne disease with an expanding geographic distribution caused by parasites from the genus Cytauxzoon (phylum Apicomplexa, class Piroplasmea, order Piroplasmida). Parasites of the genus Cytauxzoon are closely related to those of the genera Babesia and Theileria. Cytauxzoon felis is a species infecting domestic cats and wild felids in the United States and South America and detailed clinical, parasitologic, and epidemiologic information is available on this infection. Other Cytauxzoon parasites which are genetically different and of decreased pathogenicity have been described in Europe in domestic cats and wild felids but limited ­information is available on these species. Cytauxzoon felis is transmitted by the tick Amblyomma americanum in the United States. Amblyomma cajennense and other ixodid ticks have been presumed as vectors in South America. Other routes of transmission have not been well studied for this infection. The tick vector for Cytauxzoon spp. infection in felids in Europe remains unknown as well as other possible modes of transmission.

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The wildlife animal reservoir for C. felis infection in the United States is the bobcat (Lynx rufus) where infection is mostly subclinical and a persistent erythroparasitemia is commonly found. Conversely, clinical illness is frequently found in infected domestic cats. Cytauxzoon felis infection in the cat varies in its clinical manifestation from a severe acute fatal disease to subclinical chronic infection. Cytauxzoon species are detectable in two main forms in the mammalian host: schizonts in histiocytic cells and merozoites in erythrocytes. A phase of asexual schizogenous reproduction, within the host’s mononuclear phagocytic cells, is responsible for the pathologic processes resulting in clinical illness. Large schizonts can occlude small blood vessels and cause thrombosis in the visceral organs and the brain. The onset of clinical disease occurs usually 1–3 weeks after tick‐transmitted infection and the entire disease course is rapid, with cats frequently succumbing to infection within days. Epidemiology The geographic distribution and seasonality of cytauxzoonosis due to C. felis is largely correlated with relevant tick vector activity and presence of bobcats. This infection is present in the south‐central, southeastern, and mid‐ Atlantic regions of the United States. Infected domestic cats are commonly from rural or suburban wooded areas. The prevalence of C. felis infection in domestic cats is ­variable in different published studies. In healthy cats, prevalence of C. felis infection ranged from 0.3% to 28%. Cytauxzoonosis accounted for 1.5% of all feline admissions to the veterinary teaching hospital at Oklahoma State University between 1998 and 2006 [21–23]. Signalment Disease due to C. felis can occur in cats of any age and either gender with no identified breed predilections. Likewise, no signalment predispositions have been found in feline infections due to other species of Cytauxzoon. History and Clinical Signs Outdoor cats are more likely to be infected during spring and summer due to the presence of tick vector. Immune suppression has not been found to be a risk factor for infection with C. felis. Cytauxzoon spp. infection has been associated with living in a cat colony and outdoor lifestyle in domestic cats from Europe. The clinical signs of C. felis infection are nonspecific and include an acute onset of anorexia, lethargy, and fever. Other clinical signs include increased vocalization, weakness, dehydration, icteric or pale mucous mem-

branes, pigmenturia, mild to moderate lymphadenomegaly, splenomegaly and hepatomegaly, generalized pain, tachypnea and tachycardia with or without respiratory distress, dull mentation, and seizures. Cytauxzoon spp. appear to mainly cause a subclinical infection with evidence of erythroparasitemia in domestic and wild felids in Europe. So far, evidence of clinical manifestations of a tissue schizogonic phase, as found for C. felis, has not been observed in Cytauxzoon spp. infection in Europe. Diagnosis Laboratory abnormalities due to C. felis infection include pancytopenia, hyperbilirubinemia, mild to moderate normocytic normochromic nonregenerative anemia, thrombocytopenia, lymphopenia, neutropenia, mild to moderate elevation of liver enzyme activities (ALT and ALP), prerenal azotemia, electrolyte disturbances (hypocalcemia, hypokalemia, and hyponatremia), hyperglycemia, mild hypoproteinemia with hypoalbuminemia and hypocholesterolemia. Detection of Cytauxzoon in stained blood smears is useful in the diagnosis of erythroparasitemia. Infection may also be visualized by microscopic identification of schizont‐laden macrophages in cytology of fine needle aspirates of lymph nodes, spleen or liver. PCR is a valuable tool to detect low parasitemia with higher sensitivity than blood smear. Furthermore, PCR and genetic characterization can distinguish between infection with Cytauxzoon and small Babesia spp. Serology is not used in the clinical setting. Therapy The combination of the antimalarial atovaquone (15 mg/ kg PO q8h) and the macrolide azithromycin (10 mg/kg PO q24h) for 10 days is currently considered the best treatment option for acute cytauxzoonosis due to C. felis, with an approximately 60% survival rate in sick cats, while other treatments such as imidocarb dipropionate (3.5–5 mg/kg IM repeated at 14‐day intervals if needed) have been reported to result in lower survival rates [23]. Medical management of this disease requires supportive treatments. Prognosis The prognosis for acute cytauxzoonosis in domestic cats due to C. felis is guarded to poor. Prevention So far, there is no vaccine commercially available for the prevention of Cytauxzoon infection. Prevention is focused

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on avoidance of tick bites. A combination of tick control that targets the specific tick vectors and indoor lifestyle is the best strategy to reduce tick exposure.

­Babesiosis Etiology and Pathophysiology Babesia are tick‐borne protozoan parasites of erythrocytes that infect domestic and wild animals, and humans. Babesia belongs to the phylum Apicomplexa, class Piroplasmea, and order Piroplasmida [24]. Canine infections caused by different Babesia species are protozoal tick‐borne diseases with worldwide distribution and global significance. Large‐form Babesia spp. include Babesia rossi, B. canis, and B. vogeli. These species were previously considered subspecies of B. canis. They are identical morphologically but differ in the severity of clinical manifestations which they cause, their tick vectors, genetic characteristics, and geographic distribution. Therefore, they are currently considered separate species. Another yet unnamed large Babesia species most closely related to B. bigemina was found to infect immunocompromised dogs in the United States. The small Babesia spp. include B. gibsoni, B. conradae, and B. vulpes (formerly referred to as B. microti‐like piroplasm and Theileria annae). Babesiosis of domestic cats has mostly been reported in South Africa where infection is mainly due to B. felis, a small Babesia species that causes anemia and icterus. In addition, B. cati was reported from a cat in India and sporadic cases of infection in domestic cats by unnamed Babesia parasites have been reported in France, Germany, Thailand, and Zimbabwe. B. canis infection was reported from three cats in a study from Spain and Portugal in domestic cats with molecular evidence for infection. Additionally, infection of domestic cats with a genetically distinct species related to B. canis and named B. canis subsp. presentii was described in Israel [25]. Dogs and cats are infected when Babesia sporozoites are injected with saliva into the host’s skin during a blood meal. The parasites invade the erythrocytes and form ring‐shaped trophozoites which replicate within the erythrocyte and form merozoites observed as pairs of attached pear‐shaped parasites in some Babesia species. Merozoites may further divide, forming eight or more parasites in the same erythrocyte and eventually lysing the cell, discharging into the blood to invade additional erythrocytes. Ticks feeding on infected blood take up merozoites and sexual parasite development in the tick gut is followed by sporogony in its salivary gland or ovaries. The parasite can be transmitted from the tick salivary glands during a tick bite or transovarially through its oocytes, as found in some Babesia species.

Babesia gibsoni infection has also been shown to be transmitted via blood transfusion and transplacentally. Furthermore, several studies have provided evidence that B. gibsoni is likely transmitted directly from dog to dog via bite wounds, saliva, or ingested blood [25,26]. Epidemiology The geographic distribution of the causative agents and thus the occurrence of babesiosis is largely dependent on the habitat of tick vector species, with the exception of B. gibsoni where evidence for dog‐to‐dog transmission indicates that infection can be transmitted among fighting dog breeds independently of the limitations of vector tick infestation. Babesia vogeli and B. gibsoni have worldwide distribution, whereas B. rossi and B. canis have been mostly restricted to Africa and Europe, respectively. The unnamed large Babesia sp. most closely related to B. bigemina and B. conradae has been reported only from North America whereas B. vulpes was reported in Europe and North America. Signalment Puppies and dogs of any age with concomitant disease or immune suppression suffer clinical babesiosis due to B. vogeli more frequently compared to healthy adult dogs. Breed and gender do not appear to be predisposing factors. Babesia canis, B. rossi, and B. vulpes have been reported to more frequently affect young to adult outdoor hunting and shepherd dogs. Breed and gender do not appear to be predisposing factors. Disease due to B. gibsoni is found mostly in fighting dogs and breeds such as the pit bull terrier and tosa. Gender and age do not appear to be predisposing factors. Gender, breed, and age have also not been reported to be predisposing factors for B. conradae infection. Splenectomized and immune‐compromised dogs are more susceptible to infection with any Babesia species. History and Clinical Signs The main clinical findings reported in dogs suffering from B. canis infection include dehydration, lethargy, anorexia and fever, Babesia vogeli usually causes a subclinical infection or mild to moderate clinical disease which often accompanies other concomitant diseases or immunosuppressive conditions or affects splenectomized dogs. Severe fatal hemolytic anemia has been reported in puppies. The most virulent large‐form Babesia is B. rossi. Babesiosis due to B. rossi can manifest with uncomplicated common clinical signs as described for other babesial species or with complicated clinical disorders

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including acute renal failure with anuria, icterus, hypotension, acute respiratory distress syndrome (ARDS), vomiting, diarrhea, pancreatitis, myalgia, rhabdomyolysis, ascites, pulmonary edema, encephalomyelitis, and peracute shock. The clinical findings associated with B. gibsoni infection include fever, splenomegaly, lymphadenomegaly, hepatomegaly, and lethargy. Babesia conradae infection has been described as more virulent than B. gibsoni infection, resulting in higher rates of mortality. The most common clinical findings reported in dogs infected with B. vulpes from the northwest of Spain include fever, lethargy, weakness, and pigmenturia. Diagnosis The main clinicopathologic findings reported in dogs suffering from B. canis infection include mild to severe thrombocytopenia, hyperfibrinogenemia, mild to moderate normocytic-normochromic nonregenerative anemia, hemolysis and neutropenia. Hemoglobinuria has also been described in  naturally infected dogs. Common laboratory abnormalities in dogs infected with B. vogeli, B. gibsoni and B. conradae are regenerative anemia and thrombocytopenia. Although they are both small Babesia species, more pronounced anemia occurs in B. conradae infection than with B. gibsoni infection. Dogs with B. vulpes infections in northern Spain have demonstrated moderate to severe regenerative anemia, thrombocytopenia and azotemia. Hyperglobulinemia and proteinuria can also occur in Babesia infected dogs. The hemolytic anemia and thrombocytopenia associated with Babesia infections appear to occur by multiple mechanisms including immune-mediated destruction. Positive Coombs’ testing, spherocytosis, positive saline agglutination tests and anti-platelet antibodies can occur. Therefore, it is important to rule-out babesiosis in patients with suspected idiopathic immune-mediated hemolytic anemia and thrombocytopenia. Detection of large or small species of Babesia in stained blood smears has been the standard for diagnosis for many years. This method is reliable when a moderate to high parasitemia is present, but there is not always a correlation between the level of parasitemia and the severity of clinical signs. A fresh smear is recommended for the accurate diagnosis of infection. Serology can indicate past infection or a present persistent one. Some serologic cross‐reactivity exists between Babesia species. False‐negative results are likely in peracute or acute infection and therefore convalescent antibody titers are needed to prove acute infection. Some evidence suggests that seroconversion may not occur in immunocompromised or young animals. The PCR is particularly useful in the diagnosis of babesiosis in dogs with a low parasitemia level including suspected carrier dogs or chronically infected animals and for speciation of parasites. However, a negative PCR

does not rule out infection. Several molecular methods including seminested PCR, reverse line blotting (RLB), and restriction fragment length polymorphism (RFLP) PCR permit discrimination between species. Therapy Large Babesia spp. infections of dogs and cats are commonly treated with one dose of imidocarb dipropionate at 5–6 mg/kg (dog) and 2.5 mg/kg (cat) IM or SC with good clinical response. Babesia gibsoni and B. conradae infections are often resistant to imidocarb dipropionate and diminazene aceturate. The treatment of choice for these small Babesia species is a combination of the antimalarial atovaquone and the macrolide azithromycin. This combination is also likely to be a good treatment option for B. vulpes infection. The most commonly used dose of atovaquone is 13.5 mg/kg, administered PO q8h with fatty food to maximize drug absorption, in combination with azithromycin at 10 mg/kg PO for 10 days. Clindamycin in combination with metronidazole and doxycycline for a minimum of three months has also been used for treatment of B. gibsoni, although its overall efficacy is uncertain. The use of clindamycin alone is not recommended by some authors given the potential for selection of resistance and may subsequently interfere with alternative treatments. The treatment of choice for B. felis in cats is ­primaquine phosphate. Published dosages range from 0.5 to 1.0 mg/ kg PO, IV, or IM once, or administered daily on three consecutive days. Clinical and parasitologic cure are commonly not achieved in small babesial spp. infections in dogs and cats and clinical relapses may occur frequently. Medical management of infection may ­ require supportive treatments including blood transfusions, intravenous fluids, and the use of antiinflammatory drugs [24–27]. Prognosis The prognosis of dogs and cats infected with large forms of Babesia is generally good with appropriate therapy. Canine infection with small Babesia spp. may be more resistant to treatment and have a poorer prognosis. Prevention The prevention of babesiosis relies mostly on topical and environmental acaricidal treatments aimed at reducing exposure to vector ticks and transmission of the pathogen to dogs and cats. As Babesia species are transmitted by blood product transfusions, it is recommended to screen canine blood donors for Babesia infection on a regular basis and consider screening cat blood in relevant areas. Nonvectorial dog‐to‐dog transmission of

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babesiae by fighting is preventable and can be responsible for the spread of babesiosis into previously nonendemic areas. A vaccine against Babesia is commercially available in some countries in Europe.

­Giardiasis Etiology and Pathophysiology The genus Giardia includes intestinal flagellated protozoan parasites that infect a large number of host species, ranging from mammals to amphibians and birds. The most important Giardia species, G. duodenalis, infects cats, dogs, livestock, and humans with a broad host range and considerable public health significance. Giardia duodenalis is currently considered a multispecies complex. Multigeneic sequence analyses have identified different genetic assemblages of G. duodenalis including assemblages A and B in man and animal species, assemblages C and D in dogs, assemblage E from ruminants and pigs, assemblage F from cats, and assemblage G from rodents. To date, no association has been made regarding specific assemblages and the induction of ­certain characteristic clinical signs or the degree of their severity [2,3]. The life cycle of Giardia is direct and has two stages: the trophozoite and a cyst. The infective stage of the parasite, the cyst, is encysted when released into the feces and is immediately infectious. Cysts remain infectious for months in cool, damp conditions and accumulate in the environment. When ingested by the host, cysts excyst in the duodenum, releasing motile trophozoites. The latter undergo repeated mitotic division and form environmentally resistant cysts. Cysts pass through the intestine in feces and are spread by contaminated water, food, and fomites, and by direct contact. Giardia is transmitted mostly by ingestion of cysts in contaminated food or water. Following the ingestion of cysts and their exposure to the gut environment, each cyst releases two trophozoites which attach to the villi of the small intestinal epithelium. Trophozoites multiply by binary fission and eventually form cysts which are shed in the feces. The parasite presence in the intestine induces diarrhea by hypersecretion and malabsorption with increased intestinal permeability. Giardia infection in most dogs and cats is subclinical, but some animals suffer from severe diarrhea and weight loss which can become chronic. Epidemiology Infection rates with Giardia in dogs and cats vary in the 5–30% range. The infection rates found in a large multicountry study in Europe were 24.8% in dogs and 20.3% in

cats. Shedding Giardia is often subclinical and has frequently not been associated with clinical signs in population surveys. However, diarrhea and illness due to Giardia infection have been associated with puppies and kittens of young age as well as immune‐suppressed animals and pets living in crowded conditions. The relationship between human and animal Giardia infection is not clear. Humans certainly infect each other frequently and although they share the same G. duodenalis assemblages with animals with whom they have close contact, such as household dogs, it is not known how frequently infection is actually acquired from household animal contact or whether both human and pets acquire it from a common source, such as contaminated water [2]. Signalment There is no breed or gender predilection. Young, immune‐suppressed animals or those living in crowded conditions are more predisposed to the disease. History and Clinical Signs Most feline and canine Giardia shedders are infected subclinically. Those affected clinically suffer primarily from diarrhea and weight loss if disease becomes chronic. Diarrhea can be acute, intermittent or chronic and is mucoid, soft to watery and rarely bloody. Diarrhea can worsen and become bloody in animals co‐infected with other intestinal parasites such as Ancylostoma spp., Coccidia or Tritrichomonas foetus. Vomiting and fever are rare. Reinfection with Giardia after successful treatment may occur. Diagnosis Giardia infection can be diagnosed by stool examination to identify cyst and trophozoite stages in direct fresh stool smears or by flotation for cysts. Sheather’s sugar centrifugation and zinc sulfate centrifugation can be used for concentrating cysts. Rapid detection of Giardia antigen can be made using immunochromatographic kits, or by immunofluorescence, ELISA, and PCR in the parasitology laboratory [28]. Therapy Dogs and cats can be treated with febendazole at 50 mg/ kg PO q24h for 3–5 days, or with combined febantel, pyrantel, and praziquantel according to the febantel component at 37.8 mg/kg PO q24h for 3–5 days, or with metronidazole at 15–25 mg/kg PO q12–24h for 5–7 days. Since Giardia strains may have different antiprotozoal

110  Protozoal and Protozoa‐Like Infections

susceptibilities, treatment with one of these drug options should be replaced by another if it is not successful in effectively relieving clinical signs. Furthermore, co‐infections should be diagnosed and treated and sanitary management with quaternary ammonium disinfectants should be instituted to prevent reinfection. Prognosis The prognosis of treated dogs and cats is usually good and clinical signs are ameliorated, although some animals could suffer from persistent or recurrent infection. Prevention Disinfection of kennels, cages, and the animal’s direct environment, boiling or filtering drinking water and removal of feces are important for controlling the spread of infection among animals that live in close proximity. Bathing of dogs and cats in infected kennels may decrease transmission of giardiasis between individual animals.

­Miscellaneous Infections Amebiasis Amoebas are unicellular protozoal organisms with motile cytoplasm and a flexible cell wall. Molecular phylogenetic studies have shown that amoebas do not form a single taxonomic group and are found in many lineages of eukaryotic organisms. In older nomenclature classifications, most amoebas were placed in the class or subphylum Sarcodina, a group of single‐celled motile organisms that possess pseudopods or move by protoplasmic flow. However, the amoeboid organisms are not classified together in one group any longer. Amoebas can be classified as enteric and residing in the gastrointestinal tract or nonenteric and associated with visceral and CNS invasion. Amoebas of different species are a rare cause of disease in dogs and cats. Different species of Acanthamoeba, Balamuthia, Hartmannella, and other amoebic genera and species may infect dogs and cats and colonize various organs, causing pneumonia, meningoencephalitis, renal disease, gastrointestinal disease of the small or large bowel, dermatitis, nasal disease, and keratitis [29,30]. Affected cats and dogs may be febrile and show a variety of clinical signs including neurologic abnormalities. Diagnosis is based on finding amoebae in excretions as cysts or trophozoites, or in tissues by cytology or histopathology and genetic characterization by specific PCR assays. Treatment consists of metronidazole for enteric infection and other antiprotozoal and antifungal drugs

such as amphotericin B, ketoconazole, and miltefosine for deep organ infection. Trichomoniasis Trichomonas spp. are flagellated protozoa that belong to the order Trichomonadida, reproduce by binary fission, and do not produce cysts. Cats and more rarely dogs can be infected with Tritrichomonas foetus which is transmitted by the direct fecal–oral route and causes diarrhea [31,32]. Tritrichomonas foetus survives in feces and humid environment outside the animal’s body for up to a few days. The parasite has been reported in numerous countries and has a worldwide distribution, with infection rates of up to 32% in cat populations from different countries in Europe [32]. Subclinical infection with T. foetus is common and there is not a direct relationship between infection and clinical signs. Infection is more common in cats from multicat settings and breeding catteries and clinical signs are more common in young cats under 12 months of age. Tritrichomonas foetus attaches to the intestinal epithelium and induces a lymphoplamsmacytic and neutrophilic inflammatory response with large bowel diarrhea and frequent passage of soft liquid feces which may contain mucus and fresh blood [33]. Irritation of the anus may be present with swelling and edema. Diagnosis is made by identification of the organism in fresh feces by direct microscopic examination in which typical forward movement directed by the parasite’s flagella and undulating membrane can be detected. Stained thin fecal smears may aid in the detection of the parasite with its typical form which is different from the Giardia trophozoites (Figure  110.5), when its presence is suspected

Figure 110.5  Tritrichomonas foetus in a stained fecal smear from a naturally infected cat (May Grunwald–Giemsa stain 1000× magnification).

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based on wet smear microscopy. PCR is able to diagnose infection with high sensitivity and specificity, and culture in an “in pouch” medium is also used for diagnostic purposes. The drug of choice for treatment of T. foetus in cats is ronidazole at 30 mg/kg PO q24h for 14 days. Ronidazole is neurotoxic to cats at higher doses and is also teratogenic and should not be used to treat pregnant or lactating queens or young kittens. Pneumocystosis Pneumocystis is a genus of pathogens most closely related to Ascomycetes fungi. Current research has indicated that Pneumocystis species appear to be host specific. Five species have been described, including Pneumocystis jirovecii which infects humans and P. carinii which is a pathogen of rats, although much of the previous scientific literature has related to P. carinii as a human pathogen. Dogs may be infected with a different Pneumocystis species or strain provisionally termed P. canis. Pneumocystis causes pneumonia in dogs and its transmission is thought to be air‐borne in inhaled respiratory aerosol from infected hosts. Young dogs less than 1 year of age of either gender may be infected and Cavalier King Charles spaniels, Shetland sheepdogs and miniature dachshunds are predisposed to disease.

Immunoglobulin deficiency and other congenital or acquired states of immune suppression have been reported in infected dogs [34]. Trophozoites and cysts of Pneumocystis are found in the lung alveolar spaces where the pathogen and its associated inflammatory response damage the alveolar septa. Dogs with pneumocystosis suffer from respiratory distress, tachypnea, weight loss, tachycardia, and exercise intolerance which progress and worsen over weeks to months. Thoracic radiographs show diffuse interstitial to ­alveolar patterns of lung disease and fluid collected on bronchoalveolar lavage or lung aspirates may show Pneumocystis trophozoites and cysts by cytology. Lung histopathology and PCR are also helpful in confirming infection. Pneumocystis does not grow in fungal culture and serologic tests have not provided a reliable diagnosis for canine infection. The antimicrobial treatment of choice for canine pneumocystosis is trimethoprim‐sulfonamide at 15 mg/ kg PO q8h for three weeks. Pentamidine isethionate at 4 mg/kg IM q24h for two weeks is also used for pneumocytosis treatment, as well as atovaquone at 15 mg/kg PO q24h for three weeks, and a combination of clindamycin and primaquine (3–13 mg/kg PO q8h for three weeks). Supportive therapy to ease the respiratory distress is necessary in acute pneumocystosis.

­Selected Reading and References 1 Lappin MR. Update on the diagnosis and management

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of Toxoplasma gondii infection in cats. Top Compan Anim Med 2010; 25: 136–41. Baneth G, Thamsborg SM, Otranto D, et al. Major parasitic zoonoses associated with dogs and cats in Europe. J Comp Pathol 2016; 155(1 Suppl 1): S54–74. Esch KJ, Petersen CA. Transmission and epidemiology of zoonotic protozoal diseases of companion animals. Clin Microbiol Rev 2013; 26: 58–85. Hartmann K, Addie D, Belák S, et al. Toxoplasma gondii infection in cats: ABCD guidelines on prevention and management. J Feline Med Surg 2013; 15: 631–7. Dubey JP, Schares G. Neosporosis in animals – the last five years. Vet Parasitol 2011; 180: 90–108. Lyon C. Update on the diagnosis and management of Neospora caninum infections in dogs. Top Compan Anim Med 2010; 25: 170–5. Ishigaki K, Noya M, Kagawa Y, Ike K, Orima H, Imai S. Detection of Neospora caninum‐specific DNA from cerebrospinal fluid by polymerase chain reaction in a dog with confirmed neosporosis. J Vet Med Sci 2012; 74: 1051–5. Madeira MF, Almeida AB, Barros JH, et al. Trypanosoma caninum, a new parasite described in dogs in Brazil: aspects of natural infection. J Parasitol 2014; 100: 231–4.

9 Watier‐Grillot S, Herder S, Marié JL, Cuny G,

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Davoust B. Chemoprophylaxis and treatment of African canine trypanosomosis in French military working dogs: a retrospective study. Vet Parasitol 2013; 194: 1–8. Desquesnes M, Dargantes A, Lai DH, Lun ZR, Holzmuller P, Jittapalapong S. Trypanosoma evansi and surra: a review and perspectives on transmission, epidemiology and control, impact, and zoonotic aspects. Biomed Res Int 2013; 2013: 321237. Rjeibi MR, Ben Hamida T, Dalgatova Z, et al. First report of surra (Trypanosoma evansi infection) in a Tunisian dog. Parasite 2015; 22: 3. Solano‐Gallego L, Koutinas A, Miró G, et al. Directions for the diagnosis, clinical staging, treatment and prevention of canine leishmaniosis. Vet Parasitol 2009; 165: 1–18. Pennisi MG, Cardoso L, Baneth G, et al. LeishVet update and recommendations on feline leishmaniosis. Parasit Vectors 2015; 8: 302. Solano‐Gallego L, Miró G, Koutinas A, et al. LeishVet guidelines for the practical management of canine leishmaniosis. Parasit Vectors 2011; 4: 86.

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15 Otranto D, Dantas‐Torres F. The prevention of canine

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leishmaniasis and its impact on public health. Trends Parasitol 2013; 29: 339–45. Baneth G. Perspectives on canine and feline hepatozoonosis. Vet Parasitol 2011; 181: 3–11. Allen KE, Johnson EM, Little SE. Hepatozoon spp. infections in the United States. Vet Clin North Am Small Anim Pract 2011; 41: 1221–38. Baneth G, Sheiner A, Eyal O, et al. Redescription of Hepatozoon felis (Apicomplexa: Hepatozoidae) based on phylogenetic analysis, tissue and blood form morphology, and possible transplacental transmission. Parasite Vectors 2013; 6: 102. Johnson EM, Panciera RJ, Allen KE, et al. Alternate pathway of infection with Hepatozoon americanum and the epidemiologic importance of predation. J Vet Intern Med 2009; 23: 1315–18. De Tommasi AS, Giannelli A, de Caprariis D, et al. Failure of imidocarb dipropionate and toltrazuril/ emodepside plus clindamycin in treating Hepatozoon canis infection. Vet Parasitol 2014; 200: 242–5. Cohn LA, Birkenheuer AJ. Cytauxzoonosis, In: Greene CE, ed. Infectious Diseases of the Dog and Cat, 4th edn. St Louis, MO: Elsevier, 2012, pp. 764–71. Carli E, Trotta M, Chinelli R, et al. Cytauxzoon sp. infection in the first endemic focus described in domestic cats in Europe. Vet Parasitol 2012; 183: 343–52. Sherrill MK, Cohn LA. Cytauxzoonosis: diagnosis and treatment of an emerging disease. J Feline Med Surg 2015; 17: 940–8. Solano‐Gallego L, Baneth G. Babesiosis in dogs and cats – expanding parasitological and clinical spectra. Vet Parasitol 2011; 181: 48–60.

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cats: ABCD guidelines on prevention and management. J Feline Med Surg 2013; 15: 643–6. Irwin PJ. Canine babesiosis. Vet Clin North Am Small Anim Pract 2010; 40: 1141–56. Ayoob AL, Hackner SG, Prittie J. Clinical management of canine babesiosis. J Vet Emerg Crit Care 2010; 20: 77–89. Gruffydd‐Jones T, Addie D, Belák S, et al. Giardiasis in cats: ABCD guidelines on prevention and management. J Feline Med Surg 2013; 15: 650–2. Kent M, Platt SR, Rech RR, et al. Multisystemic infection with an Acanthamoeba sp in a dog. J Am Vet Med Assoc 2011; 238: 1476–81. Carlesso AM, Mentz MB, da Machado ML, et al. Characterization of isolates of Acanthamoeba from the nasal mucosa and cutaneous lesions of dogs. Curr Microbiol 2014; 68: 702–7. Gruffydd‐Jones T, Addie D, Belák S, et al. Tritrichomoniasis in cats: ABCD guidelines on prevention and management. J Feline Med Surg 2013; 15: 647–9. Yao C, Köster LS. Tritrichomonas foetus infection, a cause of chronic diarrhea in the domestic cat. Vet Res 2015; 46: 35. Tolbert MK, Stauffer SH, Gookin JL. Feline Tritrichomonas foetus adhere to intestinal epithelium by receptor‐ligand‐dependent mechanisms. Vet Parasitol 2013; 192: 75–82. Kanemoto H, Morikawa R, Chambers JK, et al. Common variable immune deficiency in a Pomeranian with Pneumocystis carinii pneumonia. J Vet Med Sci 2015; 77: 715–19.

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111 Coccidia Chris Adolph, DVM, MS, Diplomate ACVM (Parasitology) Zoetis Inc., Tulsa, OK, USA

­Cystoisospora spp. Etiology/Pathophysiology Within the last few years, canine and feline enteric coccidia belonging to the genus Isospora were reorganized and are now recognized by the genus Cystoisospora. This designation is now well accepted but not well integrated. The reader should recognize that both terms refer to the same genus. Coccidia are obligate intracellular parasites classified in the phylum Apicomplexa, and are commonly identified infecting the gastrointestinal tract of dogs and cats. Dogs are the definitive host to four species of Cystoisospora: C. canis, C. ohioensis, C. burrowsi, and C. neorivolta. The latter three species are generally grouped together as C. ohioensis complex. Cats are the definitive host for two species of Cystoisospora: C. felis and C. rivolta. Cystoisospora spp. are very species specific; dogs are not infected with cat Cystoisospora spp. and cats are not infected with dog Cystoisospora spp. To truly understand the pathophysiology of these organisms, one must first understand the life cycle. The typical Apicomplexa life cycle consists of sexual and asexual reproductive events that occur inside and outside the host. A sporozoite is the infective stage and is the end‐ product of sporogony, a form of asexual reproduction. For Cystoisospora spp., this occurs in the environment outside the host. Sporozoites are contained within a sporulated oocysts. If this environmental sporulated oocyst is ingested by the target host, the sporozoites emerge from the oocyst, or excyst, in the small intestine. It should be noted that Cystoisospora spp. have the ability to utilize paratenic hosts as a means of completing the life cycle. If a paratenic host ingests the sporulated oocysts, the sporozoite will invade the extraintestinal soft tissues and encyst. This stage is infective to the final host if the paratenic host

is ingested. In the small intestine, the sporozoite excysts from the sporocyst and invades small intestinal cells. The site, location, and number of asexual generation vary with each species of Cystoisospora. Once the sporozoite invades the cell wall, it essentially hijacks the host cell and can evade the host  immune response. After another round of asexual reproduction by endogeny to produce first‐generation schizonts (aka meronts) containing merozoites, the host intestinal cell is ruptured, releasing the merozoites to invade other cells. This process continues for a programmed number of cycles until the last cycle which is the sexual phase known as gametogony. Here, the merozoites differentiate into either a microgametocyte (male) or a macrogametocyte (female). The microgametocyte leaves the cell in search of a macrogametocyte to fertilize. The end‐product of the sexual stage is a gametozoite known as an oocyst. Pathologic changes to the host occur when developing stages (merozoites and gametozoites) emerge and rupture the epithelial cells. This leads to gastrointestinal bacteria colonizing the ruptured enterocyte. Epidemiology Transmission of Cystoisospora spp. is fecal–oral. Oocysts that are shed are not immediately infective to the next host. The oocyst needs time to sporulate in the environment (eight hours) to be infectious. Cystoisospora spp. also infect the definitive host via predation of a ­paratenic host. When this route is utilized, a paratenic host c­onsumes sporulated oocysts, but asexual and s­ exual development do not occur. Sporozoites then enter extraintestinal tissue and form monozoic cysts, commonly found in mesenteric lymph nodes. Once a definitive host consumes the paratenic host, the life cycle continues as described previously. Infections are generally self‐limiting.

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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The distribution of Cystoisospora is worldwide. Due to the fecal–oral route of transmission, cases are associated with crowded and unhygienic environments. In the United States, prevalence of canine Cystoisospora spp. has been estimated to be between 3.1% and 4.4% while feline Cystoisospora spp. prevalence has been estimated to be between 3.9% and 6.3%. Signalment In most surveys, Cystoisospora spp. are detected in puppies and kittens less than 6 months of age. There does not appear to be a breed predilection. Patients that are most susceptible to clinical disease include nursing, recently weaned, and immunocompromised dogs and cats. History and Clinical Signs Clinical symptoms are more common in puppies and kittens acquired from unsanitary or overcrowded ­ ­environments. Cystoisospora spp. can cause large or small bowel diarrhea, although patients may be asymptomatic despite shedding oocysts. Other clinical symptoms reported include anemia, dehydration, vomiting, anorexia, and weight loss. Physical exam findings of patients with Cystoisospora spp. are nonspecific and can include abdominal pain, increased gas or fluid in the ­gastrointestinal tract, and thickened loops of bowel.

and examined for the presence of characteristic oocysts (Figure 111.1). It is important to differentiate oocysts of Cystoisospora spp. from those of Eimeria spp., which are not pathologic to canine and feline patients. Coprophagy of the feces of animals such as cattle, sheep, and chickens may lead to spurious detection of Eimeria spp. in canine and feline fecal samples. Both Eimeria and Cystoisospora are ­species specific and therefore pathogenic only to their target species. Therapy Asymptomatic adult patients demonstrating fecal oocysts may be monitored for development of clinical signs. Patients that meet criteria of history, clinical signs, and detection of the organism are treated symptomatically for diarrhea, and concurrently with antiprotozoal medications such as ponazuril, sulfonamides, or amprolium (Tables 111.1 and 111.2). Table 111.1  Enteric coccidia of the dog and cat Diagnostic stage

Size (μM)

Canine   C. canis   C. ohioensis   C. neorivolta   C. burrowsi

Oocyst Oocyst Oocyst Oocyst

30 × 38 19 × 23 11 × 13 17 × 20

History, clinical symptoms, and detection of the organism are needed for a definitive diagnosis of coccidiosis. Fecal flotation, centrifugal or passive, can be performed

Feline   C. felis   C. rivolta

Oocyst Oocyst

30 × 40 20 × 25

(a)

(b)

Diagnosis

Organism

Cystoisospora spp.

Figure 111.1  (a) Sporulated and (b) unsporulated Cystoisospora ohioensis. Source: Courtesy of the National Center for Veterinary Parasitology, Oklahoma State University.

110 Coccidia

Table 111.2  Treatment of enteric coccidiosis in dogs and cats Antiprotozoal agent

Treatment regimen

Ponazurila

20 mg/kg PO twice 1–7 days apart OR 50 mg/kg PO once

Sulfadimethoxineb

50–60 mg/kg PO daily for 5–20 days

Trimethoprim‐ sulfonamidec

15–30 mg/kg PO every 12–24 hours for 5 days

Amproliumd

300–400 mg PO daily for 5 days (canine) 60–100 mg PO daily for 7 days (feline)

Furazolidone

8–20 mg/kg PO every 12–24 hours for 5 days

recover from Cystoisospora spp. infections have species‐ specific immunity, but remain susceptible to infection with other Cystoisospora spp. infections. For example, a puppy that acquires a C. canis infection and recovers remains susceptible to infection with C. ohioensis. While patients with coccidiosis respond to appropriate ­therapy,  untreated infections can lead to death from dehydration. Public Health Implications Due to the species specificity, Cystoisospora spp. of dogs and cats are not zoonotic to humans.

a

 Ponazuril is coccidiocidal and may be superior to other drugs.  Sulfadimethoxine is the only approved drug for treatment of coccidiosis in the United States. c  Trimethoprim‐sulfonamide combinations may cause acute hepatic necrosis, keratoconjunctvitis sicca, macrocytic anemia, and type III hypersensitivity reactions. d  Amprolium can cause anorexia, depression, diarrhea, and CNS disease due to induction of thiamine deficiency. PO, by mouth (per os). b

Symptomatic therapy begins by withholding food i­nitially, followed by small, frequent meals with a bland diet once symptoms diminish. A class of medications commonly utilized to treat coccidiosis are the sulfonamides, which may be potentiated for better efficacy. Sulfadimethoxine is labeled in the US for dogs with bacterial enteritis associated with coccidiosis. Sulfonamides block folic acid synthesis and are effective against the schizont, an asexual stage. Sulfonamides are coccidiostatic, therefore fecal analysis may remain positive for oocysts during the course of therapy. Toxicity with use of sulfonamides has been reported and includes crystalluria, renal toxicity, keratoconjunctivitis sicca, type 3 hypersensitivity disorders, acute hepatic necrosis, and hematopoietic disorders. Dobermans are at increased risk for immune‐mediated complications. Ponazuril, a medication used to treat equine protozoal myeloencephalitis, has recently gained popularity as an off‐label treatment for canine and feline coccidiosis. The mechanism of action is incompletely understood. A key improvement over previous treatments is that ponazuril is coccidiocidal rather than coccidiostatic, and fecal flotation can be negative 1–2 days post treatment. Amprolium, another coccidiostat, has also been used for canine and feline coccidiosis. It is a competitive inhibitor of thiamine and is most effective against the first‐generation schizont. Prognosis Prognosis following proper diagnosis and treatment of canine and feline coccidiosis is good. Patients who

­Cryptosporidium spp. Epidemiology/Pathophysiology There are many species of Cryptosporidium that infect a wide range of hosts, including dogs and cats. This genus is considered by some parasitologists to be more closely related to a group of the Apicomplexa known as the gregarines, which infect mostly invertebrates. Recently, ­gregarine‐like stages have been described in multiple Cryptosporidium spp. This may lead to a reclassification of this genus in the future, and could possibly explain why most of the medications effective against coccidia are ineffective against Cryptosporidium spp. The major host of C. canis is the dog; for C. felis, the  major host is the cat. Unlike Cystoisospora spp., Cryptosporidium spp. are not host specific, although dog and cat Cryptosporidium spp. tend to favor the major host. This is an important distinction as other species, such as C. parvum, can be zoonotic. The life cycle is similar to that of Cystoisospora. The oocyst that is infective to the host contains four sporozoites that are not contained within sporocysts. The sporozoite will penetrate the host epithelial cell where trophozoites undergo schizogony to form merozoites. These merozoites will, in turn, invade other epithelial cells and undergo another round of schizogony. This is followed by the sexual phase, or gametogony, where microgametocytes are produced, fuse, and form zygotes. Zygotes can ultimately form either a thick‐walled oocyst that is infective (sporulated) when it leaves the host, or a thin‐walled oocyst that remains within the host and leads to autoinfection. Pathologic changes occur when epithelial cells are damaged by emerging stages. Malabsorption ensues, and diarrhea is the primary clinical presentation. The severity of the infection is limited by developing immunity. Infection can be self‐limiting if a host has a competent immune system. However, clinical disease may be prolonged in dogs and cats with a compromised immune system.

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Epidemiology Transmission of Cryptosporidium spp. is fecal–oral. Oocysts that are shed are immediately infective to the next host. Oocysts are stable in the environment, and  high population densities as well as unsanitary ­conditions favor transmission. Young and immunocompromised animals are considered to be at greater risk. The infective dose is low; very few oocysts are required to produce an infection leading to clinical disease. The distribution of Cryptosporidium spp. in dogs and cats is worldwide. Prevalence of Cryptosporidium spp. in dogs and cats varies, but has been estimated at 5%.

Intravenous fluid administration may be needed to treat severe dehydration. Published data regarding the treatment of cryptosporidiosis are lacking. Treatment protocols at this point are anecdotal and treatment needs to be calibrated to the  needs and response of the individual patient. Azithromycin (10 mg/kg PO, SID until resolution), nitazoxanide (25 mg/kg PO, BID for seven days), paromomycin (150 mg/kg PO, SID for at least five days), and tylosin (10–15 mg/kg PO, BID‐TID for 14–21 days) have been used. Paromomycin has the potential for nephrotoxicity and should not be used in dehydrated patients.

Signalment

Prognosis

Most cases of cryptosporidiosis have been reported in dogs and cats with a compromised immune system, those co‐infected with other pathogens, or with preexisting conditions of the gastrointestinal tract (inflammatory bowel disease [IBD], lymphoma, feline leukemia virus [FeLV], canine distemper). Gastrointestinal symptoms may be more common in cats than in dogs.

In an immunocompetent animal, infection with Crypto­ sporidium spp. usually resolves. Conversely, treatment of immunocompromised animals can be very difficult, and extended supportive measures may be necessary.

History and Clinical Signs The majority of canine and feline patients with detectable Cryptosporidium spp. oocysts are asymptomatic. When clinical disease is present, small bowel diarrhea, anorexia, and weight loss are common. Water loss secondary to malabsorption can lead to severe fluid ­ imbalance. Diagnosis A combination of a direct saline smear and fecal flotation is preferred as the initial step, as oocyst excretion coincides with the onset of clinical signs. However, these techniques have poor sensitivity due to the small size of the oocyst. Fecal smears may be stained to enhance detection. Direct fluorescent antibody, enzyme‐linked immunosorbent assay (ELISA), and polymerase chain reaction (PCR) are also available. These detection methods provide greater sensitivity than that observed with other detection methods. Therapy Direct treatment of the organism is difficult and the main goal of therapy is supportive. Initiation of highly digestible diets, fiber, and probiotics may be of benefit.

Public Health Implications Health professionals should be aware that there have been cases of Cryptosporidium spp. transmission from animals to people. In a majority of cases, the pathogen was C. parvum and the source of infection was farm animals. However, C. felis and C. canis have been reported in the feces of some people. With no established treatment protocol for these cases, it is recommended that immunocompromised people avoid contact with animal feces, in particular the feces of strays and young pets ( meropenem) Nephrotoxicity (imipenem)

Fluoroquinolones

Articular cartilage erosions/blisters (growing animals) Retinal toxicity (cats, dose‐dep.)

Macrolides

Theophylline (CYP3A inhibition, erythromycin, clarithromycin)

Lincosamides

Esophageal structures (cats, clindamycin)

Metronidazole

Neurotoxicity/cerebellovestibular ataxia

Potentiated sulfonamides

Tetracyclines

Antacids/sucralfate (decreased oral absorption) Theophylline (CYP1A inhibition)

Idiosyncratic (dogs) Blood dyscrasias ●● Hepatoxicity ●● Polyarthritis ●● Cutaneous skin eruptions ●● KCS Dose dependent (dogs) ●● Nonregenerative anemia ●● Inhibition of thyroid hormone synthesis ●●

Tooth enamel discoloration (young, tetracyclines, oxytetracycline) Esophageal strictures (cats, doxycycline)

Antacids/sucralfate (decreased oral absorption)

ACh, acetylcholine; CNS, central nervous sysem; KCS, keratoconjunctivitis sicca.

The clinical situation where the risk of the systemic side‐effects of nephrotoxicity and ototoxicity may be avoided is the use of orally administered neomycin in the treatment of hepatic encephalopathy. The AGs have poor oral bioavailability. In patients with hepatic encephalopathy, the goal of therapy is to target the ammonia‐ producing bacteria within the gut. However, if the integrity of the gastrointestinal tract is compromised, systemic absorption and the associated side‐effects may theoretically occur. Amphenicols (e.g., Chloramphenicol) Chloramphenicol has a wide spectrum of activity against gram‐positive and gram‐negative organisms. Many anaerobic bacteria are also sensitive to chloramphenicol, including Clostridium, Bacteroides (B. fragilis), and Fusobacterium species. Other susceptible bacteria include Chlamydia spp., Mycoplasma spp., and Rickettsia rickettsii. In dogs and cats, chloramphenicol can result in bone marrow suppression. This side‐effect is considered dose-

dependent and more likely if patients, especially cats, are treated for an extended duration. In people, the major route of chloramphenicol metabolism and elimination is glucuronidation. This suggests that decreased glucuronidation may contribute to the increased risk of toxicity in cats. In addition, 25% of the dose is reported to be excreted unchanged in the urine of cats compared to most other species, in which elimination is primarily via hepatic metabolism. Therefore, dose reduction should be considered in cats with kidney disease. In dogs, chloramphenicol is a potent inhibitor of the hepatic microsomal cytochrome P450 enzyme system, specifically the subfamily CYP2B11. Through this CYP inhibition, the metabolism of barbiturates, like phenobarbital, and propofol may be impacted. Dogs concurrently prescribed chloramphenicol and phenobarbital have an increased risk of phenobarbital toxicity and dogs anesthetized with propofol may have prolonged recoveries if they are also being treated with chloramphenicol. Chloramphenicol is associated with an irreversible aplastic anemia in people that is an idiosyncratic ­toxicity. Due to this unpredictable toxicity in people,

114  Antimicrobial Therapy in Dogs and Cats

the use of chloramphenicol in veterinary medicine is limited primarily to patients with cultured bacterial ­isolates that have maintained their sensitivity to chloramphenicol but are multidrug‐resistant organisms (e.g., methicillin‐resistant Staphylococcus pseudintermedius). When prescribing chloramphenicol, owners need to be educated about the associated risk of exposure and toxicity to themselves and the need to wear gloves ­ ­whenever the drug is being handled to avoid accidental ingestion or exposure. Beta‐Lactams Penicillins

The broader spectrum (e.g., amoxicillin, ampicillin) or potentiated penicillins (e.g., amoxicillin/clavulanate and ampicillin/sulbactam) have an increased spectrum of activity against gram‐negative organisms, including some of the beta‐lactamase‐producing bacteria and anaerobes. Ticarcillin has a strong gram‐negative spectrum with activity against Pseudomonas spp. Methicillin/oxacillin has activity against the beta‐lactamase‐­producing bacteria (e.g., Staphylococcus spp.) and is used primarily for susceptibility testing. Beta‐lactams have relatively limited side‐effects and are clinically well tolerated. The most common side‐ effect is gastrointestinal upset, which is most commonly associated with amoxicillin/clavulanate, especially in cats. Hypersensitivity or allergic reactions to the penicillins are not very common in small animal medicine, in comparison to people. The beta‐lactams are largely excreted in the urine and in patients with kidney disease, dose reduction may be necessary. However, due to the wide margin of safety of these drugs, dose reduction is often not routinely pursued in patients with mild to moderate azotemia. Cephalosporins

The cephalosporins have a broad spectrum of activity against gram‐positive organisms, with the exception of  the Enterococcus spp. which are not susceptible. The  cephalosporins as a group have some anaerobic activity but relative to the penicillins and potentiated penicillins (e.g., amoxicillin/clavulanate), their anaerobic spectrum is limited. Generally, the gram‐negative spectrum of activity increases from the first‐ to third‐ generation cephalosporins. However, some of the less frequently dosed third‐generation (e.g., cefpodoxime) and extended‐generation cephalosporins (e.g., cefovecin) have a bacterial spectrum more consistent with the first‐generation cephalosporins, including activity against many gram‐positive and some gram‐negative organisms but no activity against Enterococcus or Pseudomonas species.

Cefpodoxime is an oral formulation dosed once a day, and cefovecin an injectable slow‐release formulation that delivers antibacterial activity for 14 days. Cefovecin’s extended persistence may provide convenient dosing but in the event of an adverse effect, the drug cannot be withdrawn and in the case of treatment failure, continued drug exposure may contribute to antimicrobial resistance. The clinical use of cefovecin should be limited to fractious or noncompliant patients that cannot be medicated daily. The fourth‐generation cephalosporins have both gram‐positive and gram‐negative activity and their use should be limited to patients with a cultured susceptible gram‐negative or gram‐positive isolate, patients with susceptible mixed infections, or a serious to life‐threatening susceptible multidrug‐resistant and/or nosocomial infection(s). The later generation cephalosporins are expensive, require frequent dosing, and in some cases, a constant rate infusion is necessary for effective therapy. Carbapenems Carbapenems are considered powerful antibiotics with a very broad spectrum of activity. Therefore, their use should be dictated by the results of bacterial culture and antimicrobial susceptibility testing. The carbapenems are expensive and must be administered parenterally. Side‐effects, more common with imipenem than meropenem, include central nervous system (CNS) toxicity and nephrotoxicity. Imipenem is available co‐formulated with cilastatin (an inhibitor of imipenem metabolism) which minimizes nephrotoxicity. In the absence of cilastatin, imipenem is converted to a nephrotoxic ­ metabolite in the proximal renal tubule, resulting in necrosis. In many patients, the use of meropenem may be better tolerated as it is more soluble and can delivered in smaller volumes relative to imipenem. Fluoroquinolones The spectrum of activity of fluoroquinolones classically targets aerobic gram‐negative organisms. Pradofloxacin is a third‐generation fluoroquinolone that has recently been approved in the US for the treatment of skin infections in cats. Pradofloxacin has an extended spectrum against many gram‐positive organisms and anaerobes in addition to having a good gram‐negative spectrum. Fluoroquinolones are associated with significant side‐effects and clinically important drug interactions. Articular cartilage erosions or blister lesions occur in growing animals, especially dogs treated with high doses. Clinical signs of lameness and joint swelling have been reported as early as two days following therapy in

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experimental dogs given four times the recommended dose of ofloxacin. The use of fluoroquinolones is generally avoided in young puppies ( monohydrate) can cause local esophagitis and ulceration, leading to stricture formation. A similar mechanism of stricture formation has been proposed in cats. Administering a liquid formulation versus capsules of doxycycline to cats is recommended to avoid stricture formation. Clinically important drug interactions are also associated with orally administered tetracyclines. Tetracyclines administered in association with drugs or vitamins containing di‐ or tri-valent cations (e.g., calcium, magnesium, iron) result in dramatically lower oral absorption of tetracycline antimicrobials. Common drugs that contain aluminum and/or magnesium should not be co‐ administered with any of the tetracycline a­ntibiotics,

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including bismuth subsalicylate (Pepto‐Bismol®), sucralfate, and many antacids. This interaction may be less clinically significant for doxycycline due to its lower affinity for calcium relative to other tetracyclines. Additional Considerations To optimize antimicrobial therapy, clinicians should have a working knowledge of the available drugs, including their spectrum of activity, clinically important side‐ effects, and drug–drug interactions. Prior to writing any antibiotic prescription, consideration of some important questions can help optimize clinical outcomes. Define the need for antimicrobial therapy. Is a bacterial infection suspected or identified? Define the spectrum of activity needed. Where is the site of infection? What is the most likely infecting bacterial pathogen(s)? Are additional diagnostics necessary such as culture and ­ antimicrobial susceptibility testing? Define the patient characteristics which may limit efficacy or increase the risk for an adverse event or drug–drug interaction. Does my patient have underlying kidney or liver disease? What other drugs are being administered to this patient?

Often, in otherwise healthy patients with uncomplicated bacterial infections in which the infecting pathogen(s) are predictable (e.g., canine female lower urinary tract infections), empiric antimicrobial therapy is effective. However, bacterial culture and antimicrobial susceptibility become mandatory for any patient following their first empiric antibiotic therapeutic failure. Other clinical situations in which antimicrobial therapy becomes more complicated include patients with recurrent infections, multidrug‐resistant infections, or in immunocompromised patients in which bacterial culture and antimicrobial susceptibility testing are critical to optimize efficacy and minimize further antimicrobial resistance. In cases of chronic recurrent or multidrug‐resistant infection, repeating bacterial culture and antimicrobial susceptibility testing during therapy and/or following therapy becomes essential to confirm responsiveness to antimicrobial therapy and/or confirm the resolution of infection. Consensus guidelines for treatment of bacterial infections in dogs and cats are available at the International Society for Companion Animal Infectious Disease website (www. iscaid.org)

­Further Reading Hillier A, Lloyd DH, Weese JS, et al. Guidelines for the diagnosis and antimicrobial therapy of canine superficial bacterial folliculitis (Antimicrobial Guidelines Working Group of the International Society for Companion Animal Infectious Diseases). Vet Derm 2014;25:163–e143. Lappin MR, Blondeau J, Boothe D, et al. Antimicrobial use Guidelines for Treatment of Respiratory Tract Disease in Dogs and Cats: Antimicrobial Guidelines Working Group of the International Society for Companion Animal Infectious Diseases. J Vet Intern Med 2017;31:279–294. Lees P. Pharmacokinetics, pharmacodynamics and therapeutics of pradofloxacin in the dog and cat. J Vet Pharmacol Ther 2013; 36: 209–21. Ramirez CJ, Minch JD, Gay JM, et al. Molecular genetic basis for fluoroquinolone‐induced retinal degeneration in cats. Pharmacogenet Genomics 2010; 21: 66–75.

Trepanier LA. Idiosyncratic toxicity associated with potentiated sulfonamides in the dog. J Vet Pharmacol Ther 2004; 27: 129–38. Trepanier LA. Cytochrome P450 and its role in veterinary drug interactions. Vet Clin North Am Small Anim Pract 2006; 36: 975–85, v. Weese JS, Blondeau JM, Boothe D, et al. Antimicrobial use guidelines for treatment of urinary tract disease in dogs and cats: antimicrobial guidelines working group of the international society for companion animal infectious diseases. Vet Med Int 2011; 2011: 263768. Weese JS, Blondeau J, Boothe D, et al. International Society for Companion Animal Infectious Diseases (ISCAID) guidelines for the diagnosis and management of bacterial urinary tract infections in dogs and cats. The Veterinary Journal 2019;247:8–25.

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115 Antifungal Therapy Daniel S. Foy, MS, DVM, DACVIM (SAIM), DACVECC College of Veterinary Medicine, Midwestern University, Glendale, AZ, USA

Until relatively recently, treatment of systemic mycoses was limited to intravenous amphotericin B and oral keto­ conazole. However, in the last two decades, significant progress has been made in the development of first‐­ generation triazole drugs, newer second‐generation ­triazole drugs, and echinocandins. New antifungal therapies are often constrained by the fact that fungal organisms are eukaryotic, and therefore greater potential exists for host toxicity. For example, traditional antifungal medications that target ergosterol, or its production, can cause toxicity in mammalian cells via inhibition of cholesterol production or damage to cell membranes. Newer therapies are being directed at com­ ponents unique to fungal organisms, thereby sparing mammalian cells. Although in vitro testing frequently finds antifungal drugs to be fungicidal, most appear to be fungistatic in vivo, thus the host immune system must eliminate the organisms.

­Amphotericin B Since its discovery in 1956 and increased availability in the early 1960s, amphotericin B has become, and remains, the reference treatment for invasive fungal infections. Amphotericin B is a macrocyclic polyene antibiotic origi­ nally extracted from Streptomyces nodosus. This drug forms micelles with fungal ergosterol, which creates channels in the fungal membrane, alters cell permeability, and allows leakage of ions and cellular components from the fungal organism. The effectiveness of amphotericin B is due to its greater affinity for ergosterol, the major sterol of fungal cell membranes, relative to cholesterol. Effective amphotericin B treatment requires intrave­ nous (IV) administration, with the major limiting factor in its use being cumulative nephrotoxicity; however, the only absolute contraindication to use is anaphylaxis.

The cause of nephrotoxicity remains incompletely understood, but is potentially related to both direct tox­ icity to epithelial cell membranes and renal vasoconstric­ tion. In people, serum creatinine is monitored during treatment; a clinically significant increase is considered a new elevation above the normal range, or an increase of greater than 20% from the baseline value. The inherent nephrotoxicity of the original ampho­ tericin B formulation led to the development of three new formulations: liposomal preparation, lipid complex, and colloidal dispersion with cholesterol sulfate. Liposomal amphotericin B (lip‐amB) achieves higher plasma concentrations than the original formulation; this is thought to be due to decreased uptake by the reticu­ loendothelial system (RES). Liposomes containing amphotericin B fuse with the fungal cell membrane, lead­ ing to fungal cell death. Lip‐amB has been evaluated for safety in healthy beagle dogs, and when dosed at 1 mg/kg/ day for 29 days, no azotemia was noted, and minimal renal tubular necrosis was seen on histopathology. Amphotericin B lipid complex (ABLC) is composed of a suspension of amphotericin B complexed with two phos­ pholipids: dimyristoylphosphatidylcholine and dimyris­ toylphosphatidylglycerol. ABLC is taken up by the cells of the RES, and subsequently concentrates in the liver, lungs, and spleen. The lipid complexes are likely disrupted by phospholipases at sites of inflammation or infection, lead­ ing to the release of amphotericin B. Repeated dosing of up to 5 mg/kg/day in research dogs found ABLC to be 8–10‐fold less nephrotoxic, on the basis of renal values and histology, than conventional amphotericin B. Importantly, although mean glomerular filtration rate (GFR) decreased over the course of treatment, only one of 10 dogs that received 8–12  mg/kg ABLC showed a decrease in GFR below the reference range. The third lipid‐incorporated preparation to be devel­ oped was amphotericin B colloidal dispersion (ABCD).

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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This formulation is composed of amphotericin B inserted between cholesterol sulfate bilayers, creating a disc‐like structure. Similar to ABLC, the RES rapidly takes up ABCD. In one canine study, ABCD led to high concen­ trations in the bone marrow, liver, and spleen in healthy dogs, while concentrations remained low in the kidneys and lungs. All three modified amphotericin B preparations have greater hydrophobicity, which likely results in greater delivery to the site of infection, and decreased delivery to the kidneys. All three modifications appear to maintain efficacy relative to conventional amphotericin B, while decreasing nephrotoxicity in humans. Amphotericin B remains an important treatment option in veterinary medicine, as many of the newer drugs available for peo­ ple are cost‐prohibitive in veterinary patients.

­Azoles The azole drugs exert their effect by inhibition of lanos­ terol 14‐alpha demethylase, leading to ergosterol deple­ tion and accumulation of aberrant and potentially toxic sterols in the cell membrane. Azoles are classified as imidazoles or triazoles depending on whether they ­

­ ossess two or three nitrogen molecules within their azole p ring. Individual azole drugs are summarized in Table 115.1. Ketoconazole Ketoconazole was the first azole released, and is the only imidazole antifungal agent remaining in use for the treat­ ment of systemic mycoses. It is highly lipophilic, and is ~99% plasma protein bound in humans, which impairs distribution to the brain and cerebrospinal fluid. Ketoconazole shows optimal dissolution and absorption at an acidic gastric pH in both humans and dogs, and should not be given with antacids. In dogs, ketoconazole has been used historically to treat various systemic mycoses, including blastomycosis and histoplasmosis. Although ketoconazole has been successful in treating systemic mycoses, it should be combined with amphotericin B to yield response rates that are comparable to itraconazole alone in dogs with systemic blastomycosis. When dosed at 5–10 mg/kg/day for three weeks, ketoconazole remains an inexpensive and effective treatment for Malassezia dermatitis in ­veterinary patients. Gastrointestinal upset or decreased appetite account for over half of the adverse events reported in dogs

Table 115.1  Summary of azole antifungal drugs, including formulation, indications, and side‐effects Drug (trade name)

Class and formulation

Indications

Side‐effects

Ketoconazole (Nizoral®)

Imidazole (oral, topical)

Topical mycotic infections Malassezia dermatitis Third‐line treatment for systemic mycoses

GI upset Dose‐dependent increases in ALT Potent CYP3A inhibitor Inhibitor of p‐glycoprotein Absorption impaired by antacids

Fluconazole (Diflucan®)

First‐generation triazole (oral, injectable)

Candidiasis and cryptococcosis Systemic mycoses with ocular or CNS involvement Possibly first‐line treatment of blastomycosis

GI upset Dose‐dependent increases in ALT Requires dosage reduction in renal failure

Itraconazole (Sporanox®)

First‐generation triazole (oral)

First‐line for nonlife‐threatening systemic mycoses that do not involve CNS

GI upset Dose‐dependent increases in ALT CYP3A inhibitor Absorption impaired by antacids

Voriconazole (Vfend®)

Second‐generation triazole (oral, injectable)

Invasive aspergillosis Likely efficacious against most systemic mycoses

Visual and neurologic abnormalities CYP3A inhibitor Induces its own metabolism over time in dogs

Posaconazole (Noxafil®)

Second‐generation triazole (oral)

Aspergillosis, candidiasis, and cryptococcosis Limited data but likely effective against other systemic mycoses

GI upset Headache Prolongation of QT interval CYP3A inhibitor

Clotrimazole

Imidazole (topical)

Sinonasal aspergillosis Malassezia otitis

Poor oral bioavailability

Enilconazole

Imidazole (topical)

Sinonasal aspergillosis

Poor oral bioavailability

115  Antifungal Therapy

treated with ketoconazole, with 7% of dogs exhibiting nausea or vomiting and 5% showing only inappetence; similar signs are seen in treated cats. Hepatotoxicity and cutaneous reactions are much less frequently observed. Hepatotoxicity from ketoconazole typically manifests as mild to moderate increases in alanine aminotransferase (ALT) that are reversible with drug discontinuation. Ketoconazole is a potent inhibitor of both the P450 enzyme CYP3A and p‐glycoprotein, and therefore has many potential drug interactions. For example, ketocon­ azole leads to increased plasma concentrations of iver­ mectin and midazolam in dogs, and cyclosporine in dogs and cats. It inhibits the adrenal production of many adre­ nal steroids such as cortisol and testosterone, and should be avoided in breeding animals. Fluconazole Fluconazole is a first‐generation triazole drug that was initially released in 1990 for the treatment of candidiasis and cryptococcosis. Its in vitro susceptibility profile sug­ gests low potency as an antifungal agent; however, it is relatively water soluble, minimally (~10%) protein bound, and distributes well throughout the body, leading to bet­ ter efficacy in vivo. Fluconazole effectively penetrates the blood–brain barrier, as well as the blood–ocular and blood–prostate barriers, in both people and veterinary patients. Fluconazole absorption is not affected by con­ current use of antacids, and may be a better choice in patients requiring H2 blockers or proton pump inhibitor therapy. Fluconazole also does not require food for opti­ mal absorption. As fluconazole is approximately 70% excreted unchanged in the urine, the dosage should be reduced in patients with compromised renal function. As in humans, fluconazole has been recommended for veterinary patients with systemic mycoses affecting the central nervous system (CNS) or eyes. Fluconazole has been used successfully in dogs and cats with cryptococ­ cosis and blastomycosis, and the introduction of generic fluconazole has significantly reduced its cost. Moderate, reversible ALT increases have been observed in dogs receiving fluconazole for treatment of blastomycosis. Fluconazole is a teratogen in animals, and should be avoided during pregnancy. Fluconazole is empirically prescribed at a dosage of 5–10 mg/kg per day in dogs, while cats may receive a dosage of 50 mg per day. Because of its predictable phar­ macokinetics, fluconazole does not require therapeutic drug monitoring. Itraconazole Itraconazole is another first‐generation triazole that was released in 1992, and rapidly became the oral treatment of choice for both histoplasmosis and blastomycosis in

humans. Today, itraconazole remains the preferred azole in people for nonlife‐threatening systemic mycoses that do not involve the CNS. It is lipophilic and is highly pro­ tein bound (>99%). Like ketoconazole, the absorption of itraconazole is diminished in the presence of antacids so it is recommended that itraconazole be administered with food, and that antacids are avoided during itracona­ zole therapy. Itraconazole has become the drug of choice for ­treatment of systemic mycoses in dogs and cats, and is effective for the treatment of blastomycosis, histo­ plasmosis, cryptococcosis, and coccidioidomycosis. Although itraconazole does not effectively penetrate the blood–brain or blood–ocular barriers, it may achieve adequate levels to treat CNS or ocular infection when there is associated inflammation and compro­ mise of the barriers. Itraconazole has been dosed at 5–10 mg/kg per day, and may be administered as a single dose or divided into twice‐daily dosing. The most common side‐effects are  gastrointestinal upset and hepatocellular toxicity. Although a loading dose of itraconazole has been sug­ gested when starting therapy, no difference has been found with or without an initial loading dose. Substitution with generic itraconazole has been shown to reduce plasma itraconazole concentrations to the sub­ therapeutic range in people, and has led to recrudes­ cence of infection. A recent study in dogs found that neither generic nor compounded itraconazole achieved bioequivalence with branded itraconazole. As with ketoconazole, itraconazole is an inhibitor of CYP3A and has many potential drug interactions. It has been shown to increase the concentrations of cyclo­ sporine, digoxin, and midazolam in people. Unlike keto­ conazole, itraconazole does not appear to be a significant inhibitor of cortisol or testosterone synthesis at clinically relevant doses. Itraconazole is available as an oral cap­ sule and solution. Newer Triazoles Voriconazole is a second‐generation triazole that is structurally similar to fluconazole. It has become the drug of choice for treatment of invasive aspergillosis in humans. Despite its similarity to fluconazole, voricona­ zole is poorly water soluble and moderately protein bound. Hepatotoxicity, skin rash or eruptions, and peripheral neuropathy have been reported. Like ketoconazole and itraconazole, voriconazole is a substrate inhibitor of CYP3A, and can increase plasma concentrations of other drugs in humans. It is also teratogenic in animals and should not be used during pregnancy. In dogs, voriconazole undergoes extensive metabolism; it also ­ induces its own metabolism over time. The in vitro

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potency of voriconazole against veterinary fungal iso­ lates appears to be favorable when compared to itracona­ zole. Voriconazole is available as an oral tablet or solution, or as an IV preparation. The use of voricona­ zole has been reported in cats with unilateral orbital aspergillosis; however, it has also been reported to cause neurologic deficits in cats, including ataxia, hindlimb paralysis, and visual abnormalities. Posaconazole is a lipophilic second‐generation triazole derived from itraconazole. It was approved for use in people for prophylaxis of invasive Aspergillus and Candida infections and for treatment of oropharyngeal candidiasis. Posaconazole also appears to have efficacy against Blastomyces, Histoplasma, and Coccidioides spp. As with itraconazole, administration of posaconazole with a meal appears to improve bioavailability; however, like fluconazole, alterations in gastric acidity do not appear to affect posaconazole absorption. The most common adverse effects reported in human patients are headache and gastrointestinal upset; hepa­ totoxicity and QT interval prolongation are reported but much less common. Although posaconazole appears to have a narrower drug interaction profile, inhibition of  CYP3A4 has been demonstrated in humans. Posaconazole is available only in oral formulations. Topical Azoles Both clotrimazole and enilconazole are classified as imi­ dazoles within the azole class of drugs; however, both drugs have minimal systemic bioavailability due to a high first‐pass effect. These drugs are thus confined to topical use, such as clotrimazole in otic suspensions for the treatment of Malassezia otitis in dogs and cats. Both clotrimazole and enilconazole are effective, when instilled into the nasal passages, for treating sinonasal aspergillosis in dogs; clotrimazole has also been reported to be effective in cats with nasal aspergillosis. Although some dogs require multiple treatments, more than 85% of dogs can be cured with up to three treatments.

­Echinocandins Echinocandins are a relatively new class of antifungal medications; the first compound, caspofungin, was approved by the FDA in 2001. Echinocandins inhibit ­glucan synthase and prevent the synthesis of beta‐1,3

glucan, an essential component of the cell wall in certain fungi. In susceptible fungi, the integrity of the cell wall is compromised, leading to cell lysis. The clinical use of the echinocandins is limited to  Candida and Aspergillus spp. in human patients; Cryptococcus neoformans and the zygomycetes are typi­ cally resistant. The mycelial forms of Blastomyces ­dermatitidis and Histoplasma capsulatum appear to be susceptible to the echinocandins, although the yeast forms are not, due to the predominance of alpha‐glucan, which is not a target of echinocandins, in the yeast cell wall. Echinocandins have poor oral bioavailability and are only available in intravenous formulations. The side‐ effects associated with echinocandins are typically mini­ mal, with fever, gastrointestinal signs, phlebitis, and headache being most commonly reported.

­Terbinafine Terbinafine belongs to the allylamine group of antifungal agents, and is most frequently used in people for the treatment of dermatophytoses and toenail onychomyco­ sis. Its antifungal activity is mediated via noncompetitive inhibition of squalene epoxidase, an enzyme involved in fungal ergosterol synthesis, with more than 4000‐fold selectivity for fungal versus mammalian P450 enzymes. Terbinafine is very well absorbed from the gastrointesti­ nal tract and then rapidly diffuses from the bloodstream into the dermis and epidermis. Terbinafine is highly ­lipophilic, which leads to its high concentration in hair follicles, skin, nail plate, and adipose tissue, with levels in the stratum corneum exceeding those in plasma by a fac­ tor of 75 within 12 days of therapy. Terbinafine has shown high in vitro efficacy against many dermatophytes, including Trichophyton and Tinea spp. It has also been combined with echinocandins or triazoles in a multimodal approach to systemic mycoses in people. Side‐effects are generally limited to gastroin­ testinal upset and, rarely, hepatotoxicity. Terbinafine is currently available in tablet form as well as a topical cream or gel. In dogs, terbinafine has in vitro activity against Microsporum and Trichophyton isolates, with little e vidence of acquired resistance during treatment. ­ Terbinafine appears to be equivalent or superior to keto­ conazole for the treatment of Malassezia dermatitis in dogs, with a reduction in both yeast counts and pruritus.

­Further Reading Foy DS, Trepanier LA. Antifungal treatment of small animal veterinary patients. Vet Clin North Am Small Anim Pract 2010; 40(6): 1171–88.

Greene CE. Antifungal chemotherapy. In: Infectious Diseases of the Dog and Cat, 4th edn. St Louis, MO: Elsevier, 2012, pp. 579–88.

115  Antifungal Therapy

Mawby DI, Whittemore JC, Genger S, Papich MG. Bioequivalence of orally administered generic, compounded, and innovator‐formulated itraconazole in healthy dogs. J Vet Intern Med 2014; 28(1): 72–7. Mazepa AS, Trepanier LA, Foy DS. Retrospective comparison of the efficacy of fluconazole or itraconazole

for the treatment of systemic blastomycosis in dogs. J Vet Intern Med 2011; 25(3): 440–5. Quimby JM, Hoffman SB, Duke J, Lappin MR. Adverse neurologic events associated with voriconazole use in 3 cats. J Vet Intern Med 2010; 24(3): 647–9.

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Special Topics

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116 Nosocomial and Multidrug‐Resistant Infections Jason W. Stull, VMD, MPVM, PhD, DACVPM1 and J. Scott Weese, DVM, DVSc, DACVIM (SAIM)2 1

 Department of Veterinary Preventive Medicine, Ohio State University, Columbus, OH, USA Ontario Veterinary College, University of Guelph, Guelph, ON, Canada

2 

Nosocomial infections, typically referred to as “hospital‐ associated (or healthcare-associated) infections” (HAIs), are infections acquired by patients during hospitalization and are an inherent risk in veterinary medicine. In human hospitals, HAIs are a well‐recognized contributor to illness and death, with substantial associated financial costs. Although veterinary data for this field are limited, similar (or even higher) HAI rates have been reported compared to human studies. Between 2003 and 2008, 82% of veterinary teaching hospitals in North America and Europe reported at least one HAI outbreak. Many of these ­ outbreaks required restricted patient admissions or ­hospital closure. Animals suffering from HAIs may have an increased hospital stay and suffer permanent health consequences (including death). Some veterinary HA pathogens can be transmitted to staff or pet owners, resulting in human illness. A wide range of pathogens may be involved in HAIs, but most of the focus is on the emerging epidemic of multidrug‐resistant bacteria because of dramatic increases in infections, limited antimicrobial options, and potential public health consequences. Perhaps most important to this topic is the assumption in human medicine that 10–70% of all HAIs are preventable. The proportion of HAIs that are preventable in ­veterinary medicine is unknown, but likely to be similar. The routine use of simple infection prevention practices can reduce HAIs and related multidrug‐resistant ­organisms (MDROs).

­Identification of Hospital‐Associated Infections Early identification of HAIs is critical for effective ­disease control and prevention. It is not unusual for veterinary HAI outbreaks to go unnoticed because of a lack

of centralized data reporting or communication. Key elements of early HAI identification include a hospital‐­ specific surveillance program and routine use of diagnostic culture and susceptibility data to establish practice‐­specific levels of pathogen prevalence and antimicrobial resistance. Surveillance programs allow for early recognition of HAI transmission and outbreaks. They also provide “benchmarking” data that can be used to identify changes in disease rates (e.g., understanding the background surgical site infection rate so that any increases can ­ be  promptly investigated). These programs can also ­provide important information for better understanding and  responding to other disease occurrences, such as MDROs. The specifics of a surveillance program will be driven by the needs and disease risks of the practice. There are three main types of surveillance: active, passive, and syndromic. Active surveillance involves collection of data specifically for infection control purposes. This type of surveillance is expensive and time‐consuming and, given the small population of susceptible patients and unclear response, not typically justifiable in a veterinary hospital. As the prevalence of MDROs increases in the animal population and more is known about HAIs, active surveillance practices may become useful in some situations. Passive surveillance involves the use of data that are already available. For example, it may be used to monitor surgical site infections by reviewing case records for surgical patients after a procedure or collating susceptibility data from urinary E. coli to assess empiric antimicrobial choices. Syndromic surveillance involves detection of identifiable syndromes (e.g., ­coughing, diarrhea). It is an easy tool for identification of certain high‐risk hospitalized cases and can also identify higher‐risk patients prior to admission, so that specific infection control practices can be employed.

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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­ ommon Presentations and Risk C Factors for HAIs In veterinary medicine, urinary tract infections (UTIs), pneumonia, surgical site infections (SSIs), bloodstream infections (BSIs), and gastrointestinal disease (infectious diarrhea) are likely to be the most common presentations for HAIs. Urinary Tract Infections Catheter‐associated UTIs are one of the more common HAIs in small animal veterinary medicine. Pathogens involved may be either endogenous to the patient, arising from the rectum or perineum, or from the hospital environment or people through contamination of the drainage system or bag. Biofilms (a complex structure of  microorganisms and extracellular matrix) can be ­produced by bacteria on the surface of urinary catheters, resulting in poor antimicrobial penetration, ­antimicrobial resistance, and treatment failure. Catheter‐­ associated UTIs are best prevented through careful attention to aseptic technique during placement and maintenance of urinary catheters as well as by avoidance of urinary catheterization unless necessary and at least daily reassessment of whether a urinary catheter is still required. Culture of catheters or urine from catheterized patients in the absence of clinical or cytologic evidence of disease is not recommended. Pneumonia Hospital‐associated pneumonia has been minimally investigated in the veterinary field, in large part because of the limited use of mechanical ventilation (a major contributor in human medicine). Recumbent position, mechanical ventilation, use of endotracheal/nasogastric tubes, and factors that increase aspiration pneumonia (e.g., laryngeal or esophageal disorders and decreased mentation or recumbency) likely increase HAI risk. Recumbent, sedated, or debilitated patients may benefit from having the cranial portion of the body elevated.

endogenous flora of the patient. Therefore, the pathogens responsible for SSIs will vary by type of surgery (e.g., gram‐negative rods occurring in abdominal surgeries, staphylococci in orthopedic procedures). Methods such as clipping the patient immediately before surgery after induction, following strict aseptic technique, ­cleaning and care of clipper blades after each use, minimizing personnel flow in the operating room, and ­prudent use of antimicrobials are important in reducing the likelihood of SSIs. Bloodstream Infections In human medicine, most BSIs are associated with intravascular devices, with duration of catheterization being the most important risk factor. Despite this risk, studies have not found a decrease in the incidence of catheter‐ related (CR) BSIs with prophylactic catheter changes (e.g., every three days). The current recommendation is for catheters to be removed as soon as medically indicated, but for routine changes to be avoided, since a catheter that has remained free of complications for a few days may pose a lower risk than a newly placed c­ atheter. A similar approach is appropriate in veterinary medicine. Veterinary studies have revealed that jugular and intravenous catheters are frequently contaminated with enteric or environmental pathogens. Factors associated with intravenous catheter contamination in dogs and cats include longer duration of catheter placement and patient immunosuppression. Contamination may occur from the hands of people placing or handling the catheter, the patient’s own flora, or the hospital environment. The use of aseptic technique, including appropriate skin preparation, hand hygiene immediately before placement, use of gloves, and minimal contact with the catheter site, is important to reduce CR HAIs. Culture of catheter insertion sites is not recommended since skin bacteria are expected to be present. As HAI outbreaks have been associated with contaminated materials used in skin preparation, all containers holding disinfectant and materials used for skin preparation should be routinely (i.e., daily to weekly) disinfected, sterilized or ­discarded, rather than continuously refilled.

Surgical Site Infections Surgical site infections are an inherent risk of breaching the body’s normal barriers and are estimated to occur in approximately 2–7% of veterinary surgical patients. Factors likely to increase SSI risk include longer duration of surgery or anesthesia, inappropriate antimicrobial therapy, greater contamination of the surgical site, endocrine disease, increased number of people in the operating room, and orthopedic surgeries and procedures with an implant. For most SSIs, the source of pathogens is the

Infectious Diarrhea In small animal veterinary facilities, gastrointestinal HAIs may involve a number of pathogens, including Salmonella spp. Given the high frequency with which animals may subclinically shed gastrointestinal pathogens, efforts must be aimed at both early identification of animals at increased risk for shedding pathogens linked with HAIs (e.g., recent consumption of raw egg or meat products) and routine use of infection control practices.

116  Nosocomial and Multidrug‐Resistant Infections

­Pathogens of Concern In veterinary medicine, current knowledge of many aspects of the epidemiology of important MDROs and HA pathogens is unclear. Based on reported veterinary HA outbreaks or supposition from the human literature, several important pathogens responsible for HAIs (including MDROs) are identifiable (see later). Pathogens involved in HAIs are often opportunistic and found in healthy animals. The frequency of colonization with each pathogen likely varies with geography, animal species, and veterinary practice (in part influenced ­ by  antimicrobial use/pressure). Environmentally stable pathogens have a demonstrated clear “advantage,” increasing the chance of transmission. Given the close interaction between veterinary staff and patients, as well as the often poor hand hygiene practices documented in veterinary practices, human commensals with zoonotic potential are important in veterinary HAIs. Finally, increased resistance to antimicrobials is a common ­feature. The specific resistance profiles and treatment options for common MDR pathogens have recently been summarized. Staphylococcus spp. Staphylococcus pseudintermedius and, to a lesser extent, S. aureus are frequently the cause of veterinary HAIs. As both are often carried in the nasopharynx, on the skin and in the gastrointestinal tracts of dogs and people (respectively), this is expected given the roles of endogenous bacteria and hands of healthcare workers in HAIs. The emergence of methicillin resistance in these species (MRSP and MRSA) has had profound implications for HAI prevention and control. Methicillin resistance is mediated by the mecA gene, which confers resistance to beta‐lactam antimicrobials (penicillins, cephalosporins, and carbapenems); isolates are frequently resistant to additional antimicrobial classes. Methicillin‐resistant Staphylococcus pseudintermedius (MRSP) has rapidly spread globally in canine populations, often with high levels of antimicrobial resistance, something that is of tremendous concern as S. pseudintermedius is the leading opportunistic pathogen in dogs (and to a lesser degree cats). Recent prior hospitalization and beta‐lactam antimicrobial administration have been associated with MRSP infections, suggesting nosocomial transmission may be a factor in MRSP disease. Methicillin‐resistant Staphylococcus aureus (MRSA) is less of an issue from a veterinary HAI standpoint, although its role as an important human pathogen increases concern for zoonotic transmission. Risk factors for MRSA colonization or infection in companion

a­ nimals include antimicrobial use, prior hospitalization, ownership by veterinary or human healthcare workers/ students, and longer hospitalization (>3 days). A high proportion of veterinarians and staff are colonized with MRSA compared to the general public, likely due to ­deficiencies in standard infection control and hygiene practices that allow for zoonotic transmission. As such, they may serve as a source for HAIs in their patients if infection control practices are not observed. Escherichia coli Escherichia coli is a common component of the commensal gastrointestinal microflora and is an important pathogen. Many community and hospitalized companion animals shed MDR E. coli in their feces. Factors associated with dogs shedding or acquiring MDR E. coli during hospitalization include duration of hospitalization (>3 days) and treatment with antimicrobials shortly before or while hospitalized. Of particular concern is the ability for some strains of E. coli to produce beta‐lactamase, notably the extended‐spectrum beta‐lactamase (ESBL) producers, which provide resistance to a broad range of beta‐lactam antimicrobials, including third‐ generation cephalosporins. Resistance to additional ­antimicrobial classes is not uncommon. ESBL‐producing E. coli has been identified as the source of veterinary HAIs, occurring as SSIs and CR UTIs. Carbapenemase‐producing Enterobacteriaceae (or carbapenem‐resistant Enterobacteriaceae; CRE) (including E. coli), conferring resistance to additional antimicrobial classes, have recently become a significant public health concern. CR E. coli have been identified in small animals, with suggested nosocomial transmission. This currently appears to be a rare occurrence, but one that is likely to increase as CREs increase in prevalence in the human population. Enterococcus spp. Enterococci are commonly found in the gastrointestinal tract of animals and humans. E. faecium and E. faecalis are most often involved in HAIs, notably in compromised hosts. Enterococci are inherently resistant to a number of antimicrobial classes, including cephalosporins, some penicillins, fluoroquinolones, clindamycin, and trimethoprim, and may acquire resistance to other antimicrobial classes. Vancomycin‐resistant enterococci (VRE) are an increasing concern in human medicine. To date, VRE appear to be rare in companion animals; ­however, other MDR enterococci are regularly recognized and have been identified in HAIs. Enterococci are often identified as UTIs (including catheter‐associated) although infections at other anatomic sites occur

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(SSIs, BSIs, pneumonia). The high degree of antimicrobial resistance, ability to propagate for extended periods in small animal hosts as a commensal, and environmental persistence make enterococci particularly challenging when involved in HAIs.

Box 116.1  Key methods to manage multidrug‐ resistant infections and reduce the risk of hospital‐ associated infections Reduce the prevalence of MDROs in patients Prudent antimicrobial use ●● Therapy directed by culture and susceptibility testing ●● Withholding therapy (as appropriate) for patients only colonized and not infected ●● Discontinue use of higher risk devices (e.g., urinary and intravenous catheters) when no longer medically required ●● Infection control practices ●●

Salmonella spp. Salmonella has been identified as a source of sporadic illness and hospital‐associated outbreaks in small animal hospitals. A notable concern with Salmonella HAIs is the occurrence of zoonotic transmission with accompanying human infections. Factors increasing the risk of Salmonella shedding in small animals include consumption of raw meat diets and exposure to livestock. As with E.coli, ESBL‐producing strains are a concern for antimicrobial resistance and have been identified in small animals. Acinetobacter spp and Pseudomonas spp. As opportunistic pathogens in small animals and able to persist in the environment for extended periods, Acinetobacter and Pseudomonas are important in veterinary HAIs. Documented HAIs involving A. baumannii include intravenous and urinary catheters, surgical drain infections, SSIs, pneumonia, and BSIs. SSIs, including cardiovascular device infections, have been attributed to P. aeruginosa. Identification of hospital clusters of Pseudomonas infections should prompt investigation of potentially contaminated environmental, equipment (e.g., endoscope) or consumable (e.g., catheter preparation supplies) sources.

­Management Although complete prevention of HAIs and MDROs is our goal, given the nature of medicine, bacterial adaptation, and numerous mechanisms for antimicrobial resistance, it is inevitable they will continue to occur. Methods to manage MDR infections and reduce the risk of HAIs and nosocomial transmission are paramount (Box 116.1). Efforts should be directed toward prompt identification, appropriate therapy, and infection control practices. Establishing and maintaining an infection control program is vital to integrating these efforts and protecting the health of pets and people. Therapy Antimicrobial therapy is an important part of the treatment of many MDROs and HAIs. Although available antimicrobial choices may be limited, the general

Reduce within‐hospital exposure ●● Hand hygiene ●● Environmental cleaning and disinfection ●● Personal protective equipment (e.g., gloves, gowns) ●● Cohorting patients with similar infectious disease risks ●● Isolation of known or suspected infectious cases Prompt recognition and response ●● Culture and susceptibility testing ●● HAI surveillance (e.g., passive surveillance for HAIs, reporting of pathogens and syndromes of interest) ●● Education of staff and clients HAI, hospital‐associated infection; MDRO, multidrug‐resistant organism.

­ rinciples of therapy are the same as for antimicrobial‐­ p sensitive pathogens: ensuring the appropriate drug (and dose) is selected based on the patient and infection. As infections caused by MDROs appear clinically identical to those caused by antimicrobial‐susceptible bacteria, culture and susceptibility testing are fundamental in this process. Local therapy (e.g., topical therapy, antimicrobial‐impregnated materials, intraarticular injection) is often overlooked, and in many cases can provide adequate drug levels for treatment and may be an alternative for drugs that are not an option for systemic therapy. However, the possibility of systemic absorption of locally administered drugs must be considered. In addition to treating MDROs, it is important to prevent their development. Prudent antimicrobial use will help reduce the risk for HAIs at both the individual patient and hospital population levels. Clinicians should avoid using antimicrobials when a bacterial infection has not been confirmed by culture. Antimicrobials used in the initial treatment of an infection should be selected based on the effectiveness against the most likely organisms causing the infection and penetration into the body site affected. Guidelines are available to assist veterinarians in antimicrobial selection.

116  Nosocomial and Multidrug‐Resistant Infections

Colonization versus Infection Following exposure to an opportunistic pathogen, animals may become colonized (pathogen multiplies ­ without tissue invasion/damage), infected (multiplies with tissue damage and often signs of clinical disease), both, or neither. In animals, decolonization therapy for MDROs has not been shown to be effective and unsuccessful attempts result in promotion of further resistance. As such, therapy should be reserved for animals with clinical disease, regardless of the MDRO involved. Animals without clinical signs of disease, assuming they are otherwise immunocompetent and expected to demonstrate clinical signs should infection occur, may be carefully monitored. Staff and clients should be informed of the colonization status of patients, as these patients may nonetheless shed the pathogen, serving as a source for HAIs or zoonotic infections.

fingers, backs of hands and base of the thumbs, as these areas are often missed. Hand hygiene should occur before and after every animal contact and after ­removing gloves. Appropriate PPE use reduces the risk of contamination of personal clothing, reduces exposure of skin and mucous membranes of veterinary staff to pathogens, and reduces transmission of pathogens between patients by veterinary personnel. The use of PPE is especially important when handling animals with known or suspected MDROs. This includes the use of a barrier gown (disposable or laboratory coat that is not worn elsewhere) and gloves. As animals with MDROs may be colonized or contaminated at locations distant to the infected site, these clothing precautions should be worn when ­having  contact with any MDRO patient or its housing environment.

Patient Housing

Cleaning and Disinfection

In order to protect other patients and clinic staff, attention to patient housing and animal flow is important in managing patients with MDROs. Patient cohorting entails housing together and maintaining a general flow (e.g., assigning waiting and examination areas) of patients that have similar infectious disease risks, including risk of shedding infectious organisms as well as vulnerability to infection. Additionally, animals infected or suspected to be infected with MDROs should be isolated. The specific protocols will vary with the pathogen, severity of disease, and facility, but should include use of personal protective equipment (PPE), cleaning and disinfection, and restricted personnel access. Materials and equipment used for isolation patients should be dedicated to the patient during its hospital stay, after which they should be cleaned and disinfected.

Evidence suggests environmental contamination in human hospitals increases the risk for HAIs and interventions that reduce environmental contamination reduce HAIs. This connection is less well established in veterinary medicine, but it is logical to assume that it exists. Examples of fomites that have been identified as  reservoirs of HA pathogens include stethoscopes, computer keyboards, thermometers, examination tables, and floors. The role of these items in HAIs is not known but it is prudent to minimize contamination. Several key steps must be taken to ensure a disinfectant is effective, including ensuring the surface/item is clean and the product is applied at the manufacturer’s suggested dilution and contact time (amount of time the disinfectant is in contact with the item before being removed). Disinfectants should be selected based on a number of criteria including the product’s spectrum of activity, susceptibility to inactivation by organic matter, and potential pathogens in the environment. Resources are available to guide disinfectant selection (see Chapter 117). Antimicrobial resistance does not necessarily indicate further resistance to environmental disinfectants, although the consequences for inadequate removal/ decontamination are greater. As such, it is prudent to consider the use of a “broad‐spectrum” disinfectant (e.g.,  oxidizing agents), for contact surfaces of patients with HAIs or MDROs.

Hand Hygiene and Contact Precautions Hand hygiene and use of PPE, such as nonsterile gloves and gowns, are simple techniques that can reduce the risk of HAIs. A number of studies indicate that veterinarians and staff typically do a poor job at performing hand hygiene between patients or using PPE when indicated; improvement in these areas is desperately needed. Hand hygiene is described as the single most effective and underutilized infection control measure. Use of hand hygiene limits the spread of organisms between patients and between patients and staff. Biocidal soap and water and alcohol‐based hand sanitizers (AHS) are most frequently used for this purpose. Individuals should ensure hands have soap or AHS contact for at least 15 seconds, paying attention to fingertips, between

Zoonotic Concerns During their careers, many veterinarians report a major animal‐related injury resulting in lost work or hospitalization. Although not responsible for the majority of

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these injuries, HAIs such as MDROs (e.g., MRSA, Salmonella) do occur. Educating staff and clients on zoonotic disease risks and enforcing hospital infection control protocols to reduce these risks will be beneficial to the health of people and patients.

­Conclusion Hospital‐associated infections are reported in veterinary medicine and their frequency is likely to increase with  the rise in intensive care practices. Prolonged

­ ospitalization and the use of invasive devices and proh cedures increase the risk of HAIs. All staff members should be educated on the risks and signs associated with HAIs so that cases can be detected early and managed appropriately. A multifaceted approach is necessary to address HAIs and MDROs in small animal veterinary medicine, including prudent antimicrobial use, strengthening ­surveillance in companion animal species, improving infection control practices, instilling an infection control culture amongst veterinary staff, and improving healthcare worker and public education regarding antimicrobials.

­Further Reading Benedict KM, Morley PS, van Metre DC. Characteristics of biosecurity and infection control programs at veterinary teaching hospitals. J Am Vet Med Assoc 2008; 233: 767–73. Canadian Committee on Antibiotic Resistance. Infection Prevention and Control Best Practices for Small Animal Veterinary Clinics. www.wormsandgermsblog.com/ files/2008/04/CCAR‐Guidelines‐Final2.pdf (accessed June 26, 2019). Papich MG. Antibiotic treatment of resistant infections in small animals. Vet Clin North Am Small Anim Pract 2013; 43: 1091–107.

Stolle I, Prenger‐Berninghoff E, Stamm I, et al. Emergence of OXA‐48 carbapenemase‐producing Escherichia coli and Klebsiella pneumoniae in dogs. J Antimicrobial Chemother 2013; 68: 2802–8. Weese JS, Blondeau JM, Boothe D, et al. Antimicrobial use guidelines for treatment of urinary tract disease in dogs and cats: Antimicrobial guidelines working group of the International Society for Companion Animal Infectious Diseases. Vet Med Int 2011; 2011: 263768.

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117 Management of Infectious Disease in Kennels and Multicat Environments: Creating a Culture of Compliance Frank Bossong, DVM College of Veterinary Medicine, Western University of Health Sciences, Pomona, CA, USA

Dog kennels and multiple‐cat environments exist in ­various types of venues. Whether a boarding facility, a cattery, dog breeding kennel, animal control shelter or humane society, challenges arise in minimizing and managing infectious disease in environments where multiple individuals co‐habit in confined areas. Minimi­ zing the risks, understanding modes of transmission, promoting prevention, taking a structured approach to disinfection, and, most importantly, creating a culture of compliance for established protocols can be helpful in the prevention and control of infectious disease in these settings. Not only will the health of the canine and feline residents be promoted, but such an approach protects the human caretaker from zoonotic disease transmission.

­Risks Some of the major risks for the spread of infectious dis­ ease include poor facility design, overcrowding, intro­ duction of new individuals with undetermined health status, untrained/noncompliant staff, and lack of policy, procedures, and/or biologic risk management (BRM) protocols for managing infectious disease. Overcrowding increases exposure through direct con­ tact between animals and increases stress, which can lead to decreased immunity. Stress can also lead to reac­ tivation of latent infection, which causes increased shed­ ding, and increased infection across the population. The presence of younger individuals and others with naive immune systems (not fully vaccinated) makes the popu­ lation more susceptible to disease outbreak. The housing of animals according to immune status should be imple­ mented, with the naive individuals (young and/or unvac­ cinated) housed separately from the adults. Minimum space requirements should be considered for each facility. The Humane Society of the United

States and the Animal Welfare Act have set minimum standards to follow. Dogs in group housing should have a minimum of 4 ×4 feet of floor space per individual. ­Co‐housed cats should have at least 10.8 square feet per cat and an additional 2.5 square feet for cats that are ­co‐housed together. Ventilation is also important, with an optimum exchange rate of air inside a building replaced with fresh outside air 8–15 times per hour. Safe outside housing is ideal in terms of air circulation and has the added benefit of sunlight and ultraviolet light, as they reduce the number of microorganisms and can inactivate viruses, bacteria, fungi, and mycoplasma. The materials in the facility need to be nonporous to allow for proper cleaning and disinfection of surfaces.

­Modes of Transmission Understanding of the modes of transmission is helpful in determining the preventive measures that need to be taken. Aerosol transmission is affected by ventilation and distance between individuals. Oral ingestion of con­ taminated food/water and oral contact with inanimate objects (fomites) such as toys, cages, bedding, etc. are two of the most common routes of disease transmission. Direct contact or animal‐to‐animal contact (nose to nose) is also common. Vector transfer of disease via arthropods is another cause for concern. Environmental transmission necessitates proper sanitation protocols.

­Prevention Prevention is paramount to success in the management of infectious disease outbreaks. The staff and supervisors of facilities need to understand and identify the risks and then develop infection control policies and procedures for the facility and education and training protocols for the staff.

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Upon intake to a facility, animals must be evaluated via physical exam and checking of medical records in order to identify any problem(s) that may be present and develop a preventive health plan for each animal. Staff need to be aware of the clinical signs to look for, which diagnostic tests need to be run (e.g., FeLV/FIV test, SNAP® Parvo Test) and what preventive measures need to be taken (e.g., vaccination protocol followed). If a dis­ ease is diagnosed, it is neccessary to follow a protocol on how to treat and where to house ill animals, and how to monitor for additional spread in order to succeed in managing an outbreak. Numerous pathogens need to be addressed and under­ standing the agent, mode of excretion, incubation, shed­ ding, and carrier states is key to managing them. The Koret Shelter Medicine website is one valuable resource for providing specific profiles and protocols for address­ ing diseases (www.sheltermedicine.com). Along with these management strategies, prevention involves vaccination and diagnostic testing protocols and the use of proper sanitation to prevent transmission. Improving immune status of individuals should be addressed via vaccination, good nutrition, and decreas­ ing stress. Prevention is also accomplished by screening at a designated receiving area and establishing proper isolation and quarantine areas. Staff should understand that with most diseases, isolation of 14 days is a minimal holding time for ill animals. Less than seven days is of little benefit for most infectious diseases.

­Vaccination Vaccination provides an additional safeguard to the health and well‐being of cats and dogs. The 2017 American Animal Hospital Association (AAHA) Canine Vaccine Guidelines and the 2015 World Small Animal Veterinary Association (WSAVA) Guidelines for the Vaccination of Dogs and Cats provide a comprehensive look at which vaccines are core and noncore, and how protocols may differ between canines seen in general veterinary practice and those in shelter housing (kenneled). For felines, the 2013 American Association of Feline Practitioners Feline Vaccine Advisory Panel Report is an excellent resource for the core, noncore, and not recom­ mended feline vaccinations. Timing can be critical. In shelters, core vaccines should be given as soon as possible and if vaccination status of an incoming animal is unknown, some might argue that it is best to assume that it is unvaccinated. In boarding ken­ nels, proper timing of vaccination (such as Bordetella bronchiseptica) well in advance of boarding an individual is warranted. Understanding the role of maternal immunity

and how it affects vaccination protocol and proper follow‐ up of additional boosters in puppies and kittens is crucial for adequate protection of these individuals.

­Cleaning and Disinfection The importance of sanitation is often underestimated and even though immunization for infectious disease is a valuable tool, it is good to remember that one “cannot vaccinate one’s way out of an outbreak.” If the pathogen load is great enough, a vaccinated animal can still suc­ cumb to disease. It is important to know which disinfectants are the right choices for routine disinfection protocols and what is appropriate when an outbreak occurs. Determine whether broad‐spectrum activity or targeting of a spe­ cific isolated organism is needed and then choose the appropriate disinfectant, correct concentration, applica­ tion, and contact time. It is important to know which dis­ infectants are needed to kill bacteria, enveloped viruses, nonenveloped viruses, fungus/ringworm, parasite eggs, and spores. These are listed from easiest to hardest to kill. Disinfectant failure occurs for a number of reasons. Often, the wrong choice is made for a given pathogen or there is under‐ or overdilution of the product. Failure to rid a surface of organic material, lack of adequate appli­ cation, wrong temperature, humidity and especially lack of contact time (no less than 10 minutes with most disin­ fectants) are additional pitfalls to avoid. The establish­ ment of a “disinfection action plan” includes assessment, cleaning, washing, disinfection, and evaluation. Washing is considered to be the most crucial step in the disinfec­ tion process and can result in the removal of 99% of bac­ teria present in a contaminated area. Hand washing cannot be overemphasized. Human hands play a major role in the transmission of infectious disease. A “bare below the elbows” policy allows all indi­ viduals an easier opportunity to wash the hands, wrists, and forearms after contact with animals. Running water and soap should be provided in as many areas of a facility as possible, especially in isolation areas. Understanding the limitations of hand sanitizers is important. These alcohol‐based products do not kill nonenveloped viruses like canine parvovirus and panleukopenia and are easily deactivated in the presence of organic material on unclean hands. However, the World Health Organization (WHO) recommends that these alcohol‐based products are cur­ rently “the only means for rapidly and effectively inacti­ vating a wide array of potentially harmful microorganism on hands.” Personal protective equipment (PPE) is also important for staff at animal facilities as clothing is a common fomite. Using gloves, lab jackets or a designated uniform and proper disposal or decontamination of these

117  Management of Infectious Disease in Kennels and Multicat Environments: Creating a Culture of Compliance

items are crucial to preventing the spread of pathogens. This is especially important when handling sick animals or when working in a designated isolation area. Quaternary compounds are commonly used and some of the brand names are misleading in terms of what organisms they are effective against. The Center for Food Security and Public Health (www.cfsph.iastate.edu) pro­ vides charts of both the characteristics and antimicrobial spectrum of various products. These should be routinely used as a reference when decisions about appropriate disinfectant choice are needed. In general, quaternary compounds are not effective against canine parvovirus and panleukopenia, both nonenveloped viruses that can have fatal consequences for individuals in kennels and catteries respectively. The hydrogen peroxide‐based oxi­ dizing agents are effective against some of these microbes. Bleach can be very effective but only if proper dilution, length of efficacy after mixing, and proper application protocols are followed. Concentrations greater than 1:32 can be harmful to mucous membranes, thus increasing susceptibility to disease. When cleaning and disinfecting cages, feline patients should be removed from the area when deep cleaning is involved. It is considered a stand­ ard of care to remove dogs from kennel runs when they are being sprayed down. Phenolic disinfectants (“Lysol”) should not be used in catteries as they are toxic to felines.

­Managing Outbreaks Recognizing an outbreak as early as possible provides the best prevention. Controlling the outbreak requires isolation, proper diagnosis, and treatment. In an out­ break situation, an isolation area can help prevent cross‐ infection between animals. Separate equipment needs to be stored in this area and the correct PPE and proto­ cols need to be followed by staff. Protective outer equip­ ment should be used when cleaning, and it should be cleaned and changed daily. Disposable shoe covers are used when heavy infection load is suspected and thrown away after use.

­Culture of Compliance One of the biggest risks to proper disease control in ken­ nels and multicat facilities is the lack of an established policy and procedure and the lack of staff compliance with established protocols. Staff need to be aware that their hands and fomites like clothing and equipment may be the biggest risk to the spread of disease in a facility. Outbreaks can be avoided and/or their impact minimized if good protocols exist. However, if there is no compliance from the staff, even the best protocols will be ineffective.

Many protocols are ready to follow and the staff has the ability to perform them. Their motivation to do so, on the other hand, may or may not result in compliance. The idea of establishing an “infection control culture” may be fundamental in providing effective infectious dis­ ease control. What people “read, hear, see, and talk about on a regular basis” are what characterize the idea of “cul­ ture.” The “read” component would be the establishment of a written infection control manual which would be used to train new staff and which existing staff could refer to when needed. The “hear” component comes from supervisors in terms of staff updates about protocol changes and encouraging conversation about the protocol during regular staff meetings. Posters about hand washing and use of hand sanitizers, for example, help as a visual reminder. Additionally, supervisors and senior staff can serve as an example to follow, further supporting the “see” component of the equation. Giving the staff adequate time to perform hand hygiene, proper cleaning and disin­ fection, changing PPE and other tasks demonstrates the importance that is placed on these practices, and thereby contributes to the infection control culture. The concept of compliance is a critical factor here. As per the Compendium of Veterinary Precautions, “a written check­ list, which specifies the frequency of cleaning, disinfection procedures, products to be used, and the staff responsible, should be developed for each area of the facility…”.

­Protection from Zoonotic Pathogens The emergence of new or evolving pathogens in kennels and catteries will remain a challenge to managing ­disease. For example, in kennels, Streptococcus zooepidemicus is one such pathogen and clinical signs are similar to those of kennel cough. The bacterium is able to spread between dogs as outbreaks at kennels often affect large numbers of animal within a short time. No vaccine exists but good husbandry and isolation of infected individuals are a must. Again, early detection, isolation, and disinfection provide the best management practice and have been found to be fundamental for the control of S. zooepidemicus outbreaks. Good hygiene standards will also minimize the risk of transmission of S. zooepidemicus to dog handlers.

­Conclusion When following effective protocols for managing infec­ tious disease in kennels and multicat environments, the outcome is not only reduced transmission from animal to animal but also the reduction of potential zoonotic transmission. Again, S. zooepidemicus provides an

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excellent example. In order to improve control of this disease, it is crucial to raise awareness, to ensure that cases are recognized early. This will allow the imple­ mentation of suitable protocols to minimize the spread of infection along with measures to protect staff from zoonotic infection. The National Association of State Public Health Veterinarians has produced the Compendium of Veterinary Standard Precautions (www.nasphv.org), which provides a resource for PPE,

environmental infection control, and isolation of a­nimals with infectious diseases. Extensive resources are readily available to enhance the management of infectious disease in kennels and multicat environ­ ments. With the establishment of an infection control culture as well as a culture of compliance, the spread of infectious disease can be minimized, thus protecting the health of cats and dogs as well as their human caretakers.

­Further Reading Miller L, Hurley K. Infectious Disease Management in Animal Shelters. Ames, IA: Wiley‐Blackwell, 2009. Miller L, Zawistowski S. Vaccination strategies in the animal shelter environment. In: Shelter Medicine for Veterin­arians and Staff. Ames, IA: Blackwell Publishing, 2004. Newbury S, Blinn, M, Bushby P, et al. Guidelines for Standards of Care in Animal Shelters. www.sheltervet. org/assets/docs/shelter‐standards‐oct2011‐wforward. pdf (accessed June 26, 2019).

Priestnall S, Erles KE. Streptococcus zooepidemicus: an emerging canine pathogen. Vet J 2011; 188: 142–8. Scherk MA, Ford RB, Gaskell R, et al. AAFP Feline Vaccine Advisory Panel Report. J Feline Med Surg 2013; 15: 785. www.wsava.org/sites/default/files/WSAVA%20Vaccination %20Guidelines%202015%20Full%20Version.pdf www.catvets.com/guidelines/practice‐guidelines/feline‐ vaccination‐guidelines https://www.aaha.org/aaha-guidelines/vaccination-canineconfiguration/vaccination-canine/

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118 Disorders of Sodium and Water Homeostasis Julien Guillaumin, DVM, DACVECC1 and Stephen DiBartola, DVM, DACVIM (SAIM)2 1 2

Department of Clinical Sciences, Colorado State University, Fort Collins, CO, USA Department of Veterinary Clinical Sciences, Ohio State University, Columbus, OH, USA

Sodium (Na) and water disorders traditionally have been viewed by veterinarians as difficult concepts. An appreciation of the basic physiology underlying these concepts will facilitate their understanding. The objectives of this chapter are to review the pathophysiology of changes in volume and composition of the body fluid compartments associated with hyponatremia and hypernatremia as well as a clinical approach to fluid therapy that takes into account these changes in sodium concentration.

­Water Balance Made Easy Imagine two compartments separated by a semipermeable membrane. Each compartment has the same osmolality (number of solute particles per kilogram of solvent). If solute (without water) is added to compartment #1 without changing the volume of the compartment, the concentration of osmoles in that compartment will increase transiently. The increase in osmolality in compartment #1 will result in movement of water (by osmosis) from compartment #2 to compartment #1 until the osmolality of the two compartments is equalized. The final osmolality will be the same in both compartments, and higher than the initial osmolality. The same concept will apply if the volume of water or number of sodium ions in body fluids is changed: water will flow from one compartment to the other in order to maintain the same osmolality in both compartments. Thus, there are only four ways to change serum sodium concentration: adding free water, removing free water, adding sodium, or removing sodium. Hyponatremia is the most common sodium disturbance in small animals. It can result from an excess of free water (e.g., excessive water consumption, intravenous fluid therapy with hypotonic fluids, antidiuretic hormone [ADH]

secretion) or from loss of sodium (in excess of water), which is rare because sodium usually is followed by water. Rare examples include salt‐losing nephropathy and cerebral salt‐wasting syndrome. Hyponatremia usually means that an excessive amount of water is present in body fluids, and the generalization “hyponatremia means too much water” applies in most situations. Treatment of hyponatremia requires understanding and resolution of the primary process. Conversely, the generalization “hypernatremia means not enough water” also is true most of the time. Hypernatremia may arise either as a consequence of loss of free water (i.e., evaporation from the skin or ­respiratory tract) or hypotonic fluid, or by the addition of sodium (e.g., intravenous fluid therapy with 0.9% saline or hypertonic saline). Treatment of hypernatremia mainly is based on slow volume replacement to avoid detrimental transcellular shifts of water, as will be ­discussed later.

­ odium and Water Disturbances: S A Little Physiology Goes a Long Way The Darrow–Yannet diagram (Figure  118.1) illustrates the partition of total body water (TBW) into compartments. Although some variation exists due to species, age or sex, it is estimated that: ●● ●●

total body water is 60% of body weight (BW) total body water is divided into two compartments: –– intracellular fluid (ICF): approximately 2/3 of TBW – 40% of body weight –– extracellular fluid (ECF): approximately 1/3 of TBW – 20% of body weight –– ECF is further subdivided into the interstitial compartment (3/4 of ECF) and the intravascular compartment (IV) (1/4 of ECF).

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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­Osmoregulation and Volume Regulation

ECF ICF

Int

IV

B A

Intracellular fluid compartment

Extracellular fluid compartment

Figure 118.1  Partition of total body water (TBW) into body compartments. See text for details.

To understand changes in serum sodium concentration, it is important to remember that movement of water between ICF and ECF (arrow A on Figure  118.1) depends on osmolality, whereas movement of water within the ECF (arrow B on Figure  118.1) depends on Starling’s forces, which will not be reviewed here. An estimation of osmolarity is given by the following equation: Plasma osmolarity mOsm / L 2 Na K BUN mg/dL / 2.8 glucose mg/dL /18 It is noteworthy that osmolality is a function of the number of solute particles and not their molecular weight. If using international units (mmol/L) for BUN or glucose, the dividing factors shown above are not necessary. The dividing factors convert conventional units into international units by dividing the conventional units by molecular weight (i.e., 28 g/mol for urea or 180 g/mol for glucose) and multiplying by 10 to convert deciliters (dL) to liters (L). A very abundant molecule such as albumin (mean plasma concentration of 4.5 g/dL) has a minimal effect on osmolality because its molecular weight is very high (66 000 Da equivalent to a molar mass of 66 000 g/mol), which makes the particle number relatively low. Albumin contributes only approximately 0.75 mOsm/kg to plasma osmolality. By the same token, colloid osmotic pressure refers to the osmolality due to proteins, and contributes approximately 1–2 mOsm/kg. The osmolarity calculated by the equation above and expressed in mOsm/L is always lower than the osmolality measured by freezing point depression osmometry and expressed in mOsm/kg. The so‐called osmolal gap is the contribution of osmoles other than sodium, potassium, urea, and glucose and usually is 20 mEq/L, indicating that the patient is not in a sodium‐retaining state (i.e., not hypovolemic) and that  the ADH secretion is in fact inappropriate (see Table 118.2). The decreased urine output will not respond to fluid administration, which is the first indication for the clinician faced with a patient with decreased urination, and measurements of urine sodium concentration and osmolality are necessary for the diagnosis of SIADH. The usual treatment for SIADH is fluid restriction and treatment of the primary cause, allowing the body to reset its ADH trigger (usually within 24–72 hours). Most criticalists will treat patients with SIADH using a low dosage (e.g., 0.2–0.5 mg/kg) of furosemide. Clinical Manifestations of Hyponatremia It is not possible to predict serum sodium concentration based on history and physical examination. Measurement of serum sodium concentration is needed as well as some other clinicopathologic data to rule out pseudohyponatremia. Clinical signs of hyponatremia are nonspecific (e.g., nausea, vomiting), and related to dysfunction of the central nervous system, such as lethargy, disorientation, and diminished reflexes. Most human patients do not manifest clinical signs until serum sodium concentration is below 125 mEq/L, although rapid decrease in serum

Table 118.2  Syndrome of inappropriate ADH secretion summary Identify appropriate trigger Postoperative nausea ●● Pain or stress ●● Neoplasia (common) ●● Pulmonary disease ●● Neurologic disease ●● Trauma ●● Drugs (e.g., NSAIDs, narcotics, vincristine) ●●

Treatment Stop/decrease fluid therapy ●● Treat primary trigger ●● May provide small dose of furosemide ●●

Diagnosis Plasma hypoosmolality a ●● Normovolemia (i.e., absence of hypovolemia) ●● Increased urine specific gravity/osmolality (>100 mOsm/kg) ●● Urine sodium concentration >20–40 mEq/L indicating absence of an RAAS trigger (i.e., absence of hypovolemia)b ●●

Notes Low urine output not responsive to fluid therapy ●● Age is a risk factor in humans ●● Case reports of SIADH have been published in veterinary medicine ●●

NSAID, nonsteroidal antiinflammatory drug; RAAS, renin‐angiotensin‐aldosterone system; SIADH, syndrome of inappropriate ADH. a  Spot urine sodium concentration can be influenced by administration of steroids or diuretics, osmotic diuresis (e.g., glucosuria, mannitol administration, postobstructive diuresis), chronic kidney disease and bicarbonaturia (e.g., proximal renal tubular acidosis). b  A weight gain is common in SIADH (author’s personal observation).

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As mentioned earlier, the Adrogué–Madias formula that calculates the change in serum sodium concentration with infusion of a liter of a given fluid also can be used: Change in serum sodium concentration Na K content in 1 Lof solution Na measured / 0.6 body weight kg 1 In the example of a 10 kg dog and using 5% dextrose: Change in serum Na Change in serum Na Change in serum Na

0 175 / 0.6 10 1 175 / 7 25







This formula indicates that 1 L of 5% dextrose in water will decrease the serum sodium concentration by 25 mEq/L. To decrease it to normal (i.e., 145 mEq/L) from 175 mEq/L, 1.2 L of 5% dextrose in water will be necessary, given over 30–60 hours (at a rate of 0.5–1 mEq/ L/h) and thus the infusion rate should be 20–40 mL/h. The water deficit formula has been challenged for several reasons and may underestimate the true water deficit by as much as 50%. First, it assumes that total body water remains unchanged during hyponatremic states. Also, it does not account for low solute fluid losses (i.e., hypotonic fluids), such as urine and gastrointestinal secretions (i.e., it only accounts for free water loss). Likewise, it does not account for any ongoing hypotonic losses. The Adrogué–Madias equation results in the same volume of free water, and thus also has been challenged. The important clinical implication is that serial monitoring (q4–6h) is necessary to reevaluate serum sodium concentration and serum osmolality, and adjust the fluid infusion rate, which is invariably necessary. Ongoing losses of hypotonic fluid also can persist and make the calculation unpredictable. Some clinicians (personal communication, Dr Steve Haskins) use the quick rule of thumb that “3.7–mL/kg/h of free water will decrease the serum sodium concentration by 1 mEq/L per hour” and reassess the serum sodium concentration after 4–6 hours. For the example above, the animal would have received 37 mL/h. As mentioned previously, acute changes in serum sodium concentration (e.g., accidental sodium loading in parenteral fluids) can be corrected more rapidly. This is also true for patients with severe clinical signs due to chronic changes in serum sodium concentration, when the serum sodium concentration can be corrected rapidly until clinical signs resolve, before adopting a more chronic approach. The approach described earlier does not account for any isotonic fluid losses or for the patient’s maintenance fluid requirement. If 0.45% NaCl is used (sodium con-

centration of 77 mEq/L), the rate should be adjusted because only half of this solution represents free water. The patient above should receive 36–71 mL/h, which represents a substantial volume and potentially could worsen brain edema. Thus, the lowest tonicity fluid (and hence volume) is always recommended. If 5% dextrose in water is contraindicated (e.g., hyperglycemic patients) and only if a central venous catheter is in place, some clinicians have advocated using intravascular sterile water. When administered through a central intravenous catheter, sterile water does not cause clinically relevant intravascular hemolysis. Shock Resuscitation of the Chronically Hypernatremic Patient

Resuscitation of volume‐depleted hypernatremic patients, or those with concomitant isotonic fluid losses, should be done carefully, with the sodium content of the fluid matching the animal’s serum sodium concentration within 10 mEq/L to avoid decreasing the serum sodium concentration too rapidly. Consider again the 10 kg dog with a serum sodium concentration of 175 mEq/L, and add the clinical situation of hypovolemic shock. If lactated Ringer’s solution (LRS) is used and the dog requires administration of 1 L of LRS over a one‐hour period, we can calculate the impact of 1 L of LRS on the patient’s serum sodium concentration. LRS has a sodium concentration of 132 mEq/L, so 75% (i.e., 132/175) of the solution is isotonic to the patient’s plasma and 25% is free water. With the administration of 1000 mL of LRS, the patient actually will receive the equivalent of 250 mL of free water, decreasing its serum sodium concentration from 175 to 168 over one hour and possibly leading to cellular swelling, brain edema, seizures, and coma. The use of a fluid isotonic to the patient is necessary for shock resuscitation of a chronically hypernatremic patient. If LRS is used, the addition of 10 mL of 23.4% NaCl (4 mEq/ mL of Na) to a liter of LRS will create a balanced electrolyte solution with a sodium concentration of 172 mEq/L (= 132 + 10 × 4) that will be very close to the patient’s serum sodium concentration of 175 mEq/L. Once shock is treated, the clinician can focus on the free water deficit. A similar approach also can be used for shock resuscitation of the chronically hyponatremic patient.

­Conclusion A good understanding of sodium and water balance is necessary to appropriately identify and treat chronic and acute changes in serum sodium concentration in small animal patients. The proper approach is always the one of the conscientious clinician: a thorough history and complete physical examination, appropriate diagnostic

118  Disorders of Sodium and Water Homeostasis

tests (including serum sodium concentration and urine electrolyte concentrations if needed), appropriate calculation of sodium (or water) deficit, and careful monitoring of changes in the serum sodium concentration and

osmolality in order to avoid deleterious effects of treatment. With these concepts in mind, sodium and water balance becomes both exciting and challenging at the same time.

­Further Reading Adrogué HJ, Madias NE. Hypernatremia. N Engl J Med 2000; 342(20): 1493–9. Cheuvront SN, Kenefick RW, Sollanek KJ, Ely BR, Sawka MN. Water‐deficit equation: systematic analysis and improvement. Am J Clin Nutr 2013; 97(1): 79–85. DiBartola SP. Disorders of sodium and water: hypernatremia and hyponatremia. In: Fluid, Electrolyte, and Acid–Base Disorders in Small Animal

Practice, 4th edn. St Louis, MO: Elsevier Saunders, 2011, pp. 47–79. Hillier TA, Abbott RD, Barrett EJ. Hyponatremia: Evaluating the correction factor for hyperglycemia. Am J Med 1999; 106(4): 399–403. Worthley LI. Hyperosmolar coma treated with intravenous sterile water. A study of three cases. Arch Intern Med 1986; 146(5): 945–7.

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119 Disorders of Phosphorus and Magnesium Rosanne Jepson, BVSc, MVetMed, PhD, DACVIM (SAIM), DECVIM-CA, PGCertVetEd, FHEA, MRCVS Royal Veterinary College, University of London, London, UK

­Phosphorus Phosphorus is vital for many physiologic roles within the body, including skeletal development and structure, cell metabolism and acting as a urinary and intracellular buffer. Disorders of phosphorus homeostasis are identi­ fied with relative frequency in clinical practice and war­ rant consideration of their underlying etiology, careful monitoring and, in certain circumstances, will require therapeutic intervention. Phosphorus can be found in the body in two forms: organic and inorganic (orthophosphoric and pyrophos­ phoric acid). Approximately 85%, of total body phospho­ rus is found in an organic form as hydroxyapatite (Ca10(PO4)6OH2) in bone and as fluorapatite (Ca5(PO4)3F) in teeth. Only 14% of total body phosphorus is found intracellularly and 1% within the extracellular fluid (ECF) compartment. Phosphorus is the principal anion in cells where it is a component of phospholipids and phosphoproteins and incorporated in the structure of nucleic acids and nucle­ otides. Phospholipids (e.g., phosphatidylcholine, phos­ phatidylserine) are an important part of the lipid bilayer of cell membranes and are required for platelet aggrega­ tion. Phosphorus can also be found in cyclic adenosine monophosphate (cAMP), which is required for intracel­ lular signaling pathways and is incorporated in the high‐ energy phosphate bonds of adenosine triphosphate (ATP) and guanosine triphosphate (GTP). Within red blood cells, phosphorus is found in 2,3 diphosphoglycer­ ate (2,3 DPG) which is responsible for the release of oxy­ gen from hemoglobin at the tissue level. Intracellular organic phosphorus can be readily con­ verted to the inorganic form. Rapid translocation between the intracellular fluid and the ECF space can occur, resulting in substantial alteration in ECF concen­ trations. In the ECF, 70% of plasma phosphorus is in an

organic form circulating as phospholipids. The remain­ ing 30% of plasma phosphate exists in an inorganic form where it may be protein bound (10%), complexed to sodium, calcium and magnesium (5%) or circulating free as anions (85%; HPO42‐, H2PO4‐). Regulation of Phosphorus Homeostasis of calcium and phosphorus is intrinsically linked with many of the hormones recognized in the control of calcium also important in the regulation of phosphorus. Traditionally, it was considered that cal­ cium was tightly regulated and that phosphorus concen­ trations were maintained in a reciprocal arrangement. However, it is now appreciated that independent regula­ tion of phosphorus is also important. Phosphorus homeostasis involves three major body organ systems: intestines, bone, and kidneys (Figure 119.1). Plasma phosphorus concentration is the composite effect of dietary phosphorus intake, gastrointestinal absorption, cellular translocation, and renal excretion. In a neutral state of phosphorus balance, daily dietary intake of phos­ phorus will be countered by renal excretion such that total body phosphorus concentrations vary little. The main hormones traditionally cited in the regula­ tion of phosphorus are parathyroid hormone (PTH) and calcitriol (1,25(OH)2 cholecalciferol) (Figure  119.2). Recently, further important hormones referred to as phosphotonins have been identified. Phosphotonins, such as fibroblast growth factor 23 (FGF23), play an important role regulating renal handling of phosphorus. In a similar manner to PTH, FGF23 increases renal tubu­ lar excretion of phosphorus. However, FGF23 has an opposing action to PTH by inhibiting production of cal­ citriol and therefore indirectly reducing release of phos­ phorus from bone and decreasing absorption of phosphorus from the intestinal tract.

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

119  Disorders of Phosphorus and Magnesium

●●

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●●

PTH also increases renal absorption of calcium and stimulates production of calcitriol. The net effect of PTH on the kidney is therefore a reduction in plasma phosphate and an increase in plasma calcium. Increased phosphorus concentration also directly stim­ ulates osteocytes to produce the phosphotonin FGF23. FGF23 acts on the kidney in a similar manner to PTH, decreasing tubular absorption of phosphorus. Calcitriol increases absorption of both calcium and phosphorus from the small intestine. This process is facilitated indirectly by PTH, which increases renal production of calcitriol. FGF23 has an opposing action, inhibiting calcitriol production and therefore decreas­ ing absorption of phosphorus from the intestinal tract. Calcitriol and PTH also increase reabsorption of cal­ cium and phosphorus from bone.

Meat, bone, and fish are food sources high in phospho­ rus and dietary intake is therefore a major source of phosphorus for the body. Typical commercial canine and feline diets have a phosphorus concentration of 0.8–1.6% on a dry matter basis with a calcium to phosphorus ratio of 1.2–2.1. Most organic phosphate is hydrolyzed in the gastrointestinal tract to inorganic phosphate for absorp­ tion. Phosphate is absorbed via paracellular pathways in all regions of the small intestine. However, approximately 30% of intestinal absorption is active and occurs in the duodenum, utilizing a sodium‐phosphorus co‐trans­ porter (NaPi2b). Expression and insertion of the NaPi2b co‐transporter are under the control of calcitriol. A decrease in intestinal phosphate or an increase in calci­ triol will increase intestinal absorption of phosphate. Once absorbed, inorganic phosphate within the extra­ cellular pool is exchangeable with the skeletal stores under the influence of both PTH and calcitriol. However, because proportionally only a very small amount of phosphorus is found in the ECF, measurement of plasma phosphorus concentrations does not always provide a good reflection of total body phosphorus. Inorganic plasma phosphate that is not bound to pro­ tein is freely filtered at the glomerulus. Approximately 85% of filtered phosphate is reabsorbed by active trans­ port in the proximal tubule, with a small amount also absorbed in the distal nephron. However, during periods of low dietary intake, efficiency can increase until almost 100% of filtered phosphate is reabsorbed. Active trans­ port occurs via sodium‐phosphate co‐transporters (NaPi2a and NaPi2c) located in the brush border of proximal tubular cells. Expression of the NaPi2a co‐ transporter is under the control of PTH, FGF23, and intestinal phosphate absorption (Box 119.1). Laboratory Assessment of Phosphorus Circulating inorganic phosphorus (Pi), which can be either free or complexed to calcium, magnesium or

Box 119.1  Regulation of phosphate handling by the kidney In periods of phosphate excess, increase in PTH concentration favors internalization and lysosomal degradation of the NaPi2a co‐transporter, thus limiting phosphate reabsorption and increasing phosphate excretion in the urine. Conversely, in periods of hypophosphatemia, decrease in PTH concentrations has a permissive role, favoring insertion and maintenance of the NaPi2a co‐ transporter in the proximal tubular brush border and increasing phosphate reabsorption. PTH is therefore a phosphaturic hormone promoting excretion of phosphate in the urine. FGF23, produced by osteocytes in response to increased phosphorus concentration, also acts as a phosphaturic factor. FGF23 in conjunction with its co‐factor klotho decreases expression and activity of NaPi2a co‐ transporter in the proximal tubular cells, thus decreasing tubular reabsorption and increasing renal excretion of phosphorus. Other factors which can modulate and increase renal tubular phosphate reabsorption include growth hormone, thyroid hormone, insulin, insulin‐like growth factor 1 (IGF‐1) and 1,25 (OH)2 cholecalciferol. Reabsorption can be reduced by parathyroid hormone‐related protein (PTHrP), glucocorticoids, calcitonin, and atrial natriuretic peptide.

sodium, is the component routinely quantified by diag­ nostic laboratories. Serum phosphate concentrations are slightly higher than plasma phosphate concentrations due to release of phosphate from cells and platelets dur­ ing the clotting process. Normal phosphate reference intervals will be laboratory dependent but values of approximately 2.5–5.5  mg/dL (0.8–1.8  mmol/L) are reported for dogs and 2.5–6.0 mg/dL (0.8–1.9 mmol/L) for cats. It is widely recognized that young animals demon­ strate elevated plasma phosphorus concentrations. This is likely to be the consequence of increased growth hor­ mone facilitating renal tubular reabsorption of phos­ phate and also increased concentrations of calcitriol. In puppies up to 8 weeks of age, plasma phosphate concen­ trations up to 10.8 mg/dL (3.4 mmol/L) can be consid­ ered normal. However, an age‐related difference in phosphorus concentrations can be present until 1 year of age. This difference is more marked in the dog than the cat. Phosphate concentrations can be factitiously increased by lipemia, hyperproteinemia, and hemolysis and can be increased postprandially, particularly after a high‐pro­ tein meal. Patients should therefore ideally be fasted for

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8–12 hours prior to measurement. A number of drugs may influence plasma phosphate concentrations either as a desired therapeutic effect (e.g., intestinal phosphate binders) or as a secondary consequence of their adminis­ tration (e.g., antacids including sucralfate, insulin, and dextrose).

Box 119.2  Causes of hypophosphatemia ●●

Hypophosphatemia Hypophosphatemia can be defined as a plasma phospho­ rus concentration 0.6 mmol/L) and subsequently to bring phosphate concentrations to the lower end of the normal reference interval. Due to concern regarding precipita­ tion of calcium phosphate, phosphorus‐containing fluids should be diluted in saline for administration. Response to supplementation is individual but gradual correction can usually be achieved over a period of hours to days. Patients should be closely monitored for the develop­ ment of hypocalcemia, tetany, and hyperphosphatemia. ­ uring Hyperphosphatemia may be of particular concern d the initial phase of supplementation when pathophysio­ logic mechanisms that have been activated to ensure maximal phosphate retention and reduction in renal tubular excretion during a period of phosphorus reple­ tion have not yet adapted. In patients receiving phospho­ rus supplementation, if the calcium:phosphorus product exceeds 60–70 mg2/dL2 (4.8–5.6 mmol2/L2), soft tissue mineralization and renal failure become a concern. In patients with diabetic ketoacidosis where there is evi­ dence of both hypokalemia and hypophosphatemia, supplementation with potassium phosphate may be ­ preferred. In this situation, approximately quarter of potassium supplementation can be administered as potassium phosphate with the remainder as potassium chloride. Hyperphosphatemia Hyperphosphatemia can occur as a consequence of three main mechanisms: impaired renal excretion, excessive intestinal absorption, and transcellular shift from tissue/ bone to the ECF (Box 119.3). Clinical signs as a direct consequence of hyperphos­ phatemia are rarely observed but can relate to relative hypocalcemia, such as weakness, neuromuscular tetany, and seizures. Hyperphosphatemia may also contribute to  the risk of soft tissue mineralization if the calcium:phosphorus product exceeds 60–70  mg2/dL2 2 2 (4.8–5.6 mmol /L ). Impaired renal function, as a consequence of either chronic kidney disease (CKD) or acute kidney injury (AKI), is the most common clinical cause of hyperphos­ phatemia. Reduction in glomerular filtration rate (GFR) and loss of functioning nephrons result in decreased excretion of phosphorus by the kidney. In the early stages of CKD, initial phosphorus retention is countered by increased production of FGF23, reduction in calcitriol concentrations and, in later stages, increased PTH pro­ duction. These compensatory mechanisms initially main­ tain plasma phosphate concentrations within the reference interval by promoting tubular phosphate excretion in remaining nephrons. However, with p ­ rogressive loss of nephrons and decline in GFR, the compensatory mecha­ nisms are exceeded and hyperphosphatemia ensues.

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Box 119.3  Causes of hyperphosphatemia ●●

●●

●●

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●●

Impaired renal excretion –– Renal* ○○ Prerenal causes, e.g., hypoadrenocorticism ○○ Renal causes, e.g., chronic kidney disease or acute kidney injury ○○ Postrenal causes, e.g., uroabdomen, urinary tract obstruction –– Hypoparathyroidism –– Hyperthyroidism –– (Acromegaly) Increased phosphorus intake –– Gastrointestinal absorption ○○ Vitamin D toxicosis: rodenticide poisoning, e.g.,  cholecalciferol; psoriasis creams, e.g., calcipotriene ○○ Phosphate‐containing enemas –– Intravenous ○○ Administration of phosphorus‐containing fluids Transcellular shifts –– Cellular breakdown ○○ Tumor lysis syndrome ○○ Hemolysis ○○ Rhabdomyolysis ○○ Tissue trauma or hypoxia Physiologic –– Young animal* –– Postprandial* Laboratory error –– Hyperlipemia –– Hyperproteinemia –– Hemolysis

* Commonly identified in clinical practice.

In both dogs and cats, the prevalence of hyperphos­ phatemia increases with advancing stages of CKD. A study in cats demonstrated that 20% of cats with compensated CKD, 49% with uremic CKD and 100% with end‐stage CKD were hyperphosphatemic. Findings were similar in a group of dogs: 18% with International Renal Interest Society (IRIS) stage 1, 40% stage 2, 92% stage 3, 100% stage 4. In AKI, decline in renal function can occur without suf­ ficient time for adequate compensatory mechanisms to develop. Severe hyperphosphatemia may therefore be identified in AKI and may be proportionally greater for the degree of reduction in GFR than identified with CKD. Any prerenal or postrenal causes of reduced GFR, for example hypoadrenocorticism or uroabdomen/urinary tract obstruction respectively, which decrease filtration fraction and renal excretion of phosphorus, can result in hyperphosphatemia.

Studies suggest that up to 43% of cats with hyperthy­ roidism are hyperphosphatemic at diagnosis. In cats that remain nonazotemic, with successful treatment and return of euthyroidism, phosphorus concentrations return to normal. The exact cause of hyperphosphatemia in feline hyperthyroidism has not been fully elucidated. Thyroxine is known to directly increase renal tubular absorption of phosphate. In addition, increased bone isoenzyme alka­ line phosphatase and osteocalcin as a marker of osteoblas­ tic activity suggest that altered bone metabolism may contribute. However, in those cats that are revealed to be azotemic after return of euthyroidism, phosphate concen­ trations do not significantly change and in such patients reduced renal function may also be contributing. Growth hormone can directly increase renal tubular absorption of phosphate and therefore may theoretically contribute to hyperphosphatemia in acromegaly. However, recent studies evaluating the clinicopathologic abnormaltities in larger cohorts of cats with acromegaly have not confirmed this finding. In dogs with hypoparathyroidism, reduction in PTH concentration favors insertion of renal tubular NaPi2a co‐transporters and therefore reabsorption of phospho­ rus. Studies suggest that ~30–100% of dogs with hypoparathyroidism demonstrate hyperphosphatemia. Hyperphosphatemia has also been reported in cats with idiopathic hypoparathyroidism although this is a rare condition. However, clinical signs in primary hypopar­ athyroidism usually relate to marked hypocalcemia rather than any concurrent hyperphosphatemia. Increased intestinal absorption of phosphorus is an uncommon clinical cause of hyperphosphatemia. Ingestion of excess vitamin D, either in rodenticides containing chole­ calciferol or from psoriasis creams containing calcipotriene, can cause increased release of calcium and phosphorus from bone and absorption from the intestinal tract. Absorption of phosphorus from phosphate‐containing enemas has been reported to cause hyperphosphatemia in dogs and cats. Phosphorus is found at high concentrations intracel­ lularly. Any process that results in marked cell destruc­ tion has the potential to increase plasma phosphate concentrations. For example, hyperphosphatemia is one of a number of electrolyte abnormalities identified dur­ ing tumor lysis syndrome (hyperphosphatemia, hypocal­ cemia, hyperkalemia, hyperuricemia, oliguric AKI). Administration of chemotherapy to a patient with a large tumor burden, that is highly sensitive to the chemothera­ peutic agent or radiation dose administered, results in rapid lysis of tumoral cells and release of intracellular phosphorus to the ECF. Acute tumor lysis syndrome has previously been reported in dogs and cats with lympho­ sarcoma. A similar situation can occur during hemolysis, with release of phosphorus from red blood cells or dur­ ing rhabdomyolysis, tissue infarction or severe crush

119  Disorders of Phosphorus and Magnesium

injuries. In rhabdomyolysis, marked myoglobinuria may cause AKI and therefore reduced renal function may also contribute to hyperphosphatemia. Treatment of Hyperphosphatemia

Similar to the management of hypophosphatemia, the first step in the management of hyperphosphatemia is determining and, when possible, correcting the underly­ ing etiology. Renal excretion of phosphorus should be optimized with correction of any prerenal or postrenal component. Intravenous fluid therapy to correct hypov­ olemia and/or dehydration and to provide mild expan­ sion of ECF may improve renal excretion of phosphorus. In patients with AKI, dialysis may offer a way to correct electrolyte imbalances including hyperphosphatemia. However, in those patients with CKD, restricting intes­ tinal absorption of phosphate can control hyperphos­ phatemia. In early CKD (IRIS stage 2), this can be achieved using a phosphate‐restricted diet. However, with advancing CKD, addition of intestinal phosphate binders may be required to achieve phosphate targets, such as aluminum hydroxide, calcium carbonate +/‐ chi­ tosan, calcium acetate, lanthanum carbonate, and seve­ lamer hydrochloride. Dietary phosphate restriction is achieved by feeding a low‐protein, low‐phosphorus diet. Intestinal phosphate binders complex with dietary phos­ phate, producing an insoluble, nonabsorbable com­ pound, which is excreted in the feces. For maximum efficacy, phosphate binders should therefore be adminis­ tered with every meal and used in conjunction with phosphate‐restricted diets. Aluminum hydroxide is a commonly used and well‐ tolerated phosphate binder in canine and feline patients with CKD (30–100 mg/kg/day). Side‐effects can include constipation and in human patients with end‐stage kid­ ney disease or undergoing hemodialysis, aluminum tox­ icity and neurotoxicity are reported. Caution is warranted when using calcium‐containing phosphate binders, such as calcium carbonate, due to the potential for develop­ ment of hypercalcemia, an elevated calcium phosphorus product and dystrophic mineralization. Side‐effects of calcium‐containing phosphate binders can also include nausea and constipation. Sevelamer hydrochloride (30– 40 mg/kg PO q8h) is a resin‐based phosphate binder that contains neither aluminum nor calcium. Sevelamer binds to the mucosal surface of the intestine, allowing extended periods of phosphate binding. However, its expense, gastrointestinal side‐effects, and potential to bind other substances (e.g., vitamins, bile acids and cho­ lesterol) may make it a less desired product. Lanthanum carbonate (dogs 6.25–12.5 mg/kg PO q12h; cats 400– 800 mg/cat/day divided according to feeding schedule) is another nonaluminum, noncalcium‐based phosphate binder, which is capable of binding phosphate at both

low and high pH, being effective therefore in both the stomach and small intestine. Lanthanum undergoes bil­ iary excretion and therefore should not accumulate in patients with compromised renal function. Interventions to control hyperphosphatemia should be carefully monitored by serial measurement of plasma phosphate concentrations. For patients with CKD, the IRIS provides stage‐specific targets for phosphate con­ trol. The effect of either introducing a phosphate‐ restricted diet or phosphate binder or altering dosage of a phosphate binder in patients with CKD can be rela­ tively slow in onset due to whole‐body phosphate reten­ tion. Assessment of plasma phosphate concentration after ~2–4 weeks is therefore likely to be adequate.

­Magnesium Until recently, magnesium has received relatively little attention in small animal medicine. However, magne­ sium is known to play a vital part in many cellular func­ tions and the clinical consequences of alteration in magnesium homeostasis are receiving increased recog­ nition in small animal patients, particularly within a crit­ ical care setting. Distribution in the Body Greater than 99% of the body’s magnesium is located intracellularly, with less than 1% found within the ECF. Of the intracellular stores, ~70% is present in bone, ~20% in muscle, and ~10% other soft tissues. Within the ECF, ~55% of magnesium is ionized or free and considered the biologically active component, with a further 20–30% protein bound and 15–20% complexed to anions (e.g., phosphate or bicarbonate). The relatively smaller pro­ portion of protein‐bound magnesium in comparison to calcium means that magnesium concentrations are less likely to be influenced by changes in albumin concentration. Biologic Importance of Magnesium Magnesium is required in the mitochondria of cells for oxidative phosphorylation and for anaerobic metabolism of glucose. It also plays a role in the synthesis and degra­ dation of DNA, ribosomal binding to RNA, nucleotide synthesis, and production of intracellular messengers such as cAMP. Magnesium is an important regulator of intracellular calcium cycling in both smooth and cardiac myocytes and is therefore important for cardiac excitability, con­ traction, and conduction mechanisms. In vascular smooth muscle, intracellular magnesium concentrations

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may contribute to peripheral vascular resistance; increased intracellular smooth muscle magnesium con­ centrations enhance vasorelaxation whilst conversely decreased concentrations facilitate vasoconstriction. Magnesium is important in neuromuscular transmis­ sion and severe magnesium deficiency results in increased neuronal excitability and neuromuscular transmission. The classic neuromuscular clinical signs of hypomagnesemia are more frequently appreciated in large animal internal medicine where severe muscle tet­ any and seizure are identified in cattle with “grass staggers.” Magnesium is also a vital co‐factor for ATP. Given that ATP is the critical energy source for many cellular ion pumps, such as Na+K+ATPase, HCO3‐ATPase, and Ca2+ATPase, variation in magnesium availability may influence the concentration of intracellular electrolytes such as calcium and potassium. Regulation of Magnesium Magnesium homeostasis is incompletely understood but, in a similar manner to calcium and phosphorus, gastrointestinal absorption, renal reabsorption/excre­ tion, and exchange with skeletal stores are important factors in maintaining total body and extracellular mag­ nesium concentrations. Absorption of magnesium in the intestine primarily occurs in the ileum, with further absorption in the jejunum and colon. Intestinal absorp­ tion of magnesium is via either a passive paracellular or active transcellular route. Active transport of magne­ sium in the gut is likely to involve specific magnesium transport proteins, including those from the transient receptor potential (TRP) family such asTRPM6 and TRPM7. Magnesium is freely filtered at the glomerulus but in contrast to other electrolytes, the primary site for reabsorption is the loop of Henle (60–70%), with lesser contributions in the proximal tubule (15%) and distal convoluted tubule (10–15%). Hormonal regulation of magnesium absorption in the intestine and kidney likely occurs but remains incom­ pletely understood. Parathyroid hormone, calcitonin, glucagon, antidiuretic hormone, aldosterone, and insulin can all increase absorption of magnesium whilst prosta­ glandin E2, hypokalemia, hypophosphatemia, and acido­ sis may all reduce magnesium reabsorption. Laboratory Assessment of Magnesium Making an assessment of total body hypomagnesemia can be challenging due to the high proportion of magne­ sium that is present within an intracellular location. Currently there is no consensus with regard to the opti­ mal methodology to assess total body magnesium status.

Quantification of either total or ionized magnesium concentrations can be performed. However, total serum or plasma magnesium concentrations do not correlate well with signs of hypomagnesemia and may not corre­ late with ionized concentrations. Other factors such as albumin concentrations, sample handling, and acid–base status may influence measured total serum magnesium concentrations. As ionized magnesium equilibrates rap­ idly across cell membranes, it may be a more representa­ tive measure of intracellular magnesium but uncertainty exists as to whether it provides a true reflection of total body magnesium. Quantification of magnesium within red or white blood cells or muscle tissue, and assessment of renal handling of magnesium (e.g., 24‐hour urinary excretion or evaluation of magnesium retention) may give further information regarding magnesium status but these methodologies are not routinely available as clinical diagnostic tests. In those patients where clinical signs are compatible with hypomagnesemia, a low total or ionized magne­ sium concentration will provide support that clinical signs may be attributable to hypomagnesemia. However, it should be appreciated that intracellular hypomagne­ semia can be present with normal serum total or ionized magnesium concentrations. Hypomagnesemia A number of clinical causes of hypomagnesemia can be identified (Box 119.4) reflecting increased gastrointesti­ nal loss, reduced intestinal absorption, redistribution between the intracellular and extracellular compart­ ments, or increased renal excretion. The clinical conse­ quences of hypomagnesemia can include electrolyte and neuromuscular abnormalities and cardiac arrhythmias. However, with mild hypomagnesemia, clinical signs can be vague and attributable to general illness. In human medicine, hypomagnesemia is a commonly reported electrolyte abnormality in critical care popula­ tions and is an independent risk factor for mortality. Studies suggest that in veterinary medicine, between 28% and 50% of cats and dogs in a critical care setting also demonstrate hypomagnesemia but information regarding any association with survival, morbidity or mortality is lacking. Other aspects of illness often identi­ fied in a critical care setting (e.g., sepsis) or associated treatments (e.g., blood transfusions due to citrate admin­ istration, intravenous fluid therapy, insulin administra­ tion) may increase a patient’s predisposition to develop hypomagnesemia. Hypomagnesemia has perhaps received most attention for the role it plays in concurrent electrolyte distur­ bances. Magnesium facilitates potassium leaving the intracellular compartment and enhances renal loss of

119  Disorders of Phosphorus and Magnesium

Box 119.4  Common clinical conditions associated with hypomagnesemia ●●

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Gastrointestinal loss –– Anorexia –– Malabsorption/severe diarrhea –– Short bowel syndrome Renal loss –– Diabetes mellitus/diabetic ketoacidosis –– Chronic kidney disease –– Renal tubular acidosis –– Drugs: loop diuretics –– Renal transplantation Miscellaneous –– Eclampsia –– Redistribution ○○ Severe pancreatitis ○○ Sepsis ○○ Refeeding syndrome ○○ Catecholamine excess (pheochromocytoma) ○○ Large‐volume resuscitation with magnesium‐ replete fluid

potassium such that hypomagnesemia and hypokalemia occur simultaneously. Clinical signs in this situation usu­ ally reflect hypokalemia but potassium deficiency may prove refractory to therapy until magnesium levels have also been corrected. Hypocalcemia has also been reported in conjunction with hypomagnesemia. In human patients, magnesium deficiency has been reported to contribute to the development and severity of atrial fibrillation, supraventricular tachycardia, and ventricular tachyarrhythmias. Information regarding hypomagnesemia in dogs and cats with cardiac disease is limited and the clinical importance is therefore uncer­ tain. However, certain drugs commonly used in cardiac patients such as digoxin and loop diuretics increase magnesium loss. Patients with cardiac disease that develop hypomagnesemia may be at risk of cardiac arrhythmias, decreased cardiac contractility, and refrac­ tory hypokalemia. Consideration could therefore be given to assessment of magnesium concentrations in these situations. Magnesium deficiency due to reduced dietary intake is unlikely, particularly if commercial diets are being fed, but could occur in patients with prolonged anorexia or severe gastrointestinal disease such as chronic diarrhea, malabsorptive disease, and short bowel syndrome. Hypomagnesemia has previously been reported in dogs with protein‐losing enteropathy (PLE) and as a compo­ nent of refeeding syndrome when increased cellular demand occurs in addition to chronic depletion second­ ary to prior anorexia.

Hypomagnesemia has also been reported in associa­ tion with a number of endocrine conditions, for exam­ ple, in cats with hyperthyroidism, diabetes mellitus, and diabetic ketoacidosis. However, the clinical significance of these associations is still uncertain and a study evalu­ ating magnesium concentrations in dogs with diabetes mellitus did not find an association. Hypomanesemia may be identified in lactating bitches although is rarely clinically significant in the healthy bitch. The prevalence of hypomagnesemia in bitches with eclampsia has been reported to be 44%. Treatment of Hypomagnesemia

Magnesium supplementation should be considered when a patient’s clinical signs are attributable to hypomagnesemia. Supplementation may also be indi­ cated in those patients with refractory hypokalemia or hypocalcemia despite appropriate potassium and cal­ cium supplementation respectively. Intravenous magnesium supplementation is usually provided as magnesium sulfate or magnesium chloride. Typical dosages reported are 0.03–0.04  mEq/kg/h administered as a CRI diluted in 5% dextrose or 0.9% saline. Magnesium salt concentrations >20% should not be administered and magnesium salt solutions are not compatible with calcium‐ or bicarbonate‐containing flu­ ids. Careful monitoring should be performed during administration to avoid inadvertent overdosage and sup­ plementation continued until low normal concentrations have been achieved and maintenance magnesium requirements can be provided via the patient’s daily die­ tary intake. Caution should be exercised when providing magnesium supplementation, particularly in those patients with reduced renal function where risk of devel­ opment of hypermagnesemia is increased. The value of chronic supplementation is uncertain in veterinary med­ icine but oral supplementation with magnesium oxide (1–2 mEq/kg/day) has been reported. Reports of magnesium supplementation in the veteri­ nary literature have also included patients with tetanus where supplemental magnesium may aid in reducing muscle spasm and sedative requirements, and in patients with cardiac arrhythmias where hypomagnesmia is deemed contributory. Hypermagnesemia The kidney excretes excess magnesium. Hyperm­ agnesemia may occur with any marked reduction in GFR. It is rare for dogs and cats to demonstrate clinical signs associated with mild to moderate hypermagne­ semia. Clinical signs reported with severe hypermagne­ semia can include depression, weakness, lethargy, flaccid paralysis, and decreased reflexes. Hypotension may be

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documented secondary to decreased total peripheral resistance. Bradycardia may be identified associated with prolongation of the QRS complex and an increased P‐Q interval. Untreated, severe hypermagnesemia will result in ventricular fibrillation, asystole, and death. Iatrogenic hypermagnesemia has been reported in human medicine in children associated with excessive use of magnesium‐containing laxative agents and antacids. In veterinary medicine, iatrogenic hypermagnesemia has been reported in a dog and a cat due to oversupplementa­ tion. These patients demonstrated magnesium concentra­ tions 7–9 times the normal reference interval and clinical signs including hypotension and bradycardia. This empha­ sizes the need for careful dose calculation and monitoring

during magnesium supplementation. Increased magne­ sium concentrations have also been reported in dogs with  hypoadrenocorticism and after surgery/anaesthesia although these changes were not clinically significant. Treatment of hypermagnesemia depends on severity and should involve discontinuation of any supplementa­ tion being provided. Saline diuresis and administration of loop diuretics may increase magnesium renal excre­ tion. In an acute situation with arrhythmias secondary to hypermagnesemia, calcium gluconate should be admin­ istered (50–150 mg/kg) as a bolus over 20–30 minutes to reverse cardiac toxicity. In human medicine, in certain clinical situations, dialysis may also be used for correc­ tion of hypermagnesemia.

­Further Reading Bateman S. Disorders of magnesium: magnesium deficit and excess. In: DiBartola SP, ed. Fluid, Electrolyte and Acid–Base Disorders. St Louis, MO: Elsevier Saunders, 2012, pp. 211–19. DiBartola SP, Willard MD. 2012. Disorders of phosphorus: hypophosphatemia and hyperphosphatemia. In: DiBartola SP, ed. Fluid, Electrolyte and Acid–Base Disorders. St Louis, MO: Elsevier Saunders, 2012, pp. 195–211. Foster JD. Update on mineral and bone disorders in chronic kidney disease. Vet Clin North Am Small Anim Pract 2016; 46(6): 1131–49. Hardcastle MR, Dittmer KE. Fibroblast growth factor 23: a new dimension to diseases of calcium‐phosphorus metabolism. Vet Pathol 2015; 52: 770–84.

Simmonds EE, Alwood AJ, Costello MF. Magnesium sulfate as an adjunct therapy in the management of severe generalized tetanus in a dog. J Vet Emerg Crit Care 2011; 21(5): 542–6. van den Broek D, Chang Y, Elliott J, Jepson R. Prognostic importance of plasma total magnesium in a cohort of cats with azotemic chronic kidney disease. J Vet Intern Med 2018; 32: 1359–71. Williams TL, Elliott J, Syme HM. Calcium and phosphate homeostasis in hyperthyroid cats: associations with development of azotaemia and survival time. J Small Anim Pract 2012; 53(10): 561–71.

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120 Acute Kidney Injury Adam E. Eatroff, DVM, DACVIM (SAIM) ACCESS Specialty Animal Hospitals, Culver City, CA, USA

­Etiology/Pathophysiology Acute kidney injury (AKI) has classically been categorized into hemodynamic (prerenal), renal parenchymal (intrinsic), and postrenal etiologies. Hemodynamic causes include decreases in renal perfusion and/or excessive vasoconstriction, and are characterized as rapidly reversible if the inciting cause is eliminated shortly after occurrence. It is questionable whether hemodynamic changes resulting in azotemia represent true renal injury or an appropriate physiologic attempt to conserve fluid volume as a response to a reduced effective circulating blood volume. Neurohumoral responses to reductions in effective circulating volume result in a reduction in renal blood flow and, thus, glomerular filtration rate (GFR) as a means of conserving fluid volume. Regardless of whether the hemodynamic changes represent pathology or physiology, it is generally accepted that prolonged ischemia can contribute to renal parenchymal injury. Intrinsic causes of AKI include infectious diseases, toxins, and systemic diseases with renal manifestations. Removal of the inciting cause of parenchymal injury is often the only means of directly addressing this type of insult. Postrenal AKI is due to obstruction or diversion of urine flow, including urethral obstruction, bilateral ureteral obstruction or unilateral obstruction with a nonfunctional contralateral kidney, or rupture of any portion of the urinary tract. Restoration of urine flow may rapidly reduce the concentrations of circulating uremic toxins. However, prolonged obstruction of urine flow may lead to renal parenchymal injury. During the diagnostic evaluation for AKI, it is important to recognize that (1) the classic etiologies of AKI frequently produce insults that encompass more than one of the hemodynamic, intrinsic, or postrenal processes and (2) an etiologic cause of AKI is frequently not identified.

Table 120.1 provides an extensive, but not exhaustive, list of etiologies for canine and feline AKI. The pathophysiologic process of AKI is often multifactorial, with overlapping ischemic, inflammatory, toxic, and septic components. Classically, the clinical course of AKI proceeds through four phases. These phases are defined by experimental models of ischemic acute kidney injury and may not be representative of the multifactorial nature of the disease. The initiation phase is characterized by the first stages of renal injury. Intervention at this phase may prevent progression to more severe injury, but injury at this stage occurs on a subcellular level and may not be biochemically evident. During the extension phase, cellular injury progresses to cell death. At this stage, biochemical derangements and clinical manifestations of disease manifest. During the maintenance phase, both cell death and regeneration occur simultaneously, and the potential for and length of recovery from this phase may be determined by the balance between these processes. Removal of the initiating cause at this stage does not alter the existing damage, but may allow for the balance to shift in favor of parenchymal regeneration. The recovery phase is characterized by improvement in GFR and tubular function; this final phase may last weeks to months.

­Epidemiology The epidemiology of AKI has not been thoroughly characterized in veterinary medicine. The difficulty in establishing the population characteristics of this disease is primarily related to the lack of a standard definition for AKI and the wide spectrum of disease (ranging from clinically undetectable, subcellular damage to fulminant, excretory failure). This problem previously

Clinical Small Animal Internal Medicine Volume II, First Edition. Edited by David S. Bruyette. © 2020 John Wiley & Sons, Inc. Published 2020 by John Wiley & Sons, Inc. Companion website: www.wiley.com/go/bruyette/clinical

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Table 120.1  List of etiologies for canine and feline acute kidney injury Nephrotoxins Antimicrobial agents Aminoglycosides Aztreonam Carbapenems Cephalosporins Penicillins Polymyxins Quinolones Rifampin Sulfonamides Tetracyclines Vancomycin

Antifungal agents Amphotericin B

Antineoplastic drugs Cisplatin and carboplatin Doxorubicin Methotrexate

Antiviral agents Aciclovir Foscarnet

Antiprotozoal agents Dapsone Pentamidine Sulfadiazine Thiacetarsamide Trimethoprim‐sulfamethoxazole

Immunosuppressive drugs Azathioprine Calcineurin inhibitors (e.g., ciclosporin, tacrlimus) Interleukin‐2

Miscellaneous therapeutic agents Acetaminophen Allopurinol Angiotensin enzyme converting inhibitors Antidepressants Apomorphine Cimetidine Deferoxamine Dextran‐40 Diuretics epsilon‐Aminocaproic acid EDTA Lipid‐lowering drugs Lithium Methoxyflurane Nonsteroidal antiinflammatory drugs Penicillamine Phosphorus‐containing urinary acidifiers Streptokinase Tricyclic antidepressants Vitamin D analogs

Endogenous compounds Hemoglobin Myoglobin (e.g., trauma/ rhabdomyolysis)

Heavy metals Antimony Arsenic Bismuth salts Cadmium Chromium Copper Gold Lead Mercury Nickel Silver Thallium Uranium

Organic compounds Carbon tetrachloride and other chlorinated hydrocarbons Chloroform Ethylene glycol Herbicides Pesticides Solvents

Miscellaneous nontherapeutic agents Bee venom Diphosphonate Calcium antagonists Gallium nitrate Grapes or raisins Illicit drugs Lilies Mushrooms Radiocontrast agents Snake venom Sodium fluoride Superphosphate fertilizer Vitamin D‐containing rodenticides, “Jerky treats”

Infectious Bacterial/fungal Pyelonephritis Leptospirosis Borreliosis Feline infectious peritonitis Babesiosis Leishmaniasis Bacterial endocarditis

Immune‐mediated inflammatory Acute glomerulonephritis Systemic lupus erythematosus Renal transplant rejection Vasculitis Systemic inflammatory response syndrome Sepsis Disseminated intravascular coagulation

Obstructive Ureteral obstruction Urethral obstruction

Nonnephrotoxic insults Decreased cardiac output/ ischemia Volume depletion Congestive heart failure Arrhythmias Cardiac arrest Cardiac tamponade Fluid overload Deep anesthesia/extensive surgery Renal vessel thrombosis Hyperviscosity – polycythemia Hepatorenal syndrome

Miscellaneous Lymphoma Blood transfusion reactions Heatstroke/hyperthermia Malignant hypertension Neoplasia Hypercalcemia

120  Acute Kidney Injury

existed in human medicine, but has largely been overcome by the development of standardized classification schemes. The most widely accepted schemes are the Risk Injury Failure End Stage Kidney Disease (RIFLE) scheme, the Acute Kidney Injury Network (AKIN), and the Kidney Disease: Improving Global Outcomes (KDIGO) Clinical Practice Guideline for Acute Kidney Injury. Both sets of criteria appear to perform equally well when both sensitivity for detection of AKI and ­predictive ability of adverse outcomes are evaluated and, therefore, these schemes have become accepted within the human nephrology community as the standard means of defining AKI for epidemiologic ­ characterization. There is considerable difficulty, however, with application of these criteria to veterinary AKI because both RIFLE and AKIN schemes primarily rely on relative changes in serum or plasma creatinine concentrations. In contrast to human medicine, in which most cases of AKI are hospital acquired and as such, relative changes in serum or plasma creatinine concentrations are more easily measured, most cases of veterinary AKI currently recognized are community acquired, and a baseline serum or plasma creatinine concentration is not available. Additional criteria for identification and staging of AKI exist within these schemes, but these criteria also have limitations (e.g., lack of baseline value for GFR and practical considerations of indwelling urinary collection system for monitoring of urine output). Nonetheless, recently published retrospective data collected from hospitalized dogs, utilizing a scheme similar to AKIN (Table 120.2), suggest that an association between small changes in plasma creatinine concentration (≥0.3 mg/dL) and mortality may exist. Due to the limitations associated with defining and staging AKI based on relative changes in serum or plasma creatinine, Cowgill recently proposed a veterinary staging scheme based primarily on absolute serum or plasma creatinine concentrations (Table 120.3). This proposed scheme is being validated in a prospective study that is currently under way. Table 120.2  Veterinary Acute Kidney Injury (VAKI) staging scheme for dogs VAKI stage

Criteria

Stage 0

Creatinine increase 4.0 mg/dL

Table 120.3  International Renal Interest Society grading scheme for acute kidney injury (AKI) (dogs and cats) Grade

Creatinine Clinical description

Grade I

250) of potentially pathogenic serovars, this limitation may lead to false‐negative test results. Titers may also be negative within the first 7–10 days of illness; a fourfold rise after 2–4 weeks is used to confirm exposure when initial titers are negative. A single titer of 1:800 or greater, with appropriate clinical signs and in the absence of recent vaccination, is also suggestive of Leptospira spp. exposure. However, there is a high degree of discordance in interpretation of leptospirosis among different commercial laboratories, potentially affecting interpretation of borderline results. A strong clinical suspicion for leptospirosis must be present with titers in excess of 1:800 for serovar Autumnalis, as titers often increase parallel to vaccinal serovars and with other diseases. While the microscopic agglutination test remains the gold standard for leptospira serology, new antibody detection kits are now commercially available that allow more rapid and inexpensive detection of circulating antibodies. These newer assays do not disinguish serovars. Polymerase chain reaction assays for both blood and urine have been developed for rapid, early diagnosis in dogs, but data on their clinical utility are lacking. Serologic tests for other infectious diseases known to cause AKI, such as Rocky Mountain spotted fever (Rickettsia rickettsii), Ehrlichia canis, Lyme disease (Borrelia burgdorferi), Babesia spp., or Leishmania spp., may be useful in certain areas or when there are other consistent clinical or pathologic signs, although a positive titer does not prove causality of AKI.

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Cytology of tissue acquired by fine needle aspirates has limited utility in cases of AKI, but can aid in detection of an infiltrative etiology. Cytology for the diagnosis of renal lymphosarcoma may produce false‐negative results. Therefore, for cases in which lymphosarcoma is suspected, histopathologic assessment of tissue may be necessary to rule out this etiology. Diagnosis of amyloidosis and feline infectious peritonitis requires special cytologic techniques (e.g., Congo red staining or coronavirus immunocytochemistry, respectively), and these diagnostic techniques have not been rigorously assessed. The risk of bleeding secondary to fine needle aspiration of the kidneys is low but possible, especially when platelet dysfunction is present. Histopathologic samples can be obtained by percutaneous, ultrasonographically guided needle biopsy, laparoscopy, or surgical wedge biopsy. Histopathology may confirm a suspected etiology (e.g., ethylene glycol intoxication, renal lymphosarcoma) or it may disclose nonspecific findings. When AKI cannot be clinically distinguished from end‐stage chronic kidney disease, histopathology

(particularly Masson’s trichrome stain) can aid in assessment of the severity of fibrosis and provide insight into the potential for renal recovery. The risk of significant hemorrhage secondary to renal biopsy is high when ­uremia is severe and platelet dysfunction is present. Ethylene glycol intoxication is an emergency situation requiring immediate, specific therapy, which makes accurate and timely diagnosis crucial. Commercially available in‐house test kits are available.

­Therapy Treatment of AKI is primarily aimed at addressing the underlying cause (if it can be identified and treated) and supportive measures to minimize the clinical sequelae of uremia. This section provides treatment recommendations for cases of severe AKI, in which severe uremic manifestations and abnormalities of acid–base, electrolyte, and fluid balance dominate the clinical picture. Specific doses for many of the drugs discussed are listed

Table 120.4  Indications, doses, adverse effects, and comments for drugs frequently used in cases of acute kidney injury Drug

Indication

Dose

Adverse effects

Furosemide

Fluid overload, oliguria/anuria, hyperkalemia

Ototoxicity; volume depletion 2–5 mg/kg IV bolus, may be repeated up 3–5 times; 0.5–1 mg/ (unlikely if patient is monitored) kg/h CRI if urine production increased following bolus

Regular insulin

Hyperkalemia

0.5 units/kg IV or IM, may be repeated q4–6h provided hypoglycemia is avoided

Hypoglycemia

Hypokalemic effect modest and transient; IV dextrose must be administered concurrent with and following insulin administration

Dextrose

Hyperkalemia; avoidance of hypoglycemia following insulin administration

IV bolus of 2 g/unit of insulin administered; bolus followed by CRI (dextrose concentration and administration rate are dependent on serial blood glucose concentrations, patient’s fluid status, and accessibility of central line)

Hyperglycemia, hyperosmolarity, hyponatremia, phlebitis with high dextrose concentrations

Dextrose should be diluted to avoid phlebitis; frequent changes in dextrose CRI often necessary based on serial blood glucose measurements

Calcium gluconate (10%)

Hyperkalemia; symptomatic hypocalcemia

0.5–1.5 mL/kg of 10% solution or 50–150 mg/kg IV slowly, to effect, while monitoring ECG, may be repeated

ECG should be monitored during Worsening bradycardia and ECG changes; hypercalcemia; administration; will not affect extracellular potassium soft tissue mineralization concentration; effective in rapidly normalizing ECG, but results transient; administration of large volumes may contribute to soft tissue mineralization

Sodium bicarbonate

Severe acidemia 1/4 to 1/3 of the base deficit over 30–60 min followed by an additional 1/4 over the next 4–6 hours; additional dosing based on serial blood gas analyses

Paradoxic central nervous system acidosis; hypernatremia; fluid overload; hypochloremia; may cause or exacerbate hypokalemia if patient is polyuric; may exacerbate hypocalcemia

CRI, continuous rate infusion; ECG, electrocardiogram; IM, intramuscular; IV, intravenous.

Comments

Results are frequently not satisfactory in cases of severe AKI, but adverse effects minimal so use in anuric AKI

Requires close monitoring of blood gases and electrolytes for effective treatment and avoidance of adverse effects

120  Acute Kidney Injury

in Table 120.4. While renal replacement therapy (dialysis) is referenced, it is discussed in greater detail in Chapter 126. Fluid Therapy To ensure adequate tissue perfusion, extracellular fluid deficits should be corrected with a balanced polyionic solution. Ultimately, the type of fluid administered must be guided by monitoring of serum or plasma concentration of electrolytes because the degree of solute and free water balance varies widely in patients with AKI. Colloidal support may also be considered to reduce the total amount of fluid administered, if oliguria or anuria is suspected, although no benefit over crystalloid therapy has been documented in human or veterinary medicine. Goal‐directed therapy to restore surrogate markers of perfusion (e.g., blood pressure, venous lactate concentration, venous oxygen saturation) should be employed with endpoints set to be reached within 24 hours. If oliguria or anuria persists despite achievement of normal surrogate markers of perfusion, additional fluid administration is more likely to result in fluid overload than urine production. Avoidance of fluid overload (typically defined as fluid accumulation >10% of baseline body weight) is essential, as there is ample evidence documenting the association between fluid overload and worse clinical outcomes. Maintenance fluid administration (both volume and composition) should be guided by the volume and composition of urine produced, as well as ongoing sensible losses (vomitus, diarrhea, and yield from gastric suction) and insensible loss (respiration, formed stool). Urine volume can be determined by a variety of methods, including: ●●

●● ●● ●●

●●

indwelling urinary catheter and closed collection system collection of naturally voided urine metabolic cage weighing cage bedding and litter pans (1 mL of urine = 1 g) using body weight prior to and immediately following urination.

Urine production can be categorized as anuria (none to negligible amount), oliguria (2 mL/kg/h). Insensible losses can be estimated between 12 and 29 mL/kg/day and are dependent on a variety of factors, such as species, patient activity, and body ­temperature. Careful attention must be given to serial changes in the patient’s body weight, as peracute fluctuations in weight are most likely due to changes in fluid balance rather than changes in lean muscle or fat content. Once the patient’s fluid deficit has been corrected, care must be taken to maintain a neutral fluid balance, as well as normal surrogate markers of perfusion. Consideration

of the fluid load with enteral and parenteral nutrition and medication, as well as intravenous catheter flushes, is essential to avoidance of fluid overload, and the fluid volume administered with these treatments should be incorporated into the fluid plan. High‐maintenance fluid rates have been historically advocated for cases of AKI based on the rationale that high‐volume fluid administration beyond that which is necessary to restore normal volume status will improve GFR. However, there is no evidence supporting this claim and, in the author’s experience, this practice is often futile in restoring GFR and frequently results in fluid overload. Maintenance fluid therapy for an anuric, euhydrated patient should consist of replacement of insensible losses only. Frequently, this fluid requirement is achieved in excess by administration of medications, nutrition, and catheter flushes alone and the administration of these treatments may promote fluid overload. If the patient is diagnosed with fluid overload, all fluid therapy should be withheld. Fluid overload with concurrent oliguria or anuria is a clear indication for dialysis. Monitoring fluid status is an ongoing process that must be repeated frequently. Efforts should be made to adhere to objective monitoring parameters (e.g., body weight, venous lactate concentration, urine production) of fluid status because subjective parameters, (e.g., skin turgor, saliva production) are inaccurate and often affected by variables other than hydration status. Body weight should be measured at least twice daily to assess for trends in fluid accumulation or deficit. Central venous pressure measurement has traditionally been recommended as a surrogate marker of cardiac preload, and thus fluid status. However, a thorough understanding of the limitations of this technique is necessary for appropriate interpretation, as the correlation between central venous pressure and clinical manifestations of fluid overload is increasingly recognized as flawed. Diuretics The use of diuretics in the treatment of AKI is a controversial topic in both human and veterinary medicine. Many of the benefits of the most commonly used diuretics in veterinary AKI, furosemide and mannitol, have only been theorized or demonstrated in experimental models of AKI. In fact, there is little or no clinical evidence in human or veterinary medicine that diuretics improve outcome in established AKI. It has been postulated that the ability to respond to diuretics is a marker of less severe renal injury associated with a better prognosis. However, an increase in urine output after diuretic administration does not necessarily coincide with an increase in uremic solute excretion and, therefore, does not preclude the need for dialysis if severe uremia or acid–base and electrolyte abnormalities persist.

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In veterinary medicine, because dialysis is not readily available, diuretic administration plays a large role in volume management. Conversion from an oliguric or ­anuric state to normal urine production or polyuria may enhance the clinician’s ability to prevent or manage fluid overload and thus allow administration of necessary parenteral medications and nutrition that would otherwise contribute to fluid overload. No class of diuretics has been proven to be superior to another. However, the use of loop diuretics predominates in both human and veterinary AKI due to the relatively high efficacy and safety margin of these drugs, compared to the osmotic diuretics. For this reason, the author does not recommend the use of mannitol as a diuretic for AKI. Acid–Base Balance Metabolic acidosis is a frequent complication in AKI of varying severities, and is due to the damaged nephron’s inability to excrete hydrogen ions and reabsorb bicarbonate ions, as well as lactic acidosis secondary to compromised tissue perfusion (i.e., either volume deficit or excess). Once perfusion has been restored, provision of supplemental alkali, usually in the form of parenteral sodium bicarbonate, should be considered if severe acidemia (pH 1.0 is associated with negative patient outcomes and when the UPC is reduced to 0.5. However, standard therapy rarely leads to complete resolution of the renal lesions. Inhibition of RAAS

Hemodynamic forces influence the transglomerular movement of proteins. The RAAS can be therapeuti­cally targeted to alter renal hemodynamics, leading to a reduction in proteinuria. Decreased efferent glomerular arteriolar resistance leads to decreased glomerular transcapillary hydraulic pressure, which ideally leads to a reduction in proteinuria. Other proposed mechanisms include reduced loss of glomerular heparan sulfate, decreased size of the glomerular capillary endot­helial pores, improved lipoprotein metabolism, slowed ­glomerular mesangial growth and proliferation, and i­nhibition of bradykinin degradation.

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Box 121.1  Management of glomerular disease in dogs Inhibition of renin‐angiotensin‐aldosterone system Angiotensin‐converting enzyme inhibitors (e.g., enalapril, benazapril) Angiotensin receptor blockers (e.g., telmisartan) Aldosterone receptor blockers (e.g., spironolactone) Modified dietary intake Increased n‐3 polyunsaturated fatty acids Modified protein content Reduced sodium chloride Antithrombotic therapy Aspirin Clopidogrel Antihypertensive therapy Angiotensin‐converting enzyme inhibitors (e.g., enalapril, benazapril) Angiotensin receptor blockers (e.g., telmisartan) Calcium channel blocker (i.e., amlodipine) Manage body fluid volume Careful fluid therapy when fluid deficit present Diuretics for fluid excesses that are contributing to respiratory distress Immunosuppressive therapy for immune complex‐ mediated glomerulonephritis

Angiotensin‐converting enzyme inhibitors (ACEi; e.g., enalapril, benazapril), angiotensin receptor blockers (ARB; e.g., telmisartan), and aldosterone receptor blockers (e.g., spironolactone) are the agents that target RAAS. For most dogs with glomerular‐range proteinuria, an  ACEi is the initial agent used. Typically, an ACEi is given once daily initially, but more than half of the dogs will eventually need twice‐daily administration and perhaps additional dosage escalations. Although worsening azotemia is a concern regarding the use of ACEi, severe worsening (i.e., >30% increase from baseline) due to ACEi administration alone seems uncommon and dogs that are dehydrated may be at highest risk. Less is known about the use of an ARB in dogs with glomerular disease although they have been extensively studied in people. The ARB that has historically been used in dogs with glomerular disease is losartan, although it has not always been effective. Recently, telmisartan has been shown to be very effective in some dogs and may prove useful as the initial line of therapy. Telmisartan has a more profound effect on lowering blood pressure than ACEis and should be used carefully in dogs that have low basal blood pressures (e.g., systolic 30% increase in serum creatinine), a critical increase in serum potassium or the unlikely development of hypotension. Changes in the magnitude of urine protein are most accurately measured by assessing trends in the UPC over time. Day‐to‐day variations in the UPC occur in most dogs with glomerular proteinuria, with greater variation occurring in dogs with UPC >4. Consideration should be given to either averaging 2–3 serial UPC or measuring a UPC in urine that has been pooled from 2–3 collections. The target for reduction is a UPC of 6 mEq/L should be monitored closely and therapy should be modified when serum potassium concentrations are >6.5 mEq/L. Before modifying treatment, pseudohyperkalemia, due to the high potassium content of some blood cells which may occur in dogs with glomerular disease, should be eliminated as a cause by measuring the potassium concentration in lithium heparin plasma. True hyperkalemia can be managed by reducing the ACEi or ARB drug dosage, discontinuing spironolactone ­administration, or by feeding

121  Glomerular Disease

diets that are reduced in potassium or administering an intestinal potassium binder (e.g., kayexelate). If the target reduction in UPC is not achieved and the dog does not have the before mentioned adverse effects, the dosages may be increased every 4–6 weeks. Modified Dietary Intake

Modification of dietary intake has a central role in the management of all dogs with kidney disease and has been associated with delaying spontaneous progression of kidney disease and reducing the magnitude of proteinuria. It is well known that fat content of a diet can affect renal outcome. Most commercially available renal diets are modified with respect to polyunsaturated fatty acids (PUFA) content with the goal of slowing progression and reducing proteinuria. The consensus recommendation is that dogs with glomerular disease should be fed a diet with a reduced n‐6:n‐3 PUFA ratio of approximately 5:1. It is unknown if there is any additional benefit of supplementing n‐3 PUFA when this ratio is already being provided via the diet. If supplementation is chosen, care should also be taken to provide adequate antioxidants (e.g., vitamin E). Dogs with glomerular disease should be fed modified protein diets because this has been shown to reduce proteinuria and slow progression. Lastly, the sodium chloride content should be reduced in diets fed to dogs with glomerular disease. Salt restriction may enhance the beneficial effects of agents used to inhibit RAAS and help reduce fluid retention. Antithrombotic Therapy

Hypercoagulability is a complication of protein‐losing glomerular diseases and thromboembolism can occur in as many as 25% of dogs with glomerular disease. The pathogenesis of hypercoagulability in glomerular disease is complex and incompletely understood. Urinary loss of antithrombin has long been given as the explanation but in reality, the pathogenesis is more complex than this  and probably includes vascular stasis as well as  increased platelet aggregation, plasma procoagulant factors, and fibrinogen concentrations. Unfortunately, serum albumin concentrations, antithrombin activity, and UPC cannot be used in isolation to predict hypercoagulability in individual patients. It follows then that it remains unclear when to implement thromboprophylaxis. Consensus recommendations call for the daily administration of low‐dose aspirin (1–5 mg/kg/day) in dogs with protein‐losing glomerular diseases. Clopidogrel may be an effective alternative. Antihypertensive Therapy

All dogs with glomerular disease should undergo repeat evaluation for systemic hypertension. This evaluation should not only include measurement of systemic blood

pressure but also assessing the patient for target organ damage, evaluating for conditions that may contribute to hypertension and determining if any concurrent conditions that may complicate antihypertensive therapy are present. Personal preference and experience should dictate how the blood pressure is measured; however, the substaging by arterial blood pressure system that has been proposed by the International Renal Interest Society (IRIS) should be used to guide therapy (Table 121.2). Antihypertensive therapy should be initiated when the blood pressure exceeds 160 mmHg systolic or 100 mmHg diastolic (i.e., AP2 or above). The goal of therapy is to reduce the blood pressure to less than 150 mmHg systolic or 96 mmHg diastolic (i.e., AP0). Lowering the blood pressure is not an emergency unless there is severe ocular or central nervous system target organ damage. In most cases, blood pressure reduction can be achieved gradually over a period of several weeks. Effective treatment of hypertension should lead to a concomitant reduction in the magnitude of proteinuria in dogs with glomerular disease. Feeding a salt‐restricted diet alone will not lower the blood pressure into the target range but it may enhance the antihypertensive effects of the inhibitors of RAAS, generally ACEi. However, ACEi are generally only associated with a 10–15% reduction in blood pressure. This reduction may be all that is needed for dogs that are only moderately hypertensive (i.e., AP2). Dogs that are severely hypertensive (i.e., AP3) should have a calcium channel blocker (i.e., amlodipine) added to their RAAS inhibition therapy. Alternatively, telmisartan can be used as a single agent to manage hypertension and proteinuria. The consensus recommendation is that dogs with IRIS chronic kidney disease stages 1 or 2 should have blood pressure evaluated 3–14 days after any change in antihypertensive therapy whereas those in stages 3 or 4 should Table 121.2  International Renal Interest Society substaging by arterial blood pressure Systolic blood pressure

Substage

180 mmHg

Severely hypertensive, Severe Risk

No evidence of end‐organ damage

nc (no complications)

Evidence of end‐organ damage

c (complications)

Blood pressure not measured

RND (risks not determined)

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Section 10  Renal and Genitourinary Disease

be evaluated 3–5 days after making any change. Dogs that have hypertensive emergencies should be evaluated daily. The purpose of these evaluations is not only to assess effectiveness of the antihypertensive therapy but also to evaluate for exacerbation of azotemia. Dosages should be reduced if the serum creatinine has increased more than 30% from baseline or if the blood pressure is reduced to less than 120–mmHg systolic or 60–mmHg diastolic. Once target blood pressure is achieved, the dogs should be evaluated at 1–4‐month intervals, depending upon the stability and severity of the renal disease. Body Fluid Volume

Dogs with glomerular disease may have fluid excesses, deficits or maldistribution, which may need to be corrected if the dog has decompensated or is being prepared for anesthesia. Correcting body fluid imbalances in dogs that are hypoalbuminemic from glomerular disease is a very difficult therapeutic challenge. Fluid therapy can exacerbate edema and hypertension and diuretics can exacerbate azotemia or uremia. As such, those who have experience and expertise in critical care are the best people to deliver these therapies to dogs with glomerular disease. Fluid therapy should be given only when needed to help control clinical signs or provide support to those dogs that have inadequate fluid intake. Both colloids and crystalloid can be used. However, the decision to use colloids should not be based on the presence of hypoalbuminemia or decreased colloidal oncotic pressure alone. Colloids should be considered when crystalloids have failed to bring about the desired result. The fluid status of affected dogs should be assessed before and during fluid therapy on the basis of serial body weight and other physical examination findings (e.g., skin turgor, appearance of mucous membranes, capillary refill time, pulse rate and quality, arterial blood pressure). When the serum albumin decreases below 2 g/dL in people, the plasma oncotic pressure is sufficiently reduced to allow for transudation of fluid from the vascular compartment into the interstitial space. Dogs may be more resistant to the formation of edema, which does not generally occur until the serum albumin concentration is below 1.5 g/dL. Plasma volume may be reduced at this point, making the use of diuretics in the management of edema relatively ineffective and also dangerous because of the increased risk of acute kidney injury and vascular stasis leading to thromboembolism. Diuretics should be avoided unless needed to control respiratory distress. When needed, furosemide is the drug of choice for pulmonary edema but spironolactone may be best for pleural or abdominal effusion. Provision of adequate exercise also may help reduce the formation of edema or ascites.

Immunosuppressive Therapy Immunosuppressive therapy should be considered in dogs that have severe, persistent or progressive glomerular disease and ICGN documented via appropriate evaluation of a renal biopsy specimen. However, there are many practical and medical reasons why a biopsy might not be performed. In addition, financial limitations may prevent biopsy. The end‐result is that sometimes veterinarians must decide about the use of immunosuppressive therapy in dogs with glomerular disease without a pathologic diagnosis. In these situations, immunosuppressive therapy should not be administered if there is any doubt that the proteinuria is of glomerular origin, administration of the specific drug is medically contraindicated, or there is a high index of suspicion that the dog has a nonimmune‐mediated familial disease or amyloidosis. Immunosuppressive drugs should be considered without a pathologic diagnosis when standard therapy has been implemented but the azotemia is progressive or the serum creatinine is >3.0 mg/dL or serum albumin is 25% of baseline (partial response). A meaningful increase in serum albumin is defined as a sustained increase to >2.5 mg/dL (complete response) or either to 2.0–2.5 mg/dL or by >50% of baseline (partial response). Secondary goals include improved blood pressure regulation, resolution of edema, and stabilization of body weight. If there are no unacceptable adverse drug effects, treatment should be continued for 8–12 weeks before changing the regimen or discontinuing treatment. Those demonstrating a partial or complete response should have therapy continued for at least 12–16 weeks. If partial or complete responses are not evident by the end of 8–12 weeks, the immunosuppressive protocol should be changed or discontinued. If there is no response by 3–4 months, all immunosuppressive

therapy should be discontinued following appropriate drug tapering.

­Prognosis The prognosis for dogs and cats with glomerular disease is variable and probably based on a combination of factors. The prognosis is expected to differ with the various diseases. Although progressive disease can be expected to occur in a large percentage of animals with glomerular disease, spontaneous remission and response to specific therapy can also be expected. Furthermore, disease progression can be slow enough for the animals to lead relatively normal lives, especially when the diagnosis is established early in the disease process. In humans, azotemia, severe proteinuria, systemic hypertension, and marked tubulointerstitial lesions at presentation are the most significant predictors of an unfavorable outcome in most forms of glomerular disease. In dogs, the presence of nephrotic syndrome or azotemia is a negative prognostic indicator. Median survival time for dogs with nephrotic syndrome was only 12.5 days versus 104.5 days for dogs without nephrotic syndrome. When only nonazotemic dogs were considered, (i.e., serum creatinine 8 mEq/L

Terbutaline

0.01 mg/kg

IV bolus

If Ca gluconate given Potassium >8 mEq/L

Sodium bicarb

1 mEq/kg

Admin over 3–5 min

Potassium >10 mEq/L

c, cat; d, dog; ECG, electrocardiogram.

122  Obstructive Uropathy

polytetrafluoroethylene) can be used to initially relieve the obstruction. Given the very rigid nature of polypro­ pylene catheters, care should be taken not to use too much force in advancing. For comfort, as well as associ­ ated inflammation and irritation, these catheters should also not be left in place. With regard to optimal catheter size, there is some evidence to suggest that use of a 3.5 Fr urinary catheter may be associated with less risk of immediate reobstruction compared to 5 Fr. However, another study failed to show this association. When passing the catheter, aggressive flushing (rather than force) should be utilized to dilate the urethra and retropulse/break down any physical component of the obstruction. For the flush solution, adding sterile lubri­ cant to sterile saline and mixing across a three‐way stop­ cock (in a ratio of 10:1) may help to decrease urethral injury by allowing lubricant to be deposited throughout catheter placement (rather than just lubricating the end of the catheter). Another helpful technique is to pull the prepuce dorsal and caudally after the catheter is intro­ duced into the penile urethra (Figure  122.5). Elevating the natural downward angle of the urethra as it passes out of the pelvic canal may facilitate catheter placement. Once the initial catheter is in place, the urinary bladder can be emptied and flushed. A sterile closed collection system should be connected to allow for urine produc­ tion to be quantified, and decrease risk of ascending infection. (a)

For catheterization and deobstruction in dogs, general anesthesia can be especially important to ensure optimal urethral relaxation. The prepuce or vulva should be cleaned and flushed to decrease risk of contamination. A red rubber or Foley catheter is typically used, though in some circumstances a more rigid catheter may be needed (with potential increased risk of urethral trauma). Similar to the process described for cats, hydropulsion with lubricated saline should be the primary means of retro­ pulsing any physical obstruction. In male dogs, it is help­ ful to pinch the urethral orifice to prevent antegrade movement of flush solution. An additional useful trick (in either male or female dogs) is to have an assistant rec­ tally apply pressure to the pelvic urethral during hydro­ pulsion. This allows a build‐up of pressure which, when released, can help dislodge a luminal obstruction. If urethral catheterization fails due to urethral trauma, rupture, or persistent partial obstruction, antegrade ure­ thral access can provide a means to deobstruct the patient and facilitate catheterization (Figure 122.6). Briefly, percutaneous access into the bladder is achieved with an 18 gauge over‐the‐needle catheter by palpation or ultrasound guidance. Through the catheter, a hydro­ philic wire is directed toward the bladder trigone and out the urethra. As the tear or trauma is usually created from a retrograde direction, the wire finds the true urethral lumen easily and often can be directly advanced out the penile urethra. Once wire access is achieved in this man­ ner, the open‐ended urinary catheter is advanced over the wire into the bladder lumen. The access catheter and wire are then removed and the urethral catheter sutured in placed and cared for routinely. Ideally, this procedure is performed under fluoroscopic guidance to visualize the path of the wire and appropriate positioning of the catheter. Postobstructive Care

(b)

Figure 122.5  Once the catheter is seeded in the distal urethral, the prepuce is pulled dorsal and caudal to straighten the urethra and facilitate passage.

One major facet of postobstructive care is careful moni­ toring of urine output and maintaining fluid balance. Some patients may experience a postobstructive diuresis which can lead to significant quantities of urine produc­ tion. Proposed mechanisms for this diuresis include accumulation of osmotically active substances in the blood, pressure necrosis, medullary washout and/or antidiuretic hormone resistance. Likely owing to these mechanisms, it has been demonstrated that up to 50% of cats may have increased urine production after deob­ struction. However, in some circumstances it is unclear whether the initial rates of fluids administered may have also been contributing to increased urine production. Similar data are not available for dogs. Given the potential to produce significant quantities of urine (>5–10 mL/kg/h), it is very important to keep up with urinary losses to prevent dehydration and hypov­

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(a)

(d)

(b)

(e)

(c)

(f)

Figure 122.6  Radiographic images of antegrade urethral catheterization for urethral tear in a cat. (a) The initial retrograde urethrogram showing severe contrast extravasation throughout the pelvic canal consistent with urethral rupture during attempted urethral catheterization. (b) Percutaneous access to the bladder has been obtained with an 18‐gauge over‐the‐needle catheter and iodinated contrast has been delivered to highlight the urinary bladder. (c) A hydrophilic guidewire is then advanced through the catheter and into the urinary bladder. (d) The wire is manipulated under fluoroscopic guidance and advanced antegrade out the urethra. As the tear was made during retrograde catheterization, the soft wire will follow the normal urethral lumen and pass out the penile urethra. (e) Once the wire is exteriorized, an open-ended urinary catheter can be advanced over the wire and will follow the true urethral lumen. Once the urinary catheter is within the bladder, the wire and bladder catheter are removed. (f ) After seven days of in‐dwelling urethral catheterization, a repeat urethrogram shows the urethral tear has healed and the urinary catheter is removed.

olemia. This can be achieved by at least matching the fluid rate to hourly urine production (even though the resultant rate could be very high). In the postobstructive period there is also the potential concern for inadequate urine production (7 but 6 years

Struvite Any breed Increased risk in miniature schnauzer, bichon frise, shih tzu, Lhasa apso, Yorkshire terrier, miniature poodle (and other small‐breed dogs). The predisposition for smaller breeds may be related to their lower urine volume, fewer numbers of micturitions, and likely increased urine mineral concentrations compared with larger breeds

No age predilection (middle‐aged but generally younger than those with oxalate calculi). Calculi that occur in dogs younger than 1 year are frequently composed of struvite

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Table 123.2  Urolith composition for feline and canine samples submitted to the Canadian Veterinary Urolith Centre (Guelph, Ontario, Canada; 2009–2012)

Etiopathogenesis Hypercalciuria and Hyperoxaluria

Major composition

Canine n (%)

Feline n (%)

Total n (%)

Ammonium urate

619 (2.9)

257 (4.7)

876 (3.3)

Brushite

60 (0.3)

2 (0.04)

62 (0.2)

Calcium oxalate

10 043 (47.4)

2778 (50.7)

12 821 (48.1)

Calcium phos apatite

221 (1.0)

69 (1.3)

290 (1.1)

Calcium phos carbonate

240 (1.1)

11 (0.2)

251 (0.9)

Cystine

134 (0.6)

6 (0.1)

140 (0.5)

Silica

181 (0.9)

7 (0.1)

188 (0.7)

Sodium urate

21 (0.1)

1 (0.01)

22 (0.08)

Struvite

7226 (34.1)

2238 (40.9)

9464 (35.5)

Uric acid

5 (0.02)

0

5 (0.02)

Xanthine

5 (0.02)

5 (0.09)

10 (0.04)

X‐oxalate/Ca phosphate

237 (1.1)

28 (0.5)

265 (1.0)

Excessive urinary excretion of calcium (hypercalciuria) or oxalate (hyperoxaluria) results in increased risk of cal­ cium oxalate urolith formation. Lulich et al. reported that 6/6 miniature schnauzers with calcium oxalate urolithia­ sis had hypercalciuria. Similarly, urinary concentration of oxalate is an important factor in calcium oxalate crystal­ lization and urolithiasis. This is because increases in urine oxalate concentration have a greater effect on urinary saturation of calcium oxalate than equivalent increases in urinary calcium concentration on a molar basis. Urinary oxalate is derived mainly from endogenous production in the liver (metabolism of ascorbic acid, gly­ oxylate, and glycine) with lesser amounts from dietary intake of oxalate. Enteric (large intestine) colonization of Oxalobacter formigenes, an anaerobe which exclusively relies on oxalate metabolism for energy, is correlated with absence of hyperoxaluria or calcium oxalate (CaOx) urolithiasis in humans and laboratory animals. It has recently been demonstrated that the absence of enteric colonization of O. formigenes is a risk factor for CaOx urolithiasis in dogs.

X‐oxalate/silica

63 (0.3)

1 (0.02)

64 (0.2)

X‐oxalate/struvite

39 (0.2)

13 (0.2)

52 (0.2)

Water Type

X‐struvite/Ca phosphate

1878 (8.9)

21 (0.4)

1899 (7.1)

X‐struvite/urate

215 (1.0)

41 (0.7)

256 (1.0)

Total

21 187

5478

26 665

Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

In humans, the mineral content of water may play a role in calcium oxalate urolith formation. It is also thought that increased sodium content can increase water intake and offset its calciuretic effect. Increased water sodium, calcium, or the absence of trace minerals such as zinc, which chelates calcium, may increase the risk of calcium oxalate urolithiasis. However, in cats, the source of water (bottle, municipal, etc.) did not appear to affect the risk of calcium oxalate urolithiasis. A similar study has not been conducted in dogs. Modifiers of Calcium Oxalate Crystal Formation

In humans, urine normally contains several natural inhibitors of urolith formation and growth. Inhibitors can be either organic or inorganic substances in urine that reduce crystal formation, aggregation, or growth. A defect or deficiency in these inhibitors may contribute to urolith formation. Inhibitors of calcium oxalate include citrate, magnesium, pyrophosphate, glycosaminogly­ cans, nephrocalcin, and Tamm–Horsfall mucoprotein. Although little is known about these inhibitors in dogs, it is suspected they may play a similar role as in humans. Concurrent Medical Conditions Figure 123.2  Feline oxalate stones from a 16‐year‐old female spayed domestic short‐hair. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

Hyperadrenocorticism is associated with calcium oxalate urolithiasis in humans and dogs. In humans, glucocorti­ coid excess leads to hypercalciuria secondary to increased mobilization of calcium from bone. Glucocorticoid‐

123  Urolithiasis in Small Animals

induced decreases in tubular resorption of calcium, chronic metabolic acidosis, and decrease in urinary excre­ tion of citrate may also occur. Similar glucocorticoid‐ associated mechanisms are believed to occur in dogs. A recent study in humans demonstrated an association between calcium oxalate stones and hypertriglyceri­ demia. The predisposition of miniature schnauzers for idiopathic hyperlipidemia may one day be found to explain why this breed is at high risk of calcium oxalate stones. Struvite Urolithiasis Epidemiology

Struvite uroliths are typically composed of 100% magne­ sium ammonium phosphate hexahydrate. Struvite stones may also contain varying amounts of calcium phosphate (termed calcium apatite) or calcium carbonate phos­ phate (termed carbonate apatite). These uroliths are commonly referred to as “infection stones” or “urease stones” because, in dogs, they are primarily due to the presence of urease‐producing bacteria. Struvite stones are often multifaceted or pyramidal in appearance, espe­ cially when multiple urocystoliths are present, because the adjacent surfaces are smooth and flattened. If soli­ tary, struvite may have sharp spiculated projections (Figure  123.3). Struvite uroliths represented 35.5% (43.6% including mixed stones) of all canine and feline uroliths submitted to CVUC from 2009 to 2012 (see Table 123.2). More recently, struvite urolith submissions have increased and have exceeded the calcium oxalate

Figure 123.3  Bladder struvite uroliths retrieved from a 10‐year‐ old female spayed golden retriever. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

urolith submissions for small animals (2018 Minnesota Urolith Center [MUC] Global Data generated by Minnesota Urolith Center, February 2019). Struvites are most commonly located in the lower uri­ nary tract (LUT) (95%). Risk factors appear related to sex and associated higher bacterial infection risk, as they are more frequent in female dogs (71–85% of struvite sub­ mitted to urolith centers) than male dogs (15–29%). Certain breeds are predisposed to struvite uroliths (see Table 123.1). Etiopathogenesis

The majority of canine struvite uroliths are due to urinary tract infection, most commonly related to bacterial infec­ tion with urease‐producing organisms. However, sterile struvite urolithiasis has been reported in related cocker spaniels. In dogs, Staphylococcus pseudintermedius is cited as the most common urease‐producing bacterium associated with struvite uroliths. Other less common urease‐producing organisms include Pseudomonas spp., Klebsiella spp., Proteus, and Corynebacterium urealyticum. Urease‐producing mycoplasmas (termed urea­ plasma) such as Ureaplasma urealyticium may also cause struvite urolithiasis. Infection‐based struvite uroliths form when urease cleaves urea to form ammonia and bicarbonate. The available ammonium then combines with magnesium and phosphate, normally present in urine, to form mag­ nesium ammonium phosphate hexahydrate (struvite) crystals. Bicarbonate increases the urine pH (i.e., urine becomes more basic), which then decreases the solubil­ ity of struvite crystals. Ammonium also damages the urothelial glycosaminoglycan layer by acting as an irri­ tant. This allows struvite crystals and bacteria to adhere to the urothelium and may promote production of an organic matrix for the crystal–matrix interaction, lead­ ing to stone formation. Unlike in cats, where sterile stru­ vite urolithiasis may occur solely because of excess mineral intake in the diet, infection with a urease‐pro­ ducing bacterium is associated with struvite urolithiasis in most dogs. Infection‐induced struvite uroliths may contain viable bacteria trapped in the interstices of the stone layers. Bacteria have been cultured from the core of struvite uroliths that have been stored in formalin for years. Dissolution or fragmentation of struvite uroliths may release viable bacteria into the urinary tract and this may cause reinfection unless appropriate antimicrobials and duration of therapy are provided. If the bacteria isolated from the urine and the urolith differ, a change in antimi­ crobial is only necessary if poor clinical response is observed. If a nonurease‐producing bacterium is isolated and subclinical bacteriuria is suspected, an antimicrobial

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is rarely indicated. If a urease‐producing bacterium is isolated, antibiotic therapy should be continued as long as struvite uroliths are present. If therapy duration is insufficient, UTIs will not resolve or a relapse of infec­ tion will likely occur. Purine Urolithiasis Epidemiology

Naturally occurring purine uroliths consist of uric acid (and its various salts) and xanthine. Xanthine and uric acid are successive biodegradation products in the metabolism of purines (Figure  123.4). Purine uroliths may be pure (100%) in composition, or less commonly, may contain varying amounts of other minerals. The term “urate” includes uric acid, ammonium urate, and the other salts of uric acid, mostly sodium acid urate, sodium calcium urate, and ammonium calcium urate. Ammonium urate accounts for 86% of urate uroliths. Urate uroliths are typically small, round, smooth stones that are greenish‐brown in color (Figure  123.5). There are often multiple urate uroliths present. Cross‐ sections of urate uroliths reveal a concentric appearance. Xanthine uroliths are typically similar in size and shape to urate uroliths but are usually yellow to brown (Figure 123.6). Purine uroliths accounted for 6.4% of canine uroliths analyzed at the MUC between 1981 and 2007, making it the third most common type of canine urolith in this region. In Canada, purine uroliths represented 3.44% of canine and feline uroliths submitted to the CVUC between 2009 and 2012 (see Table  123.2). The majority of urate (97%) and xanthine (94%) uroliths that are submitted for Dietary purines

Endogenous purines

Figure 123.6  Xanthine stone retrieved from a 2‐year‐old male neutered Dalmatian. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

Hypoxanthine (Xanthine Oxidase) Xanthine (Xanthine Oxidase) Excreated in the urine of Dalmatians

Allopurinol* (Xanthine oxidase inhibitor)

Uric Acid (Uricase)

Excreated in the urine of normal dogs

Figure 123.5  Urate stones retrieved from a 12‐year‐old male neutered Dalmatian dog. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

Allantoin *Increases urinary excretion of hypoxanthine and xanthine

Figure 123.4  Normal canine purine degradation pathway.

analysis have been removed from the LUT. However, con­ sidering the difficulty in removing uroliths from the upper urinary tract (UUT), nephroliths and ureteroliths may be underrepresented in submission frequency. Risk factors for purine uroliths (and types of these) include species, breed, age, and sex. Ammonium urate is the most common (approx. 95%) naturally occurring purine urolith in cats. Xanthine uroliths are uncommon specimens submitted to the CVUC, having been diag­ nosed in only five cats and five dogs (0.02% of uroliths in cats and 0.09% of uroliths submitted in dogs) from 2009 to 2012 (see Table 123.2). Uroliths composed of hypox­ anthine have not been recognized in cats. Urate stones may be found in dogs of any age but are most frequently retrieved from young to middle‐aged dogs (see Table  123.1). Pure urate uroliths occur more

123  Urolithiasis in Small Animals

often in males than in females. Female dogs may sponta­ neously void urate uroliths without being detected. In dogs, urate uroliths are most commonly retrieved from Dalmatians (61% of ammonium urate and 91% of sodium and calcium urate uroliths). In cats, the typical patient with urate uroliths is a pure‐bred neutered cat, 4–7 years old, with uroliths in the bladder or urethra. The Siamese breed seems to be overrepresented, along with the Egyptian Mau and Birman breeds. Cat breeds that had  lower odds for urate urolith submission than mixed breeds included Abyssinian, American shorthair, Himalayan, Manx, and Persian. There is no sex predispo­ sition in cats. Naturally occurring xanthine uroliths are rare and are frequently retrieved from dogs that have been treated with allopurinol. Etiopathogenesis in Dogs

The purine metabolites, uric acid and xanthine, have much lower solubility than that of allantoin. Consequently, if urine is oversaturated with these substances, uroliths can form. A high dietary intake of purines and purine precursors, along with acidic urine, aids in urolith for­ mation. Infection with urease‐producing bacteria may also promote formation of urate uroliths by increasing ammonium ion concentration from the breakdown of urea (see Figure 123.6). Dalmatian Dogs

Dalmatian dogs are predisposed to the development of urate uroliths because of their reduced transport of uric acid into hepatocytes. This results in decreased conver­ sion of uric acid to allantoin, a subsequent moderate increase in serum uric acid concentration (hyperurice­ mia), and then hyperuricosuria. The inheritance of hyperuricosuria in Dalmatians is via a simple autosomal recessive gene. The missense mutation in the SLC2A9 gene that encodes for a urate transporter is responsible for this defect, and all Dalmatians are homozygous. Despite excretion of relatively large quantities of uric acid, not all Dalmatians form urate uroliths. It is esti­ mated that 25% of male Dalmatians will have urolithiasis and it is suspected that other risk factors play a role in dogs who become urate stone formers. Non‐Dalmatian Dogs

Dogs other than Dalmatians accounted for approxi­ mately 30–60% of urate uroliths submitted for analysis. Similar to Dalmatians, urolith‐forming bulldogs and black Russian terriers have been shown to be homozy­ gous for a mutation in SLC2A9. However, the reasons for development of urate uroliths in these non‐Dalmatian breeds is unknown and does not appear associated with hepatic dysfunction.

Hepatic Dysfunction

Hepatic dysfunction is associated with a reduced ability to convert ammonia to urea and uric acid to allantoin. Therefore, dogs suffering from hepatic dysfunction may develop hyperammonuria and hyperuricuria, which may result in urate urolith formation. The incidence of urate urolithiasis appears to be higher in dogs with por­ tal vascular anomalies than with other causes of hepatic dysfunction. Xanthine

Xanthine is an important mediator in purine metabolism; the drug allopurinol binds rapidly to (and inhibits the action of ) xanthine oxidase, thereby decreasing conver­ sion of hypoxanthine to xanthine and xanthine to uric acid. The result is a reduction of serum and urine concen­ trations of uric acid, with an increase in serum and urine concentrations of xanthine. As such, most xanthine uro­ liths in dogs form secondary to therapy with allopurinol via this mechanism. This is particularly true when a diet high in purines (meat based) is fed to these at‐risk dogs. Naturally occurring xanthinuria has been reported in Cavalier King Charles spaniels and dachshunds, and is thought to reflect an inborn error of xanthine oxidase activity. Etiopathogenesis of Urate Urolithiasis in Cats

The pathophysiology of urate urolith formation in cats is unknown. Ammonium urate uroliths have been recov­ ered from cats with portovascular anomalies. Such con­ genital anomalies can be found in male and female cats and are typically are detected when cats are 99% reabsorption in the proximal renal tubules, these amino acids are lost in the urine; however, only cystine causes a problem. In cystinuric patients, the carrier proteins responsible for reabsorp­ tion are defective. The type of amino acids lost and their quantity vary depending on the case. The low solubility of cystine in acidic urine predisposes to the formation of cystine crystals and uroliths in the urinary tract. Cystinuria in humans and dogs is classified into several distinct types depending on age of onset, severity, sex, inheritance, and mutant gene. In dogs, type I‐a cystinu­ ria is an autosomal recessive disease caused by a SLC3A1 gene mutation common in the Newfoundland, Landseer, and Labrador retriever breeds. A nonsense mutation in exon 2 of the SLC3A1 (amino acid  –  aa‐transport) gene has been identified as a ­molecular basis of the defect but the underlying genetic

123  Urolithiasis in Small Animals

defect(s) in most cystinuric breeds of dogs have yet to be identified. More recently, Labrador retrievers were found to have a different mutation in the same gene, encoding for a renal basic amino acid transporter. DNA‐based genetic tests are available to detect abnormal cystine excretion for some breeds of dogs. Type II cystinuria is an autosomal dominant disease that has been reported in Australian cattle dogs, border collies (type II‐a) and miniature pinscher (type II‐b). In the latter, the mutation affects the SLC7A9 gene and encodes for intramembrane transporter protein. The molecular basis for androgen‐dependent type (type III) cystinuria is unknown. Type III cystinuria regroups several breeds of dogs such as the English bull­ dog, English mastiff, French bulldog. Type III cystinuria is typically detected later in life, is less severe clinically, and appears to involve only males. Castration seems to lower cystine excretion, therefore the mode of inherit­ ance is unclear. No gene mutation has been identified as the cause of type III cystinuria. Etiopathogenesis of cystine in cats is not well charac­ terized, but one study recently reported an SLC3A1 mutation causing cystinuria in a cat as well as three dif­ ferent SLC7A9 gene variants in four cats. Resources for cystinuria testing (urine nitroprusside/ genetic testing) can be found at PennGen Laboratories: http://research.vet.upenn.edu/penngen

crops. Plant sources implicated in silicate urolithiasis include rice and soybean hulls and corn gluten feed. Silicates may be high in certain home‐made diets and in some groundwater from aquifers. In certain regions, soil may contain high concentrations of silicate so consump­ tion of dirt and grass (pica) should be discouraged. Experimentally, beagle dogs consuming large quanti­ ties of antacids containing magnesium trisilicate were predisposed to silica uroliths. Homeopathic, other medi­ cations and vitamin‐mineral supplements may contain silica, for example silica is used in anticaking materials added to tablets. Other Types of Urolithiasis In dogs, mixed composition uroliths comprised 2.3% of uroliths submitted to the MUC, with an additional 8.8% of uroliths being described as compound. Compound uroliths are likely to form when factors initially promot­ ing precipitation of one type of mineral are replaced by factors promoting precipitation of a different mineral. This can make compound uroliths poorly responsive to medical management due to the need to treat each min­ eral type separately for dissolution or prevention. The list below includes uncommon types of uroliths submitted to major urolith centers. ●●

Silicate Urolithiasis Epidemiology

Silica (silicon dioxide) uroliths are infrequently diag­ nosed. They account for 0.9% of specimens submitted to the MUC and 0.7% of specimens submitted to the CVUC between 2009 and 2012 (see Table 123.2). Silica‐contain­ ing nephroliths are rarely submitted to urolith centers, but have been reported in native Kenyan dogs. Some silica uroliths are pure, while others are composed of different layers, with oxalate and struvite being the most commonly associated minerals. Most silica uroliths have a “jackstone” appearance; however, an outer layer (coating) of another mineral type may mask this configuration. Typically, multiple silica uroliths present and cross‐sec­ tion of the urolith usually reveals laminations. The majority of dogs with silica uroliths are male, with a mean age of 7.2 +/‐ 3.1 years (range 1–17 years).

●●

Suture induced: some stones can be induced by residual or retained suture material. Uroliths typically have a piece of suture contained within the stone, or have a hole where the suture was located (Figures 123.7 and 123.8). Dried solidified blood calculi: calculi composed solely of dried solidified blood have been removed from the urinary tract of cats.

Etiopathogenesis

Silica uroliths are thought to form due to individual ani­ mal increase in consumption of this mineral. Animal protein contains very low concentrations of silica and consumed dietary sources are likely plant origin, such as direct ingestion of soil or ingestion of contaminated

Figure 123.7  Suture‐induced stone removed from the bladder of a 7‐year‐old male neutered Yorkshire terrier. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

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urine stasis, high rate of urinary excretion of drugs that are poorly soluble in urine, and/or prolonged treat­ ment with high doses of potentially lithogenic drugs.

­Urolith Diagnosis Urinalysis Urine Sediment

Urolithiasis typically induces inflammatory urine sedi­ ment, such as pyuria (presence of white blood cells), hematuria (red blood cells), and proteinuria. Bacteria may also be present if there is a primary or secondary UTI, (i.e., infection induced by urolith‐associated trauma). Figure 123.8  Suture‐induced stone (60% struvite, 40% calcium phosphate) removed from a 4‐year‐old female spayed miniature schnauzer. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre. ●●

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Potassium magnesium pyrophosphate: the etiology of these uroliths is not definitively known, and it is theo­ rized that formation is related to a type of temporary or permanent enzymatic dysfunction causing pyroph­ osphate supersaturation of the urine. Melamine and/or cyanuric acid‐induced uroliths: after a devastating occurrence of diet‐related renal failure in dogs and cats, melamine urolithiasis or crystalluria was noted in affected patients (Figure 123.9). Drug and drug metabolite uroliths: these are most commonly related to the use of the sulfonamides or allopurinol and there are reports of fluoroquinolone (ciprofloxacin), primadone, and tetracycline uroliths. Factors predisposing to precipitation of drugs in urine include reduced volume of highly concentrated urine,

Urine pH

The urine pH of dogs and cats depends on many factors and can be affected by method of measurement. A study in dogs found that urine dipstick tended to overestimate pH, in some cases yielding an alkaline result for a sample that had a mildly acidic urine as determined by the refer­ ence method. As such, if accurate measurement of urine pH is important, a pH meter should be used. Relatively inexpensive hand‐held instruments are commercially available. Certain minerals are more soluble in acidic or alkaline pH. For these reasons, some minerals tend to precipitate and form stones in acidic (e.g., cystine, purine, calcium oxalate) or in alkaline pH (e.g., struvite, calcium carbon­ ate, and calcium phosphate stones). At present, research indicates conflicting results regarding the role urine pH plays in the formation of calcium oxalate stones. Crystals

Crystals may indicate the presence of uroliths or be nor­ mal, that is, no urolith. As such, specific therapy for crystal­ luria is not indicated unless a urolith is documented through imaging, the patient has a history of urolithiasis or urethral plugs (cats), and/or specific crystal types are noted. In order to correctly identify crystalluria, urine should be analyzed within one hour of collection. This is advised because both calcium oxalate and struvite crystals can form in vitro if there is a delay prior to analysis or due to refrigeration. Crystal types that are not normally present in the urine, and demand attention regardless of urolith presence, include cystine, xanthine, and urate (excluding Dalmatians). Urine Culture

Figure 123.9  Melamine cyanuric acid removed from the bladder of a Burmese cat. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

Urine culture and susceptibility should be performed in all patients with urolithiasis in order to detect infections and start appropriate antibiotic treatment if necessary,

123  Urolithiasis in Small Animals

that is, there are concurrent clinical signs of infection. Urinary tract infections can be the primary cause of uro­ lith formation (e.g., struvite uroliths in dogs), or they can be secondary to urolith‐induced irritation of the urinary tract. If the urine culture is negative despite a high suspi­ cion of UTI in a patient (i.e., a dog with struvite stones), culture of the bladder wall and stone nidus, at the time of stone removal, is strongly advised in order to accurately direct any needed antimicrobial therapy.

example of this is a calcium oxalate stone forming in an at‐risk breed of dog after an incidence of struvite uro­ lithiasis due to bacterial infection. Stone Layers

When a urolith is analyzed, it is named according to the constituents making up the majority of the “stone” (Figure 123.10). These layers consist of the following. ●●

Bloodwork Complete blood count (CBC) is usually normal in patients with urolithiasis. A left shift may (or may not) be present with pyelonephritis or pyonephrosis. A serum biochemistry may show evidence of liver dysfunction in a dog with hepatic vascular abnormalities (portosys­ temic shunt [PSS]), microvascular dysplasia (portal hypoplasia), or severe hepatic disease. The lab changes can include increased alanine aminotransferase (ALT) and alkaline phosphatase (ALP), and with hepatic dys­ function there can be low albumin, urea, and glucose. Bile acid (pre‐ and postprandial) measurement is strongly recommended if a young dog or cat presents with bladder stones and clinical signs compatible with PSS (i.e., signs of hepatoencephalopathy, polyuria/poly­ dipsia [PU/PD], poor growth) or increased liver enzymes. Dogs with portovascular abnormalities are more likely to have urate uroliths than those with other types of hepatic dysfunction. Middle‐aged, small‐breed dogs with a por­ tovascular anomaly and good portal perfusion can have normal chemistry panels with urate stones, making bile acid testing indicated. In cases of ureteral calculi resulting in urinary tract obstruction, blood urea nitrogen (BUN) and creatinine may or may not be increased. This will depend on the laterality (unilateral or bilateral obstruction) and under­ lying intrinsic renal damage due to the calculi. Azotemia will not be present if the contralateral (nonobstructed) kidney is normal. Most cats (88%) with unilateral ure­ teral calculi are azotemic (>90%), indicating the presence of intrinsic renal disease in the contralateral kidney.

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Nidus (center of the urolith): this may or may not be composed of crystalline material and/or a foreign body, such as suture material. Stone: the stone may or may not be composed of the same minerals as the nidus. Shell: the layer(s) that surround the stone. The shell is the outermost complete layer on the urolith. Surface crystals: these may or may not be present on the outermost surface of the urolith. They do not com­ pletely encase the urolith (incomplete layer).

Ideally, all four layers of the urolith should be analyzed separately and their respective mineral amounts listed. Only quantitative analysis ensures that all urolith layers are examined and provides an estimate of the mineral content of each visible layer. This is important since con­ fusion over urolith content and therapy and prevention can occur when only the outer layer is analyzed. This type of confusion can also occur when using the qualitative

CANADIAN VETERINARY UROLITH CENTRE Terminology for Urolith Reporting*

Shell Surface Crystals

Stone

Nidus

Stone Analysis A common pitfall for clinicians dealing with urolithiasis in small animals is attempting to guess the type of stone(s) noted on imaging based on signalment of the patient, diet, urine pH, urine culture results, and radio­ graphic density of the stone. For this reason (and if pos­ sible), it is always recommended to submit uroliths for mineral analysis. It is important to note that even if there is recurrence of urolithiasis in the same patient, the uro­ lith type may be different from the prior episode. An

* These terms will be used in all reports from the Canadian Veterinary Urolith Centre.

Figure 123.10  Stone layers description. Source: Courtesy of Andrew Moore, Canadian Veterinary Urolith Centre.

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Section 10  Renal and Genitourinary Disease

test for urolith analyses, where the entire urolith is crushed and then analyzed. An example of why this is important would be with a primary metabolic stones (i.e., urate) which may have a large (or several) outer covering (s) of struvite. If only the outer layer was assessed or if the urolith was crushed and then analyzed, then ongoing diagnostics and therapy would be misdirected, at the struvite rather than the urate portion.

linked marker test to identify dogs at risk for early stone formation is being evaluated. Imaging of the Urinary Tract Radiography

Specific Tests to Detect Cystinuric Dogs

Most uroliths are radiopaque on survey abdominal radi­ ographs. Radiopaque uroliths include calcium p ­ hosphate, calcium oxalate, struvite and silicate (Figures 123.11 and 123.12). Survey radiography is suitable for identification of these radiopaque uroliths, ­provided they are greater than approximately 2–3 mm in diameter. However, urate, cystine, and xanthine uroliths are generally character­ ized as radiolucent or only slightly radiopaque on survey radiographs. The failure rate for detection of uroliths by survey radiography was 2–27% dependent on urolith size and composition. Clinicians may try to guess the chemical composition of a urolith based on the density, shape, and number of uroliths on radiographs. One aid for this type of “guessti­ mate” is the pneumonic POCUS, which is used by clini­ cians to remember the order of the stones that are most radiodense to radiolucent: calcium phosphate, calcium oxalate, struvite, silicate, cystine, and urate calculi. Contrast‐enhanced radiographic procedures or ultra­ sound are needed to identify uroliths composed of com­ pounds similar to the radiographic density of soft tissue. Double‐contrast cystography has been associated with high sensitivity and low false‐negative detection rates (Figure 123.13). Small uroliths are detected more reliably by double‐contrast cystography or ultrasonography. However, ultrasound is superior to double‐contrast cystogram in experienced hands and for very small stones and sand.

On average, dogs with type I cystinuria have a several‐ fold higher urinary COLA excretion than non‐type I cystinuric male dogs. A simple urinary screening test to determine if a dog is cystinuric is available through the Metabolic Screening Laboratory, Section of Medical Genetics, at the University of Pennsylvania (http:// research.vet.upenn.edu/penngen). This test can detect any type I cystinuric animal, but not necessarily all dogs with non‐type I cystinuria. High‐performance liquid chromatography can be further used to determine the amount of cystine and other amino acids in urine. However, this form of quantitative amino acid analysis is currently restricted to a few laboratories (PennGen and LaboKlin) and is relatively expensive. Based on studies, dogs with either cystine levels of >200 μmol/g creatinine or COLA values of >700 μmol/g creatinine are consid­ ered cystinuric. Moreover, for Newfoundlands and Labrador retrievers, a breed‐specific mutation test is available that detects not only cystinuric dogs but also asymptomatic carriers. Finally, in mastiffs, a preliminary

Figure 123.11  Abdominal radiographs (lateral view) showing bladder stones in a dog. Source: Courtesy of Stephanie Nykamp, Radiologist, Ontario Veterinary College.

Stone Characterization

Uroliths are defined based on their mineral composi­ tion as noted earlier. The following terms are used to identify type. ●●

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Pure urolith: when 70% or more of the urolith is com­ posed of one type of crystalloid, it is named for that crystal type. In dogs and cats, most stones have one major crystal component. Mixed urolith: when less than 70% of the urolith is composed of one mineral, it is called a mixed urolith. This means that the urolith does not contain one pre­ dominant mineral type but is a combination of two or more types. Compound urolith: this is a urolith in which each layer is composed of a different mineral. Uroliths that have formed around foreign material in the urinary tract are often classified as compound. Suture material, urinary catheter, metal pellets, plant material, hair strands, and other foreign objects (ex. porcupine quill) have all been reported to be a part of uroliths of various min­ eral types.

123  Urolithiasis in Small Animals

Figure 123.12  Abdominal radiographs (lateral view) showing bladder and urethral stones in a dog. Source: Courtesy of Stephanie Nykamp, Radiologist, Ontario Veterinary College.

transducer, and found that ultrasonography may be more sensitive than survey and contrast cystography for detection of urocystoliths, with false‐negative rates of 6%. Ultrasonography allows simultaneous evaluation of the upper urinary tract for nephroliths and ureteroliths. Therefore, radiography combined with ultrasonogra­ phy are preferred over contrast radiography alone. Ultrasound can detect upper urinary tract obstruction that is characterized by renal pelvis dilation >13 mm. However, pelvis dilation can be iatrogenic (diuresis) or secondary to different diseases such as chronic kidney disease or pyelonephritis. Although a renal pelvis >13 mm may indicate ureteral obstruction, the absence of dilated renal pelvis does not rule out an upper urinary tract obstruction. The combination of proximal ureteral dilation and normal kidney pelvis diameter can be asso­ ciated with ureteral obstruction in cats. In a recent study, 25% of cats with confirmed ureteral obstruction had a renal pelvis 75% water: high moisture). Avoidance of dietary urinary acidification (pH