Zoo Animal and Wildlife Immobilization and Anesthesia -  - ebook

Zoo Animal and Wildlife Immobilization and Anesthesia ebook

0,0
699,99 zł

Opis

Zoo Animal and Wildlife Immobilization and Anesthesia, SecondEdition is a fully updated and revised version of the firstcomprehensive reference on anesthetic techniques in captive andfree-ranging wildlife. Now including expanded coverage ofavian and aquatic species, this exhaustive resource presentsinformation on the full range of zoo and wildlife species. Coveringtopics ranging from monitoring and field anesthesia to CPR andeuthanasia, the heart of the book is devoted to 53 species-specificchapters providing a wealth of information on little-known andcommon zoo and wildlife animals alike. In addition to new species chapters, the new edition brings a newfocus on pain management, including chronic pain, and moreinformation on species-specific physiology. Chapters on airwaymanagement, monitoring, emergency therapeutics, and fieldprocedures are all significantly expanded as well. This update toZoo Animal and Wildlife Immobilization and Anesthesia is aninvaluable addition to the library of all zoo and wildlifeveterinarians.

Ebooka przeczytasz w aplikacjach Legimi na:

Androidzie
iOS
czytnikach certyfikowanych
przez Legimi
Windows
10
Windows
Phone

Liczba stron: 2888




Table of Contents

Title page

Copyright page

Dedication

Contributors

Preface

Section I: General

1: Clinical Pharmacology

Introduction

Pharmacokinetics

Pharmacodynamics

Inhalant Anesthetics

Injectable Anesthetics

Opioids

Nonsteroidal Anti-inflammatories

Alpha-2 Adrenergic Receptor Agonists and Antagonists

Phenothiazines and Butyrophenones

References

2: Monitoring

Introduction

Monitoring Basics

Monitoring the Respiratory System

Monitoring the Cardiovascular System

References

3: Airway Management

Introduction

General Principles of Airway Management

Approaches to Airway Management

Maintenance of the Airway during Anesthesia

Aids to Tracheal Intubation

Methods of Tracheal Intubation

References

4: Thermoregulation

Monitoring Body Temperature

Hypothermia

Perioperative Heat Loss

Hyperthermia

References

5: Oxygen Therapy

Why O

2

?

Hypoxemia and Hypoxia

Oxygen Sources

Methods of Administering Oxygen

Detection of Hypoxemia and Monitoring the Effects of Oxygen Therapy

Oxygen Cylinder Use in Wildlife

Portable Oxygen Concentrator Use in Wildlife

Postanesthetic Effects of Hypoxemia

Oxygen Toxicity

References

6: Analgesia

Introduction

Physiology of Pain

Principles of Pain Management

Pain and Its Management in Vertebrate Species

References

7: Physical Capture and Restraint

Introduction

Mammals

Birds

Reptiles

Reptiles

Amphibians

References

Webliography

8: Zoo and Wildlife CPR

Introduction

Defining Basic and Advanced Life Support

Cardiopulmonary Arrest (CPA) and CPCR: Strategy

Open-Chest or Internal Cardiac Massage

The Alphabet of an Arrest: D, E, F

Monitoring Efficacy of CPCR

Discontinuing CPCR

Prognosis and Return of Spontaneous Circulation?

References

9: Field Emergencies and Complications

Introduction

Precapture Considerations

Animal Safety Considerations for Remote Drug Delivery Equipment

Complications

Conclusion

References

10: Euthanasia

Introduction

Considerations for Wildlife and Zoo Species

Summary

References

11: Remote Drug Delivery

Introduction

Direct Delivery for Cooperative Animals

Remote Delivery Systems for Uncooperative Animals

Conclusions

References

12: Capture Myopathy

Introduction

History

The Human Comparative

Etiology

Predisposing Factors

Pathophysiology

Clinical and Pathological Syndromes

Differential Diagnoses

Treatment

Prevention

Conclusions

References

13: Human Safety during Wildlife Capture

Introduction

Risks to Human Safety during Wildlife Capture and Handling

Emergency Preparedness

References

Section II: Invertebrates, Fish, Reptiles, and Amphibians

14: Invertebrates

Introduction

Taxonomic Groups

Pain Management

Euthanasia

References

15: Bony Fish (Lungfish, Sturgeon, and Teleosts)

Introduction

Stress and Anesthesia

Taxonomy, Anatomy, Physiology, and Behavior

Environmental and Other Factors

Vascular Access

Immobilization Methods and Techniques

Monitoring

Recovery

Resuscitation

Anesthetic Drugs

Nonchemical Anesthesia

Analgesia

Euthanasia

Field Immobilization

Postanesthetic Challenges

Acknowledgments

References

16: Elasmobranchs and Holocephalans

Introduction

Anatomy and Physiology

Capture

Physical Restraint

Vascular Access

Preanesthetic Considerations

Chemical Immobilization

2-Phenoxyethanol (2-PE)

Monitoring

Recovery Considerations and Postanesthetic Challenges

Field Immobilization

Euthanasia

Acknowledgments

References

Personal Communications

17: Amphibians

Introduction

Anatomy and Physiology

Vascular Access

General Anesthetic Considerations

Monitoring

Analgesia

Anesthetic Drugs

References

18: Crocodilian Capture and Restraint

Introduction

Capture Equipment

To Catch a Crocodilian

Capture and Handling Basics

Capture and Restraint

Restraint and Transport

Release Techniques

References

19: Crocodilians (Crocodiles, Alligators, Caiman, and Gharial)

Introduction

Physiology and Anatomy

Restraint Techniques

Drug Delivery

Monitoring

Analgesia

Tranquilizers and Sedatives

Local Analgesia

Immobilizing Drugs

Other Injectable Agents

Inhalant Anesthesia

Anesthetic Protocols

References

20: Venomous Reptile Restraint and Handling

Introduction

Snake Behavior

Management Guidelines

Equipment and Methods

Venomous Lizard Management

Acknowledgments

Products Mentioned in the Text

References

21: Squamates (Snakes and Lizards)

Introduction

Taxonomy and Biology

Anatomy and Physiology

Vascular Access Sites

Preanesthetic Considerations

Maintaining Body Temperature

Monitoring Physiological Function

Monitoring Depth of Anesthesia

Endotracheal Intubation

Ventilation

Inhalation Anesthesia

Inhalation Anesthetics

Parenteral Anesthesia

Other Techniques

Recovery

Complications

Analgesia

Selected Protocols

Field Techniques

References

22: Chelonia (Tortoises, Turtles, and Terrapins)

Introduction

Anatomy and Physiology

Perianestehtic Considerations and Patient Management

Analgesia

Anesthetic Induction and MAINTENANCE

Monitoring

Recommended Sedatives and Anesthetic Agents

Recovery

References

Section III: Bird Anesthesia

23: Avian Anatomy and Physiology

Introduction

Respiratory System

Cardiovascular System

Thermoregulation

References

24: Cagebirds

Introduction

Preanesthetic Evaluation

Preanesthetic Supportive Care

Equipment

Thermal Support

Emergencies

Anesthesia

Preanesthetics

Injectable Anesthetics

Local Anesthesia

Monitoring

Analgesia

Appendix

References

25: Penguins

Introduction

Anatomy and Physiology

Capture and Physical Restraint

Vascular Access

Endotracheal Intubation

Preanesthetic Considerations

Induction and Maintenance Protocols

Monitoring

Recovery

Field Anesthesia

Postanesthetic Challenges

References

26: Ratites

Introduction

Physical Restraint

Vascular Access

Endotracheal Intubation

Analgesia

Chemical Restraint and Anesthesia

Recovery

Field Anesthetic Techniques

Monitoring

Complications

Diseases of Concern

References

27: Raptors

Introduction

Anatomy and Physiology

Capture and Restraint

Ancillary Aspects of Anesthesia in Raptors

Vascular Access

Injectable Drugs for (Sedation) and Anesthesia

Local and Regional Anesthesia (Nerve Blocks)

Inhalant Anesthesia

Monitoring

Vascular Support

Recovery

Anesthetic Administration through an Air Sac Cannula

Special Considerations

References

28: Galliformes and Columbiformes

Introduction

Anatomy and Physiology

Physical Restraint

Vascular Access

Endotracheal Intubation

Pre-Anesthetic Considerations

Local and Regional Anesthesia

Analgesia

Induction and Maintenance Protocols

Monitoring

Field Immobilization (Wild Capture)

References

29: Free-Living Waterfowl and Shorebirds

Introduction

The Patient

Fasting

Anesthetic Environment

Physical Restraint

Respiratory Control

Monitoring

Maintenance of Body Temperature

Recovery

Anticholinergics

Inhalation Anesthesia

Oxygen and Anesthesia

Parenteral Anesthesia

Vascular Access

Analgesia

Field Techniques

References

30: Birds: Miscellaneous

Introduction

Anatomy and Physiology

Capture and Physical Restraint

Vascular Access

Endotracheal Intubation

Preanesthetic Considerations

Analgesia

Induction/Maintenance Protocols

Monitoring

Recovery Considerations

Field Immobilization (Wild Capture)

References

Section IV: Mammal Anesthesia

31: Monotremes (Echidnas and Platypus)

Introduction

Echidnas

Platypus

References

32: Marsupials

Introduction

Anatomy and Physiology

Dasyuromorphia

Peramelemorphia

Koalas (Diprotodontia)

Wombats (Diprotodontia)

Possums and Gliders (Diprotodontia)

Macropods (Diprotodontia)

Notoryctemorphia

Didelphimorphia, Paucituberculata and Microbiotheria

References

33: Insectivores (Hedgehogs, Moles, and Tenrecs)

Taxonomy and Biology

Physical Restraint

Preanesthetic Preparation

Parenteral Anesthesia

Inhalation Anesthesia

Monitoring and Supportive Care

References

34: Edentata (Xenartha)

Introduction

Preanesthetic Preparations

Physical Restraint

Induction

Maintenance

Support

Recovery and Complications

References

35: Tubulidentata and Pholidota

Introduction

Tubulidentata

Pholidota

References

36: Chiropterans (Bats)

Introduction

Physiology and Anatomy

Zoonotic Diseases

Physical Restraint

Blood Collection and Handling

Parenteral Anesthesia

Inhalation Anesthesia

References

37: Prosimians

Introduction

Special Physiology

Restraint

Vascular Access Sites

Endotracheal Intubation

Pre-Anesthetic Considerations

Nonsteroidal Anti-Inflammatory Drugs (NSAIDS)

Benzodiazipines

Opioids

Alpha-2 Agonists

Induction Agents

Inhalation Anesthesia

Monitoring

Recovery

Remote Immobilization and Field Techniques

References

38: Monkeys and Gibbons

Introduction

Taxonomy and Biology

Physiology

Human Safety

Vascular Access

Physical Restraint

Psychological Restraint (Taming)

Endothracheal Intubation

Preanesthetic Considerations

Sedation and General Anesthesia

Monitoring

Analgesia

Complications

Recovery

Field Immobilization (Wild Capture)

Suggested Protocols

References

39: Great Apes

Introduction

Taxonomy, Biology, and Anatomical Considerations

Preanesthetic Considerations

Preanesthetics/Sedatives

Induction Agents

Anesthetic Maintenance

Analgesia

Monitoring

Anesthetic Recovery

Complications

Field Anesthesia

References

40: Canids

Introduction

Species-Specific Physiology and Unique Anatomic Features

Physical Restraint

Vascular Access

Intubation

Preanesthetic Considerations

Chemical Restraint and Anesthesia

Postanesthetic Challenges

References

41: Ursids (Bears)

Species-Specific Physiology

Chemical Restraint and Anesthesia

Monitoring Anesthesia and Supportive Care

Pharmacological Considerations

Species-Specific Concerns

References

42: Procyonids and Mustelids

Introduction

Biology

Chemical Restraint, Immobilization, and Anesthesia

Inhalation Anesthesia

References

43: Viverrids

Introduction

Species Specific Physiology

Physical Restraint

Chemical Restraint and Anesthesia

Endotracheal Intubation

References

Webliography

44: Hyenidae

Introduction

Taxonomy and Biology Related to Anesthesia and Handling

Vascular Access and Sample Collection Sites

Restraint

Field Techniques

Body Weights and Blood Values

Captive Hyena Immobilization and Anesthesia

Anesthetic Maintenance

Analgesia

Complications

Recovery

Disease Issues (Affecting Anesthesia)

Field Immobilization of the brown Hyena (

H

yaena brunnea

) in Namibia

References

45: Felids

Introduction and Taxonomy

Anatomy Related to Immobilization and Anesthesia

Physical Restraint

Tranquilization or Premedication

Other Tranquilization Agents

Induction and Chemical Immobilization

A Fully Reversible Injectable Combination

Intravenous Inductions

Supplementation of Injectable Agents

Inhalation Agents

Antagonist or Reversal Agents

Monitoring

Local and Regional Anesthesia

Analgesia

Acknowledgments

References

46: Phocid Seals

Introduction

Anatomy and Physiology

Capture and Physical Restraint

Vascular Access

Endotracheal Intubation

Preanesthetic Considerations

Induction and Maintenance Protocols

Monitoring

Supplemental Drugs Used during Anesthesia

Recovery

Field Anesthesia

Analgesia

References

47: Otariid Seals

Introduction

Capture and Physical Restraint

Vascular Access

Endotracheal Intubation

Preanesthetic Considerations

Induction/Maintenance Protocols

Monitoring

Field Immobilization (Wild Capture)

Analgesia

Acknowledgments

References

48: Walrus

Species-Specific Physiology

Physical Capture and Restraint

Chemical Restraint and Anesthesia

Monitoring Anesthesia and Supportive Care

Pharmacological Considerations

Analgesia

References

49: Cetaceans

Introduction

Dolphin Sleep

History of Dolphin General Anesthesia

Anatomy and Physiology Relevant to Anesthesia

Physical Restraint

Vascular Access

Endotracheal Intubation

Anesthesia Support and Monitoring

Field Techniques and Strategies

Body Weight Ranges of Some Cetacean Species

Analgesia

Anesthetic and Sedative Drugs

Recovery

Local Anesthesia

Disease Issues Affecting Anesthesia

References

50: Sirenians (Manatees and Dugongs)

Introduction

Anatomy and Physiology

Capture and Physical Restraint

Vascular Access

Endotracheal Intubation

Preanesthetic Considerations

Induction/Maintenance Protocols

Monitoring

Recovery Considerations

Acknowledgments

References

51: Elephants

General Considerations for Elephant Sedation and Anesthesia

Immobilization of Free-Ranging Elephants

Captive Elephant Procedures

Analgesia

Summary

References

52: Nondomestic Equids

Introduction and Taxonomy

Anatomy and Physiology

Vascular Access Sites and Monitoring

Intubation

Nonchemical Capture of Equids

Chemical Restraint and Capture

References

53: Tapirs

Introduction, Taxonomy, and Natural History

Clinical Anatomy and Physiology

Preanesthetic Considerations

Preanesthetic Medication and Induction

Maintenance Anesthesia

Monitoring and Anesthetic Support

Recovery

References

54: Rhinoceroses

The Rhinocerotidae

Rhinoceros Immobilization and Capture

Rhinoceros Anesthesia in Captivity

Rhinoceros Anesthesia in the Wild

Rhinoceros Crating and Transport

Alternative Rhinoceros Anesthesia Techniques

Rhinoceros Calf Anesthesia

References

55: Nondomestic Suids

Introduction

Taxonomy

General Considerations of Suid Biology and Physiology

Trapping and Physical Restraint

Vascular Access

Endotracheal Intubation and Anesthetic Monitoring and Support

Anesthetic Combinations

Conclusions

References

56: Hippopotamidae

Introduction

Taxonomy and Anatomy Related to Immobilization

Anatomy and Physiology Related to Capture and Immobilization

Cardiovascular System

Respiratory System

Thermoregulation and Body Temperature

Specific Use of Anesthetic Monitoring Equipment on Hippos

Planning for a Hippo Immobilization Procedure

Historical Review of Hippo Immobilization

New Advances in Hippo Restraint and Immobilization

References

57: Camelids

Introduction

Patient Restraint and Handling

Instrumentation

Anesthetic Techniques

Anesthetic Recovery

Analgesic Techniques

Monitoring during Anesthetic-Induced Recumbency

Support during Anesthetic-Induced Recumbency

Summary

References

58: Giraffidae

Taxonomy and Biology

Anatomy and Physiology Related to Anesthesia

Analgesia

Physical and Mechanical Restraint

Sedation and Tranquilization

Endotracheal Intubation

Anesthesia of Giraffe

Okapi Anesthesia

References

59: Cervids (Deer)

Introduction

Species-Specific Physiology

Physical Capture and Restraint

Chemical Restraint and Anesthesia

Monitoring and Supportive Care

Pharmacological Considerations for Anesthesia of Deer

References

60: Antelope

Introduction

Taxonomy and Anatomy Related to Anesthesia

Biology and Physiology

Vascular Access Sites and Monitoring

Intubation

Review of Reports on Antelope Anesthesia

Strategies for Field Capture and Anesthesia in Remote Locations

Guidelines for Anesthesia in Specific Antelope Species

Anesthetic Regimens for Antelope in Managed Care

Induction

Recovery

Anesthesia Records

Acknowledgments

References

61: Gazelle and Small Antelope

Introduction

Unique Physiology, Anatomy, and Behavior

Physical Restraint

Vascular Access

Intubation

Preanesthetic Considerations

Induction Protocols

Maintenance and Monitoring

Recovery Considerations

Tranquilizers

Complications

Appendix 61A.1

References

62: Wild Sheep and Goats

Introduction

Species-Specific Physiology

Physical Capture

Chemical Restraint and Anesthesia

References

63: Nondomestic Cattle

Introduction

Physical Restraint

Vascular Access and Urine Collection

Chemical Restraint and Capture

Pre-Anesthetic Considerations

Anesthetic Monitoring

Endotracheal Intubation

Sedation and Tranquilization

General Anesthesia

Analgesia

Recovery

Complications

References

64: Bison

Introduction

Species-Specific Physiology

Physical Capture and Restraint

Chemical Restraint and Anesthesia

References

65: Lagomorphs (Rabbits, Hares, and Pikas)

Introduction

Physical Restraint

Vascular Access

Preanesthetic Preparation

Premedication

Analgesia

Local and Regional Anesthesia

Parenteral Anesthesia

Inhalation Anesthesia

Perioperative Monitoring

Perioperative Supportive Care

Recovery

Recommended Anesthetic Regimens for Domestic Rabbits

Recommended Anesthetic Regimens for Free-Living Lagomorphs

References

66: Rodents

Introduction

Consequences of Small Body Size

Zoonotic Diseases

Physical Restraint

Preanesthetic Preparation

Premedication

Analgesia

Local and Regional Anesthesia

Parenteral Anesthesia

Inhalation Anesthesia

Monitoring

Supportive Care

Recovery

Anesthetic Regimens for Small Domestic Rodents

Anesthetic Regimens for Free-Living Rodents

References

Index

End User License Agreement

List of Tables

Table 1.1.    Anesthetic agent vapor pressures at 20 and 24°C

Table 1.2.    Selected partition coefficients of commonly used anesthetic agents

Table 1.3.    Structure and characteristics of inhalation anesthetics

Table 5.1.    The effect on arterial oxygen tension by intranasal oxygen supplementation from portable oxygen cylinders at different flow rates in various ungulate species immobilized with various drug combinations

Table 5.2.    Recommended flow rates of intranasal oxygen to anesthetized brown bears in relation to their body mass

Table 6.1.    Commonly used analgesics in nonmammalian vertebrates

Table 6.2.    Commonly used analgesics in mammals

Table 7.1.    Principles and considerations of humane physical restraint in zoo and wild animals

Table 7.2.    Comparison of physical capture techniques for free-ranging wildlife

Table 8.1.    Cardiopulmonary cerebral resuscitation drug doses

Table 8.2.    Sample crash cart setup

Table 10.1.    Euthanasia methods for wildlife and zoo species

Table 11.1.    Description of currently available darts by manufacturer

Table 12.1.    Predisposing or contributing factors for capture myopathy

Table 14.1.    Immobilization and anesthetic drugs used in invertebrates

Table 15.1.    Selected anesthetic agents used in selected bony fish (lungfish, sturgeon, and teleosts)

Table 16.1.    Anesthetic and immobilization drugs used in elasmobranchs

Table 16.2.    Drugs used for emergency and supportive care in anesthetized elasmobranchs

Table 17.1.    Analgesic drug dosages for amphibians

Table 17.2.    Anesthetic drug dosages for amphibians

Table 18.1.    Suggested standard list of materials used for crocodilian capture and restraint

Table 19.1.    Commonly used drugs in crocodilians

Table 21.1.    Selected anesthetic, sedative and analgesic drugs used in snakes and lizards

Table 21.2.    Recommended anesthetic protocols for snakes and lizards

Table 22.1.    Analgesic agents with pharmacokinetic and pharmacodynamic data available in chelonian species

Table 22.2.    Selected sedative and anesthetic protocols used in chelonian species

Table 23.1.    Comparison of allometric equations for respiratory variables in birds and mammals

Table 23.2.    Published blood gas values for selected species of birds

Table 23.3.    Published direct blood pressure (DBP) ranges in avian species

Table 24.1.    Avian emergency drug doses

Table 24.2.    Injectable premedication/emergency, sedative, tranquilizer, and anesthetic drugs used in cagebirds

Table 24.3.    Analgesic drug dosages commonly used in birds

Table 25.1.    Species of penguins

Table 26.1.    Reported weight and height ranges for ratites and tinamous

Table 26.2.    Anesthetic agents used in adult ratites

Table 26.3.    Systolic (SBP), mean (MAP), and diastolic (DAP) arterial blood pressures (mmHg) reported for anesthetized ratites

Table 27.1.    Reported dosages of more commonly used injectable anesthetics in raptors

Table 27.2.    Inhalant anesthetics in raptors

Table 28.1.    Parenteral anesthetics used in domestic galliformes and columbiformes

Table 29.1.    Simplified anesthesia levels of birds. The stages are continuous

Table 29.2.    Mean cardiopulmonary and blood gas values with SD (±) or range (−) for normal waterfowl

Table 29.3.    Mean cardiopulmonary values given as mean with SD (±) or range (−) for waterfowl after at least 15 minutes under injectable anesthesia

Table 29.4.    Injectable drugs used in waterfowl anesthesia

Table 30.1.    Classification of the class Aves

Table 32.1.    Body weights of selected marsupial species (Van Dyke & Strahan, 2008)

Table 32.2.    Dosages (mg/kg) of analgesics and anti-inflammatory drugs used in marsupials (Vogelnest & Woods 2008)

Table 32.3.    Accessible veins in marsupials

Table 32.4.    Heart rate, respiratory rate, and body temperature of selected marsupials

Table 32.5.    Dosages (mg/kg) of tiletamine/zolazepam (TZ), xylazine/ketamine (X/K), medetomidine/ketamine (M/K), and alfaxalone for immobilization of marsupials (Vogelnest 1999; Vogelnest & Woods 2008)

Table 33.1.    Anesthetic and analgesic drugs used in insectivores

Table 34.1.    Normal adult body weights of select Edentate species

Table 34.2.    Doses for anesthetic induction agents in Edentata

Table 35.1.    Chemical restraint agents used for aardvarks

Table 36.1.    Anesthetic and analgesic drugs used in megachiropteran and microchiropteran bats

Table 37.1.    Body weight ranges for various prosimian species

Table 37.2.    Drugs commonly used for sedation and anesthesia in prosimian primates

Table 37.3.    Doses of analgesic drugs used in prosimian primates

Table 37.4.    Useful combination regimes for prosimian primates

a

Table 38.1.    Taxonomy of monkeys including approximate adult body weights

Table 38.2.    Recommended drugs and drug combinations for immobilization of monkeys and gibbons

Table 39.1.    Range of dosages of injectable anesthetic induction and reversal agents used in wild and captive great apes

Table 39.2.    Range of physiologic parameters reported in great apes under various anesthetic regimens

Table 40.1.    Taxonomic and biologic information for canids

Table 40.2.    Injectable immobilization drug dosages for canids

Table 41.1.    Recommended mean doses (mg/kg) of immobilizing agents used to facilitate capture of free-ranging American black bears, brown bears, and polar bears

Table 42.1.    General Procyonidae chemical restraint agent doses (IM)

Table 42.2.    Specific Procyonidae chemical restraint agent doses (IM)

Table 42.3.    General mustelid chemical restraint agent doses (IM)

Table 42.4.    Specific mustelid chemical restraint agent doses (IM)

Table 43.1.    Latin names, common names, body weights, and longevity of viverrid species

Table 43.2.    Anesthetic drug combinations used in viverrids

Table 44.1.    Serum chemistry ranges for captive adult spotted hyenas under KXA anesthesia

Table 44.2.    Complete blood count ranges for captive adult spotted hyenas under KXA anesthesia

Table 45.1.    Dosages for immobilization agents used in large nondomestic felids (

P

anthera

spp.) and cheetahs (

A

cinonyx jubatus

)

Table 45.2.    Dosages for immobilization agents used in small nondomestic felids (

F

elis

spp. and

N

eofelis nebulosa

)

Table 45.3.    Dosages for analgesic and NSAIDs used in nondomestic felids

Table 46.1.    Some chemical immobilizing agents used in phocid seals 1990–2010

Table 46.2.    Classification of stages of immobilization in phocid seals

Table 47.1.    Blood gas variables from the caudal gluteal vein in 10 physically restrained California sea lions (

Z

alophus californianus

)

Table 47.2.    Parenteral and inhalant anesthetic drug dosages in otariids

Table 49.1.    Common juvenile and adult weight ranges for selected species of cetaceans housed in oceanaria and aquaria

Table 49.2.    Analgesic drugs used in cetaceans

Table 49.3.    Chemical sedative and anesthetic agents used in bottlenose dolphins (

T

ursiops truncatus

)

Table 50.1.    Family Sirenia

Table 50.2.    Analgesic, anesthetic, and reversal agents for the Florida manatee (

T

richechus manatus latirostris

)

Table 50.3.    Physiological parameters for the Florida manatee (

T

richechus manatus latirostris

)

Table 51.1.    Geographic distribution, height, and weight of officially recognized elephant taxa

Table 51.2.    Procedures requiring elephant sedation or anesthesia

Table 51.3.    Doses of opiate agonists used in elephant anesthesia

Table 51.4.    Published cardiopulmonary values for etorphine-immobilized free-ranging elephants

Table 51.5.    Drugs used for standing sedation in elephants

Table 51.6.    Published cardiopulmonary values for etorphine-immobilized captive elephants

a

Table 51.7.    Commonly used nonsteroidal anti-inflammatory agents used in elephants

a

Table 52.1.    Present-day wild equids

Table 52.2.    Biological data of the wild equids

Table 52.3.    Suggested dosages for long-acting neuroleptics in selected wild equids

Table 52.4.    Selected anesthetic protocols for wild equids

Table 53.1.    Typical body weight and size for members of Tapiridae

Table 53.2.    Criteria that should be considered when designing an anesthetic protocol for tapirs

Table 53.3.    Summary of protocols utilized to immobilize free-ranging tapirs

Table 53.4.    Anesthetic protocols used in captive tapirs

Table 54.1.    Suggested doses for chemical restraint of adult

captive

rhinoceroses producing anesthetic planes from sedation to recumbency

Table 54.2.    Suggested doses for chemical restraint of adult

wild

rhinoceroses including supplemental agents used for respiratory support

Table 54.3.    Suggested opioid reversal protocols for walking, crate loading and transport of adult African rhinoceroses

Table 54.4.    Suggested doses for immobilization and anesthesia of rhinoceros calves in both

captive

and

wild

environments

Table 55.1.    Species of suids by family and weight ranges

Table 55.2.    Common immobilization protocols used in nondomestic suids

Table 56.1.    New anesthetic combination dosages for captive hippos

a

Table 57.1.    Drug protocols for sedation and anesthesia in domesticated healthy adult camelids

a

Table 57.2.    Drug protocols for sedation and anesthesia in domesticated neonatal camelids

a

Table 58.1.    Drugs used for chemical restraint in giraffe

Table 58.2.    Drugs used for chemical restraint in okapi

Table 60.1.    Drugs used by SANParks in the mass capture of various species

Table 60.2.    Drugs used in South African National Parks for the immobilization of various antelope species

Table 60.3.    Anesthetic induction regimens for selected antelope species with average adult captive body weights

Table 61.1.    Weights

Table 61.2.    Dosages used in gazelle and small antelope

Table 61.3.    Dosages for anesthetic antagonists

Table 63.1.    Average body weights of adult exotic cattle

a

Table 63.2.    Recommended anesthetic agents and protocols for wild cattle

Table 65.1.    Drug dosages (mg/kg, IM, SC or IV) for premedication and sedation of rabbits

Table 65.2.    Analgesic drugs used in rabbits and hares

a

Table 65.3.    Drugs used for induction and maintenance of anesthesia in rabbits (and hares)

Table 66.1.    Drugs used for premedication in rodents

Table 66.2.    Suggested analgesic dosages for rodents

Table 66.3.    Parenteral anesthetic regimens and dosages (mg/kg) for representative rodents

Table 66.4.    Guidelines for endotracheal tube size selection in rodents

List of Illustrations

Figure 1.1.    Effects of opioid agonists.

Figure 1.2.    Activation of κ receptor, but occupation without action at the μ receptor.

Figure 1.3.    Partial activation of μ receptor.

Figure 1.4.    No activation of receptors.

Figure 1.5.    Eicosanoid synthesis.

Figure 2.1.    Pulse oximeter placement on the tongue of a brown bear.

Figure 2.2.    Sidestream capnography on a cockatoo.

Figure 2.3.    Mainstream capnograph placed between the anesthetic circuit and the endotracheal tube of a toucan.

Figure 2.4.    Invasive blood pressure measurement in a reindeer.

Figure 2.5.    Oscillometric cuff placed over the distal limb of a tiger.

Figure 2.6.    Ultrasonic Doppler flow probe being used to monitor heart rate.

Figure 3.1.    Cave racer (

Elaphe taeniura ridleyi

) consciously intubated with a Cole tube prior to being induced with isoflurane.

Figure 3.2.    White rhinoceros (

Ceratotherium simum

) with a nasopharyngeal tube in place that was later used as a nasotracheal tube. Nasopharyngeal tubes can be useful in recovery where nasal edema may lead to obstruction or restricted upper airways in nasal obligate breathers.

Figure 3.3.    A homemade facemask being used for maintenance in a giant anteater (

Myrmecophaga tridacytyla

): such a large apparatus dead space requires high flow rates to prevent rebreathing when compared to a smaller facemask.

Figure 3.4.    An intracuff pressure monitoring device allows accurate selection of cuff pressure and a reduction in risks associated with excessive cuff pressures.

Figure 3.5.    (a and b) The crista ventralis is a projection from the ventral aspect of the cricoid cartilage found in a variety of avian species (a). This reduces the size of tracheal tube that can be used. This is not a bifurcation as can be aseen when the dorsal larynx is removed (b) (Eastern white pelican,

Pelecanus onocrotalus

).

Figure 3.6.    Intravenous cannulae can be used as tracheal tubes in extremely small patients. To attach to the anesthetic circuit a cut down 2.5-mL syringe and a standard 7.5 15-mm connector can be used as an adapter (African spurred tortoise,

Geochelone sulcata

).

Figure 4.1.  Even large animals can become hypothermic while under general anesthesia. A zebra is covered with a forced hot air warmer and a large blanket to prevent radiant heat loss. In addition, the zebra is placed on an insulated pad to prevent conductive heat loss through the concrete floor.

Figure 5.1. Intranasal oxygen supplementation from a portable battery-driven oxygen concentrator to a bighorn sheep in Alberta, Canada.

Figure 5.2.    Oxygen supplementation administered to a bighorn sheep via two nasal lines secured with tape.

Figure 5.3.    The i-STAT

®

1 Portable Clinical Analyzer and cartridges used for field analysis.

Figure 5.4.    Oxygen therapy provided when moving an immobilized desert bighorn sheep in Nevada.

Figure 5.5.    Arterial blood sampling from an auricular artery during intranasal oxygen therapy to a white rhinoceros in Kruger National Park, South Africa.

Figure 6.1.    The pain pathway.

Figure 6.2.    Topical application of a local anesthetic cream (lidocaine/prilocaine) in a White's tree frog (

Litoria caerulea

) prior to microchipping.

Figure 6.3.    Electroacupuncture in a black throated monitor (

Varanus albigularis ionidesi

) for cranial cervical trauma that had resulted in tetraparesis. The monitor had not improved with pharmaceuticals and aqua therapy, but significant clinical improvement was noted within three acupuncture treatments.

Figure 6.4.    Infraorbital nerve block with bupivicaine in a gray wolf (

Canis lupus

) prior to an endodontic procedure.

Figure 6.5.    Mental nerve block with bupivicaine in an Amur tiger (

Panthera tigris altaica

) for an endodontic procedure. This dental block would be useful for premolar extraction for aging studies as well.

Figure 7.1.    Wild elk restrained in a padded manual squeeze near Banff National Park, Canada.

Figure 7.2.    Timber wolf physically restrained with hobbles and duct tape on the muzzle for application of a radio collar.

Figure 7.3.    Wild coyote physically restrained with hobbles and nylon muzzle after capture in modified foot hold trap.

Figure 7.4.    Eurasian eagle owls being restrained for examination and weighing following initial capture with a net in a captive enclosure.

Figure 7.5.    Blue-tongued skink restrained in a towel.

Figure 8.1.    (a) The chest in a relaxed state with proper positioning of the hands over the heart. (b) Compression of the heart, between the right and left chest walls. (c) Blood is forced into the aorta and pulmonary artery as the mitral and tricuspid valves are active and act to prevent back flow of blood into the atrii. (d) During relaxation and chest recoil, the heart reexpands. (e) The valves open and blood passively fills the heart.

Figure 8.2.    (a) The hands are placed over the widest part of the chest. (b) The chest size decreases, but the heart is not directly compressed between the two opposing rib cages. (c) Increases in intrathoracic pressures push blood out of the heart and thorax into the arteries. Note that the tricuspid and mitral valves are open and the heart is acts as a passive conduit. The venous valves play a role in preventing back flow of blood. (d) The chest expands during the relaxation phase. (e) Blood reenters the thoracic vessels and heart passively during the relaxation phase of external CPCR. Note: The heart (and cardiac valves) is passive. There are antireflux valves at the thoracic inlet.

Figure 8.3.    Anatomic location of the internal thoracic artery (red) and vagal nerve (yellow) for canid and felid species, which should be avoided when opening the thorax and pericardium respectively.

Figure 8.4.    (a) ECG tracing of ventricular asystole. Note the absence of any electrical activity (b). An example of pulseless electrical activity (PEA). Note that the ECG tracing can vary from near normal to vary bizarre depending on the electrical activity present. The key to note is that there is synchronous electrical activity from the heart in the absence of a detectable pulse. (c–e) ECG tracing of coarse ventricular fibrillation progressing to fine ventricular fibrillation.

Figure 8.5.    End tidal carbon dioxide measurement and its association with cardiac arrest, effective external cardiac message (ECM), return of spontaneous circulation (ROSC) and initiation of positive pressure ventilation (PPV).

Figure 8.6.    Cardiopulmonary resuscitation.

Figure 9.1.    Preparation of a cushioned landing area for a bear cub, prior to immobilization in a tree.

Figure 9.2.    Cooling a hyperthermic lynx in a pond.

Figure 9.3.    Use of a portable oxygen concentrator to provide supplemental inspired oxygen to a brown bear cub.

Figure 9.4.    Even with lightweight, low velocity darts, impact can cause trauma. This photo shows dart impact trauma in a lynx.

Figure 9.5.    Poor dart placement results in a high risk of complications and injury, including pneumothorax.

Figure 9.6.    Using a pole to check that the bear is safely anesthetized before approaching.

Figure 11.1.    Mechanical restraint of a giraffe using a chute.

Figure 11.2.    Use of a blowpipe for short range injection of a white rhinoceros that would react if hand injected.

Figure 11.3.    Photograph and illustration of a two-chambered compressed gas dart. (a) Posterior plunger that is a one way valve allowing compressed gas to be introduced, but preventing escape. (b) Movable central syringe plunger that divides the two-chambers of the dart. (c) Side-ported needle with a silicone sleeve occluding the port.

Figure 11.4.    Photograph of two side ported needles. (a) This needle has the side port exposed. (b) A similar needle with a silicone sleeve occluding the port.

Figure 11.5.    Photograph of a two-chambered compressed gas dart being filled. (a) Posterior chamber of the dart. (b) Posterior plunger that is a one way valve allowing compressed gas to be introduced, but preventing escape. (c) Coupler that is needed to allow air to pass from the syringe to the dart. (d) Standard syringe used to force air into the dart.

Figure 11.6.    Photograph showing the difference between two types of two-chambered compressed gas darts. (a) Blow dart. (b) Molded nylon dart.

Figure 11.7.    Photograph of two needle types used in two types of two-chambered compressed gas darts. (a) A hypodermic needle that has the sharp tip occluded with solder used in blow darts. (b) A thicker, blunter, machined needle used in molded nylon darts.

Figure 11.8.    Photograph and illustration of a gunpowder explosive-powered dart. (a) Yarn tailpiece that is threaded into the back of the dart body. (b) Movable central syringe plunger, which contains the explosive charge. (c) Forward-ported needle with a wire barb is threaded into the front of the dart body.

Figure 11.9.    Illustration of the tail section and trigger mechanism of a Pneu-Dart gunpowder explosive-powered dart. (a) Movable weight. (b) Spring that holds the weight back. (c) Explosive cap.

Figure 11.10.    Photograph demonstrating the steps in assembling the tail section and trigger mechanism of a Palmer Cap-Chur gunpowder explosive-powered dart. (a) The charge has been placed into the caudal pocket of the central plunger. The needle is pressing the weight and spring to demonstrate the proper orientation of the explosive charge. (b) The plunger and explosive charge have been inserted into the posterior part of the dart body. (c) The yarn tailpiece has been threaded into place.

Figure 11.11.    Photograph of three needle types used in gunpowder explosive-powered darts. (a) Pneu-Dart needle with a green gel cone collar barb. (b) Palmer Cap-Chur needle with small metal collar. (c) Palmer Cap-Chur needle with wire barb.

Figure 11.12.    Illustration of a chemical reaction-powered dart. (a) Represents the loaded dart prior to impact. The chemical reagents are kept separated by a weight occluding the chemical chambers. (b) Represents the dart after impacting the target and during discharge of the drug. The weight was dislodged and the chemicals have mixed and produced a gas that pushes the central plunger forward.

Figure 11.13.    Photograph of two Dan-Inject blowgun type projectors. (a) A model CO2 PI pistol with an 11-mm barrel. The pistol is powered with a compressed CO2 cartridge. (b) A JM Special rifle with 11-mm barrel. It is also powered by a compressed CO2 cartridge.

Figure 11.14.    Photographs of three compressed gas-powered 12.7 mm (.50 caliber) projectors. (a) Palmer Cap-Chur Model #1300 mid range pistol powered with 2 compressed CO2 cartridges. (b) Pneu-Dart Model 178B air pump rifle. (c) Palmer Cap-Chur Model #1200 long-range rifle powered with two compressed CO2 cartridges.

Figure 11.15.    Photographs of two cartridge powered 12.7 mm (.50 caliber) projectors. These guns are very powerful and used only outdoors for long range targets. (a) Palmer Cap-Chur Model #1000 extra long-range rifle. (b) Pneu-Dart Model 196 rifle.

Figure 11.16.    Photograph of a darting kit with essential components separated into compartments. Note that a radio is included to provide emergency communications.

Figure 11.17.    Photograph of three types of dart tails. (a)Four fin tail from a Dist-Inject mini-ject easy dart. (b) Cloth tail from a Dan-Inject molded S series dart. (c) Plastic tail from a Pneu-Dart C dart.

Figure 11.18.    Photograph through the barrel of a rifle barrel showing the spiral riffling needed to cause a dart to spin on its horizontal axis during flight.

Figure 12.1.    Incised urinary bladder of the same white-tailed deer as pictured in Figure 12.4 demonstrating marked myoglobinuria (photo courtesy of Dr. Douglas Whiteside).

Figure 12.2.    Hindlimb adductor muscle of a nilgai diagnosed with capture myopathy after escaping from a zoo exhibit. The animal survived for 1 week postescape. Note the sharp demarcation between normal muscle on the left-hand side and the affected adductor muscle on the far right that has a pale, dry appearance (photo courtesy of Dr. Scott Citino).

Figure 12.3.    Gluteal musculature of a Grant's zebra diagnosed with capture myopathy. The affected muscle is on the far right. It is pale in color, appears to have a dry surface and it is isolated from normal muscle by a septum of deep fascia (photo courtesy of Dr. Scott Citino).

Figure 12.4.    Left hindlimb of a captive white-tailed deer diagnosed with capture myopathy and exhibiting ruptured muscle syndrome. The left lateral musculature is exposed with the tarsus on the left. Note the marked subcutaneous hemorrhage surrounding the hindlimb muscles proximal to the tarsus (photo courtesy of Dr. Douglas Whiteside).

Figure 13.1.    The use of ropes to approach a bighorn sheep on a cliff edge.

Figure 13.2.    Loading a dart in a “splash box” (image courtesy of Dr. Keith Amass).

Figure 13.3.    Bear deterrent spray.

Figure 14.1.    A recirculating anesthesia system containing dilute ethanol with an anesthetized cuttlefish (

Sepia officinalis

). Note that two irrigating tubes are being used, one for each set of gills (photo courtesy of J. Bolynn).

Figure 14.2.    Spiny lobster (

Panularis

spp.) anesthesia: This adult spiny lobster has been anesthetized with a eugenol immersion, followed by eugenol gill perfusion, for some diagnostic procedures (photo courtesy of M. Mehalick).

Figure 14.3.    (a) Wearing latex gloves, or other protective measures, should be taken when handling many invertebrates, like this rose hair tarantula (

Grammostola rosea

). (b) Sensible physical capture and restraint are used to obtain a weight from this rose hair tarantula (photos courtesy of M. Mehalick).

Figure 14.4.    Inhalant anesthetic chamber for terrestrial invertebrates. Pictured here is an anesthetized rose hair tarantula (5% isoflurane) (photo courtesy of M. Mehalick).

Figure 14.5.    Since the spider's respiratory intake is located in the abdominal segment it appears that at least some species can be maintained on inhalant anesthesia, as shown here, while the limbs and cranial body parts are examined or manipulated. This is a rose hair tarantula (

Grammostola rosea

) (photo courtesy of M. Mehalick).

Figure 14.6.    This blue crab (

Callinectes sapidus

) is being manually restrained while hemolymph is collected (photo courtesy of M. Mehalick).

Figure 14.7.    Hemolymph collection from a Madagascar hissing cockroach (

Gromphadorhina portentosa

): This mature male hissing cockroach has been sedated with inhalant isoflurane for hemolymph collection.

Figure 15.1.    (a) Midline caudal tail venipuncture in a grouper (

Epinephelus itajara

) using an 18-gauge spinal needle and extension set. A chamois is placed on the tail to hold the animal in place within a sling during the venipuncture. (b) Lateral approach to caudal vein in a rainbow trout (

Onchorynchus mykiss

).

Figure 15.2.    Cardiac venipuncture in a grouper (

Epinephelus lanceolatus

) using a 16-gauge spinal needle.

Figure 15.3.    Cannulation of the dorsal aorta in a rainbow trout (

Onchorynchus mykiss

).

Figure 15.4.    (a) Operant conditioning of a cobia (

Rachycentron canadum

). Note the animal is targeting on a T-shaped piece of PVC. (b) Operant conditioning of a grouper (

Epinephelus itajara

). The animal swims into a PVC box, which can then be closed/secured to reduce the functional swimming space of the animal.

Figure 15.5.    Sling designed to lift large aquatic animals.

Figure 15.6.    Squeeze device designed from PVC and plastic mesh. To the left of the squeeze is a net that reduced the spaced of the tank. The animal was corralled into the right side of the PVC/mesh (the animal is in the far right bottom corner).

Figure 15.7.    A large grouper (

Epinephalus

sp.) being anesthetized using a pump sprayer to deliver concentrated MS-222 into the oral cavity of the animal.

Figure 15.8.    A simple recirculating system that enables delivery of anesthetic water from a reservoir to the gills and recycling of the effluent back to the fish by use of submersible pump.

Figure 15.9.    A goliath grouper (

Epinephelus itajara

) positioned in dorsal recumbency in a foam holder during anesthesia with MS-222. Note the bifurcated mouthpiece being placed in the buccal cavity for delivery of oxygenated anesthetic water to both sets of gill arches, normograde.

Figure 15.10.    Laced moray eel (

Gymnothorax favagineus

) receiving retrograde ventilation in preparation for an oral surgery.

Figure 15.11.    Modified pole syringe system: a commercially available spring-loaded syringe capable of injecting 5 cc of solution is loaded and charge. The dart setup is backed into a syringe case taped to a PVC pipe, and the dart is taped with paper tape that can easily fall off when the animal is injected. Filtered food dye has been added to the contents of the syringe so injection success can be judged.

Figure 15.12.    An example of a Hawaiian sling. It is recommended that this only be used for long distances with skilled users or for euthanasia purposes due the power of the propulsion of the tip.

Figure 15.13.    Ultrasonography is a useful tool for monitoring heart rate in fish. In specimens of sufficient size, the probe can be placed into the opercular slit as in this goliath grouper (

Epinephelus itajara

). Alternatively, the probe is placed directly over the heart in smaller specimens or small-scaled/scaleless species.

Figure 15.14.    Doppler flow is also used for monitoring heart rate in fish with probes placed as described for ultrasonography. This figure demonstrates placement of the probe onto the area of the heart in a laced moray eel (

Gymnothorax favagineus

).

Figure 15.15.    While not always practical, is it possible to monitor the ECG of fish, as with this laced moray eel (

Gymnothorax favagineus

). Note the retrograde placement of a ventilation tube in the opercular slit.

Figure 16.1.    (a) Cross section of the tail of a blacktip reef shark (

Carcharhinus melanopterus

) demonstrating the more vascular or “red” muscle in comparison with the “white” muscle. Note the location of the artery and vein in comparison to the vertebral column. (b) Assisted ventilation in a sandbar shark (

Carcharhinus plumbeus

). There is a PVC tube in place to keep the mouth open. A continuous flow pump is used to direct oxygenated water over the gills. Water flow out of the gill slits determines appropriate positioning.

Figure 16.2.    (a) A sandbar shark (

Carcharhinus plumbeus

) target feeding during a training session. (b) An eagle ray (

Aetobatus narinari

) targeting over a stretcher in a training session. (c) Behaviorally conditioning a manta ray (

Manta alfredi

) to swim into and through a stretcher (photo credit: Georgia Aquarium).

Figure 16.3.    A partially deflated swimming pool in a larger medical pool used to corral a sandbar shark (

Carcharhinus plumbeus

) into a smaller area for anesthesia. The pool was measured for volume when fully inflated and in the water in order to provide accurate dosing with immersion anesthesia.

Figure 16.4.    (a) A clear vinyl bag (a round hoop at the wide end and a narrower hole, to allow for water flow, at the distal portion of the bag) used to guide a shark into the bag for transport to the surface of the water and further restraint into a sling (photo credit: Gavin Drysdale, uShaka Marine World Durbin, South Africa). (b) Restraint of a bowmouth guitarfish (

Rhina ancylostoma

) in a partially submerged boxnet (photo credit: Georgia Aquarium). (c, d, and e) Retrieval of a largetooth sawfish (

Pristis microdon

) from an exhibit utilizing a boxnet. SCUBA divers corral the animal over the submerged net which is then quickly hoisted to the surface (photo credit: Marj Awai, Georgia Aquarium). (f) A metal bridge across an enclosure with a net (wall-to-wall) that extends to the depth of the tank. The bridge and net can move across the entire tank to isolate animals or push them from one end to the other. (g) The same net as in Figure 16.4b with an underwater view of divers maneuvering the net over coral heads in the enclosure's bottom.

Figure 16.5.    A PVC safety tube placed over the tail of a freshwater stingray (family Potamotrygonidae).

Figure 16.6.    (a) Collection of blood from a ventral tail vessel in a whitetip reef shark (

Triaenodon obesus

). (b) Collection of blood from the dorsal sinus in a vessel in a zebra shark (

Stegostoma fasciatum

). (c) Underwater collection of blood from the pectoral fin vasculature of a whale shark (

Rhincodon typus

) under behavioral control using a specialized feeding technique (photo credit: Georgia Aquarium). (d) Collection of blood from the wing vessel in an eagle ray (

Aetobatus narinari

).

Figure 16.7.    (a) Target training a green sawfish (

Pristis zijsron

) to swim over a foldable restraint device. (b) The same animal, within the folded restraint device, now in the shape of a prism with the animal's head at the far end of the photo. Divers are administering high dose MS-222 over the gills of the animal through the device. (c) The same animal, fully anesthetized and pulled from the water, ready for examination.

Figure 16.8.    An underwater dart system with laser apparatus (AQUADART, Harvey et al. 1988).

Figure 16.9.    Intramuscular injection into the dorsal saddle just ventrolateral to the first dorsal fin in a giant guitarfish (

Rhynchobatus djiddensis

) (photo credit: Georgia Aquarium.)

Figure 16.10.    Cardiac ultrasound of a whitetip reef shark (

Triaenodon obesus

).

Figure 16.11.    Collection of blood from a free-ranging sand tiger shark (

Carcharias taurus

) during a population health assessment project in Delaware Bay (photo credit: Georgia Aquarium).

Figure 17.1.    Ventral pelvic region of a gray tree frog,

Hyla versicolor

. Note the verrucae hydrophilicae, or granular sculpturing of the skin, providing increased surface area for water absorption. This region can have application for drug absorption as well.

Figure 17.2.    Collecting blood from the ventral abdominal vein in a western toad,

Anzxyrus boreas

.

Figure 17.3.    Collecting blood from the axillary plexus in a bullfrog,

Rana catesbeiana

.

Figure 17.4.    Transillumination technique highlighting the ventral abdominal vein in a Borneo eared frog,

Polypedates otilophus

.

Figure 17.5.    A tomato frog,

Dyscophus insularis

, under surgical anesthesia in a liquid anesthetic bath, being monitored by Doppler and pulse oximetry. Note the nostrils are elevated above the water line with gauze pads.

Figure 18.1.    A homemade cable snare used for catching small- to medium-sized crocodilians. The catchpole is made of PVC piping.

Figure 18.2.    A homemade rope snare used for catching small- to medium-sized crocodiles. It can also be used as a jaw noose; the rope is placed around the upper and lower jaws and either twisted or the rope pulled at the distal end to tighten. The pole is made of PVC piping.

Figure 18.3.    Physical restraint of a small crocodilian (

Crocodylus mindorensis

). Note the electrician tape placed around the jaws and the hand grasping the neck and forelimbs.

Figure 18.4.    Physical restraint of large crocodilian (

Crocodylus novaeguinae

) with a restraint board. Cargo straps are used to tie the animal to the board. A towel covers the eyes to reduce struggling.

Figure 19.1.    View of the gular fold of a Chinese alligator (

Alligator sinensis

) being depressed with a tongue depressor to access the epiglottis. Note the oral speculum constructed of a piece of PVC pipe wrapped with tape.

Figure 19.2.    Intravenous blood draw via accessing the lateral coccygeal vein. The needle is inserted at a 90° angle at the lateral midline of the tail, just beneath the lateral spinus process of the vertebral body. This same approach may be used to complete intravenous injections of anesthetic drugs such as propofol.

Figure 20.1.    Full face shields and protective garments are recommended when working with all species of spitting cobras. Red-spitting cobra,

Naja pallida

, Central Florida Zoo and Botanical Gardens.

Figure 20.2.    Pinning behind the head for manual restraint is not recommended. Snakes when struggling can move fangs independently while dislodging the mandible to succeed with envenomation. Eyelash viper,

Bothriechis schlegeli

, The Orianne Society.

Figure 20.3.    Snake hooks (top to bottom): Wide-blade “python hook,” extension hook, double-handled hook, various styles, and “L hook” or pinning hook.

Figure 20.4.    Tongs and forceps (top to bottom): Midwest Tong

®

, Pilstrom Tong

®

, hemostatic forceps, endoscopic forceps, and tissue forceps.

Figure 20.5.    Puncture-resistant gloves can offer protection from snakebite of small- to medium-sized species. However, all handling precautions must be met as this method requires the handler to work in very close proximity to the snake. Tiger rattlesnake,

Crotalus tigris

, Glades Herp Farm.

Figure 20.6.    Shift box and interfacing cage for an arboreal snake. Not shown are transparent top and side panels of shift box. Used for Gold's tree cobra,

Pseudohaje goldii

, Central Florida Zoo and Botanical Gardens.

Figure 20.7.    Shift box in cage. Shift box door can be slid close with a snake hook and locked. Black mamba,

Dendroaspis polylepis

(courtesy of Medtoxin Venom Laboratories).

Figure 20.8.    Shift box/squeeze box. Inner panel manually slides toward the entrance hole in the box for tube restraint or against transparent slotted end for injections (Dallas Zoo, courtesy of Habitat Systems Limited).

Figure 20.9.    Shift box/squeeze box. Features include removable squeeze apparatus, slotted end for injections, two capped side ports for tube restraint, and top capped feeding chamber with sliding false floor for protected feeding. Not shown is removable center divider to create “U-maze” to facilitate total entry of long snakes. Jacksonville Zoo and Gardens, black mamba exhibit,

Dendroaspis polylepis

(courtesy of Habitat Systems Limited).

Figure 20.10.    Traditional shift box. This simple design enables the snake to shift out of cage eliminating direct contact. Snake can then be transferred for anesthesia or other procedures.

Figure 20.11.    Traditional restraint box for clinical and field applications. Primarily used for injections and measuring. Western diamondback rattlesnake,

Crotalus atrox

.

Figure 20.12.    Tube restraint. Position of primary handler and snake in tube just prior to restraint. Florida cottonmouth,

Agkistrodon piscivorus

, Central Florida Zoo and Botanical Gardens.

Figure 20.13.    Tube restraint completed. With large specimens, the primary handler secures the snake and tube while the secondary handler supports and restrains the body. Florida cottonmouth,

Agkistrodon piscivorus conanti

, Central Florida Zoo and Botanical Gardens.

Figure 20.14.    Tube restraint directly from a shift box containing a squeeze apparatus. In addition to tube restraint, the body of this 14′ king cobra (

Ophiophagus hannah

) is also manually held in the shift box during examination to maintain control of the body (Central Florida Zoo and Botanical Gardens, photo credit Sarah Burke).

Figure 20.15.    Anesthesia box. A clear plastic storage box with fitted with an adapter to receive the corrugated breathing tube from the anesthesia machine. The transparent container facilitates viewing to judge level of induction. Rhinoceros viper,

Bitis nasicornis

, Central Florida Zoo and Botanical Gardens.

Figure 20.16.    Prerestraint position for helodermatids. Head should be immobilized prior to manual restraint. Gila monster,

Heloderma suspectum

, Central Florida Zoo and Botanical Gardens.

Figure 20.17.    Helodermatids can be manually restrained by securing a firm grip at the base of the head with a forelimb held between the fingers to maintain control of the upper body. Once elevated, the second hand is used to support and hold the body. Gila monster,

Heloderma suspectum

, Central Florida Zoo and Botanical Gardens.

Figure 21.1.    (a) The oral cavity of a boa constrictor (

Boa constrictor

). Note the rostral location of the partly open glottis. (b). The oral cavity of a boa constrictor showing the endotracheal tube within the glottis after intubation.

Figure 21.2.    Mask induction of a monitor lizard with isoflurane. Note the use of a standard small animal face mask.

Figure 22.1.    Appropriate physical restraint of a medium-size (15 kg) alligator snapping turtle (

Macrochelys temminckii

). Note the position of the hands on either side of the shell. The use of reinforced gloves provide additional protection from the claws.

Figure 22.2.    Physical restraint of a large (30 kg) alligator snapping turtle (

Macrochelys temminckii

). The animal is controlled with one hand on the nucal scute and the other on the caudal carapace. The head of the animal is directed away from the operator.

Figure 22.3.    CT-scan 3D reconstruction of the neck of the Gopher tortoise (

Gopherus polyphemus

). The vascular structures are identified as follows: (a) carotid artery, (b) external jugular vein, (c) internal jugular vein, (d) occipital venous sinus. (e) Indicates a intravenous catheter placed in the left external jugular vein.

Figure 22.4.    Use of the subcarapacial venous sinus for blood withdrawal (a) in a Florida softshell turtle (

Apalone ferox

), and for intravenous injection of propofol (b) in a gopher tortoise(

Gopherus polyphemus

). The needle is inserted on the midline of the neck, at the junction between the skin and the carapacial nucal scute. The needle is advanced caudally while maintaining gentle negative aspiration on the syringe until blood is flowing into the chamber.

Figure 22.5.    Venipunture in a alligator snapping turtle (

Macrochelys temminckii

) using the dorsal approach to access the dorsal coccygeal vein. The main advantage of this venipuncture site is represented by its location in the “safe zone” around the animal. For safety, an assistant should be restraining the animal during the venipuncture procedure.

Figure 22.6.    Single-lumen jugular catheter placed in the external jugular vein of a green sea turtle (

Chelonia mydas

). The catheter was inserted using a over-the-wire technique. The winged proximal portion is secured to the skin with sutures.

Figure 22.7.    Placement of a long-term jugular catheter in an Aldabra tortoise (

Aldabrachelys gigantea

). In the anesthetized animal, the external jugular vein is identified by palpation of the dorsolateral aspect of either side of the neck. A hollow needle is advanced through the skin until blood is aspirated. The catheter is then inserted using the Seldinger technique: a blunt guidewire is passed through the needle (a), then the needle is removed. A dilating device may be passed over the guide wire to slightly enlarge the vessel. Finally, the central line itself is then passed over the guidewire, which is then removed. All the lumens of the line are aspirated and flushed. The catheter is finally secured with sutures to the skin (b).

Figure 22.8.    Lateral radiographic views of the placement of a inthathecal catheter in a Galapagos tortoise (

Chelonoidis nigra

). A Tuohy needle is slowly advanced between the spinous processes of two consecutive coccygeal vertebrae and placed in the vertebral canal (a) into the intrathecal space (space between dura mater and leptomeninx). The catheter is threaded through the needle and advanced a few centimeters into the vertebral canal (b). The needle is then withdrawn over the catheter. The catheter is secured to the skin with sutures to prevent it becoming dislodged. The catheter can be used for repeated administrations of analgesics and local anesthetics. The small black arrows indicate the catheter placed into the intrathecal space.

Figure 22.9.    A piece of PVC pipe used as oral speculum facilitates intubation and protects the endotracheal tube. Both the mouth gag and the endotracheal tube are secured with tape to the head of the animal to prevent dislodgment.

Figure 22.10.    Glottal apparatus of an alligator snapping turtle (

Macrochelys temminckii

) during breath holding (a), and during inspiration (b). Note the perfect seal of the closed glottis. Waiting for the animal to voluntarily open the glottis facilitates the endotracheal intubation.

Figure 22.11.    Electrocardiographic tracing of an anesthetized Galapagos tortoise (

Chelonoidis nigra

). On the tracing, the following waves are identified: (SV) sinus venosus depolarization, (P) atrial depolarization, (R) ventricular depolarization, and (T) ventricular repolarization. Note that systolic phase of the cardiac cycle is physiologically prolonged (R-T interval is approximately 1.6 seconds).

Figure 22.12.    Cardiovascular monitoring of a juvenile alligator snapping turtle (

Macrochelys temminckii

) during anesthesia. A 8-Hz Doppler pencil probe is placed on the side of the neck with the detecting crystal directed toward the thoracic inlet. The ECG is used in a standard three limb lead configuration.

Figure 22.13.    Active warming the patient with a forced warm air blower (Bair Hugger

®

, Arizant Healthcare Inc., Eden Prairie, MN). The maintenance of the patient's temperature within the preferred body temperature range is an important aspect of patient care in chelonians. Unpredictable disarrangement of the chelonian's homeostasis can occur in response to sudden changes in body temperature.

Figure 23.1.    Change in oxygen tension during 8 minutes of apnea in six chickens anesthetized with isoflurane. In one treatment, the chickens were connected to an anesthetic circuit containing oxygen (100% O

2

), and in the second, they were disconnected from the circuit at the onset of apnea (Disconnect).

Figure 24.1.    Various endotracheal tube types are used in avian clinical practice. Cole, noncuffed, and cuffed tubes have all been used successfully in caged birds (a). Catheters can be modified and used as ET tubes for very small patients (b). Intubation is straightforward as visualization of the glottis is not difficult in most caged birds (c).

Figure 24.2.    Transtracheal membrane occluding approximately 75% of the tracheal lumen in a double yellow-headed Amazon parrot. Note the significant vascularization of the membrane. These membranes have been reported most commonly in birds up to several weeks after intubation with both cuffed and non-cuffed endotracheal tubes. Various factors such as over-inflation of the cuff, ventilation techniques and endotracheal tube disinfection protocols have been evaluated but no definitive correlations have been identified to date.

Figure 24.3.    Placement of an air sac cannula. Strict asepsis should be observed when placing the cannula. Feathers over the site for cannulation should be gently plucked, contour feathers taped away from the cannula site and the area should be surgically prepared. The landmarks for air sac cannula placement are caudal to last rib, ventral to the vertebrae and, in most cases, cranial to hindlimb. The proper site for cannula insertion is marked with * (a). Once the cannula has been surgically placed, it is secured in place with suture (b). A piece of sterile gauze can be used as a filter to minimize contamination of the cannula with particulate matter (c).

Figure 24.4.    Sites for intravenous (IV) catheters include the ulnar (basilic) vein (a), medial metatarsal vein (b) or jugular vein. Small bore over-the-needle catheters (24 gauge or smaller) are most often necessary for avian patients.

Figure 24.5.    Intraosseous (IO) catheter placement in the distal ulna of a bird. Strict asepsis should be observed when placing the catheter. Feathers over the site should be gently plucked and the site should be surgically prepared (a). Wearing sterile gloves, the sterile catheter is positioned slightly ventral to the dorsal condyle of the distal ulna and parallel to the bone (b). The bone is held firmly with one hand while the hand other applies firm pressure combined with slight rotation (c). The cannula should be flushed with heparinized saline and should flow easily (d). The insertion site should be covered with an antibiotic ointment and the cannula should be secured with a bandage tape butterfly at the exit point in the skin and suture (d). A bandage can be placed over the cannula site for additional security and to prevent possible trauma or damage to the catheter (e).

Figure 24.6.    In order to provide accurate infusion rates for small avian patients, fluid infusion pumps and syringe pumps are most commonly required.

Figure 24.7.    Peripheral pulse rates can be determined in many birds at the brachial artery (a), the medial metatarsal artery, or the carotid artery (b) either by direct pressure assessment or with a Doppler ultrasonic probe. Heart rate can also be detected by placement of the Doppler ultrasonic probe over the palatine artery (c) on the dorsal palate of some species of raptors and waterfowl.

Figure 24.8.    Arterial catheterization of the brachial artery in a bird. The brachial (a) and carotid arteries are the most common sites for arterial catheterization in birds. Percutaneous placement is utilized most commonly, but sometimes a surgical cut-down procedure is required to confirm the location of the artery (b).

Figure 24.9.    Indirect blood pressure monitoring can be performed using a Doppler ultrasonic probe to detect the arterial flow, a pressure cuff to occlude arterial blood flow and a sphygmomanometer to measure pressures.

Figure 24.10.    Avian capnogram.

Figure 25.1.    Restraint technique for a penguin less than 5 kg (little penguin). The animal is grasped behind the head and then held around the body caudal to the wings.

Figure 25.2.    Restraint technique for a large penguin (King penguin) (courtesy of Village Roadshow Theme Parks).

Figure 25.3.    Restraint technique for a large penguin (King penguin) (courtesy of Village Roadshow Theme Parks).

Figure 25.4.    Emperor penguin: restraint for venipuncture (courtesy of SeaWorld San Diego).

Figure 25.5.    Maintenance of gaseous anesthesia in the emperor penguin. To prevent hyperthermia, ice is placed around the wings and feet (courtesy of SeaWorld San Diego).

Figure 26.1.    The glottis of most ratites, as in this rhea, is large and readily visible at the base of the tongue.

Figure 26.2.    A hood reduces excitement and struggling in ostriches; it is not recommended in other ratites.

Figure 26.3.    Wooden shields are used to protect handlers when approaching partially anesthetized ratites.

Figure 26.4.    The wings of emus and most ratites are small and can be fractured if used to restrain an animal. The basilic vein on the medial surface of the wing can be used for blood collection in sedated or anesthetized animals, and for catheterization for fluid administration under anesthesia.

Figure 26.5.    Intramuscular injections are administered into the upper thigh. This is done with either remote or hand injection.

Figure 27.1.    (a) Safely restrained raptor (red-tailed hawk) on the examination table. (b). Properly restrained large raptor (bald eagle). Notice protective gear worn by the handler, including leather welder's vest, gauntleted gloves, and safety glasses.

Figure 27.2.    Insertion of intubation tube in a falcon (a) and intubated raptor (gyrfalcon) (b) prepped for orthopedic surgery. Note capnography and Doppler unit included in monitoring equipment array. While the latter may have limited utility for monitoring blood pressure, it serves as a useful means of audible monitoring of heart rate.

Figure 27.3.    (a) Large-diameter air sac cannula in place for use as a breathing tube. (b) Small-diameter (8–12 Fr; 2.7–4.0 mm) air sac cannula made from red rubber tube (top) and commercially available dedicated air sac cannula tube (bottom) for use in administering gas anesthesia. Note added length to provide maneuverability without dislodging the tube from its insertion site (see Fig. 27.4c).

Figure 27.4.    (a). Positioning of patient and preparation of insertion site in last intercostal space for air sac cannula: arrow indicates site. (b) Anchoring the air sac cannula in place by suturing tape wings to body wall. (c) Air sac cannula installed and attached to anesthesia administration tubing.

Figure 27.5.    In-line bubbler bottle for humidification of transport and anesthetic gases.

Figure 27.6.    Capnographic unit in use. Note that there are collocated esophageal stethoscope and pulse oximeter transducers (not visible in this image).

Figure 29.1.    Restraint technique used for ducks and small geese. The humeri are grasped with the fingers of one hand. For heavier birds, the other hand should be placed under the feet to support the weight of the bird. Using this technique, birds should not be held for more than a minute or two to avoid damage (photograph courtesy of Scott Larsen).

Figure 29.2.    Temporary restraint of a male surf scoter (

Melanitta perspicillata

), here used to weigh the bird on a scale, by tucking the head under a wing and wrapping the bird snugly with “hook-and-loop” strapping. Care is taken to avoid interference with respiratory excursions of the keel and the duration of such restraint should be measured in seconds (photograph courtesy of Dan Esler).

Figure 29.3.    A lesser Canada goose (

Branta Canadensis

) anesthetized with isoflurane with anesthetic circuit shown. The patient, prepared for surgery, is being electronically monitored with a digital thermometer, capnograph, and pulse oximeter, and a blood sample is being drawn from the jugular vein for blood gas determination.

Figure 29.4.    Placement of a pulse oximeter transmittance probe on the bill of a female mallard duck (

Anas platyrhynchos

). This technique works when there is little or no pigment on the bill (photograph courtesy of Scott Larsen).

Figure 29.5.    Bristle-thighed curlew (

Numenius tahitiensis

) being intubated. The left hand is used to hold the upper and lower bills apart while the right hand inserts the endotracheal tube. It is sometimes easier to intubate long-billed birds by holding the head dorsal side down, which permits the slight curve of the endotracheal tube to follow the curve of the beak (photograph courtesy of Dan Ruthrauff).

Figure 29.6.    Examples of holding and recovering cages made by modifying an animal carrier by the addition of artificial turf (top) or mesh surrounding a PVC-pipe frame (bottom) to reduce fecal soiling of feathers. A turf bottom (top) is safer for a bird like the bar-tailed godwit (

Limosa lapponica

), that has a long bill and toes, to reduce the chance of entanglement. A finer mesh net should be attached to the inside of the wire-grid door and air holes to prevent damage to bills of shorebirds and ducks such as the common eider (

Somateria mollissima

), shown in the bottom picture.

Figure 29.7.    Comparison of equipment and supplies required to perform an identical number of surgeries using isoflurane (left) or propofol (right). The latter system totals about 10% of the weight of the former. The cartons containing the oxygen cylinders bear Hazardous Materials labels.

Figure 29.8.    A 0.508-mm (25-gauge), 1-cm butterfly catheter placed in the tibiotarsal vein of a yellow-billed loon (

Gavia adamsii

) and connected to a syringe holding propofol. The wings of the “butterfly” have been cut off to reduce the chance of dislodging the needle.

Figure 29.9.    Assembly of an air-driven inhalant anesthetic system that drives a vaporizer with compressed air from a low pressure, small-volume air tank filled by an electrical air compressor.

Figure 30.1.    (a) The nares of brown (

Pelecanus occidentalis

) and other pelicans are partially occluded making them predominantly mouth breathers. As in most birds the nares are located at the base of the upper bill. (b) An aggressive American white pelican (

Pelecanus erythrorhynchos

) demonstrating the potential danger of the beak to handlers. Birds will orient on the human eyes to attack because they are constantly moving. The glottis is located deep within the caudal portion of the oropharynx and very mobile. (c) The closed glottis of a brown pelican (

Pelecanus occidentalis

) deep in the caudal portion of the oropharynx. (d) The open glottis of a brown pelican (

Pelecanus occidentalis

). The cartilaginous septum in the larynx can be gently displaced by the tube for endotracheal intubation. Stabilization of the glottis from outside the pouch and a good light source will facilitate intubation. (e) An intubated pelican connected to a nonrebreathing system. A circle breathing system would also be appropriate for this-size bird. The cuff of the endotracheal tube is not inflated to prevent damage to the tracheal mucosa. This bird was induced with propofol administered to effect into a catheter placed in the medial metatarsal vein.

Figure 30.2.    (a) Inhalant anesthetic induction of a toco toucan (

Ramphastos toco

) using a small human pediatric mask placed over the nares at the base of the bill and connected to a nonrebreathing system. (b) An aracari being induced using a mask made from a plastic bottle to accommodate the long beak.

Figure 30.3.    A hospitalized sandhill crane (

Grus canadensis

) demonstrating the use of purpose-made hood to keep the animal quiet. This can also be used during physical restraint and examination, or for transportation.

Figure 30.4.    (a) The oral cavity of an aracari demonstrating the position of the glottis at the base of the thin tongue. (b) The oral cavity of a kookaburra (

Dacelo novaeguineae

) also showing the closed glottis at the base of the tongue.

Figure 30.5.    (a) Intravenous administration of propofol into the medial metatarsal vein of a purple swamp hen (

Porhyrio porphyrio

). (b) A purple swamp hen (

Porhyrio porphyrio

) anesthetized with propofol for surgical implantation of a radiotransmitter. The mid-line surgical site was infiltrated with a combination of lidocaine and bupivacaine. Meloxicam was also administered for additional analgesia. Note the use of a flexible esophageal thermometer (white line) and esophageal stethoscope for monitoring. The bird is intubated and ventilated four to six times a minute using an Ambu bag.

Figure 31.1.    A dorsally recumbent, anesthetized echidna demonstrating the technique for blood collection from the jugular vein.

Figure 31.2.    A manually restrained adult platypus demonstrating the correct position of the hand at the base of the tail to avoid the spurs in adult males.

Figure 31.3.    An anesthetized platypus induced and maintained with isoflurane in oxygen administered through a mask.

Figure 31.4.    Venipuncture can be achieved in platypuses by introducing a small-gauge needle into the sinus at the edge of the bill.

Figure 32.1.    (a) Blood collection from the jugular vein of a Tasmanian devil (

Sarcophilus harrisii

). (b) Blood collection from the saphenous vein of a ringtail possum (

Pseudocheirus peregrinus

). (c) Blood collection from the jugular vein of a ringtail possum (

Pseudocheirus peregrinus

). (d) Blood collection from the cephalic vein of a juvenile common wombat (

Vombatus ursinus

).

Figure 32.2.    A Tasmanian devil (

Sarcophilus harrisii

) demonstrating its large gape and powerful jaws.

Figure 32.3.    Physical restraint of an eastern quoll (

Dasyurus viverrinus

), a small carnivorous marsupial.

Figure 32.4.    Physical restraint of a juvenile koala (

Phascolarctos cinereus

) adapted to human contact. These animals are still capable of injuring with their claws an unwary handler.

Figure 32.5.    Blood collection from the cephalic vein of an awake koala restrained in a large bag.

Figure 33.1.    Inhalation anesthesia is preferred for anesthesia in all insectivores. This white-bellied hedgehog (

Atelerix albiventris