text:
A Short Review of the Chemical Immobilization Principles in Some Common African Wildlife Species
Uploaded by Andrés González Rodríguez
saveSave A Short Review of the Chemical Immobilization Prin... For Later
Steps in Essential Newborn Care
Sankaran's Table of Misams and Remedies
funda 4.docx
A Dying Breed Red List Species GS Paper 3 Environment and Biodiversity
Facial Anesthesia and Paralysis Due to Intra-Arterial Injection of Local Anesthesia
Administering Intramuscular Injection
57670244 Daftar Dosis Dan Sediaan Obat Untuk Anak 160207084634
BIG Job Application
IUCN Adds Wild Crops to the Red List
nj dep green acres program rules - njac 7-36
460_rev
hectors dolphin
Ultravoilet Rays
SOP 15 Table set up
Novel delivery technology for probiotic bacteria
Seize the Diary - Control
UNIVERSIDADE DE LISBOA
FACULDADE DE MEDICINA VETERINÁRIA
A SHORT REVIEW OF
THE CHEMICAL IMMOBILIZATION
PRINCIPLES IN SOME COMMON
AFRICAN WILDLIFE SPECIES
BÁRBARA PASTILHA LOPES FERREIRA
CONSTITUIÇÃO DO JÚRI ORIENTADOR
Doutor José Manuel Chéu Limão Oliveira Doutor Brendan William Tindall
Doutora Sandra de Oliveira Tavares de Sousa Jesus
Doutor Luís Ressano Garcia Pardon Lamas CO-ORIENTADOR
Doutora Sandra de Oliveira
Tavares de Sousa Jesus
DISSERTAÇÃO DE MESTRADO INTEGRADO EM MEDICINA VETERINÁRIA
Doutor José Manuel Chéu Limão Oliveira
"Do you know why is it called veterinary practice?
Because you never stop practicing it"
- Dr. B. Tindall, 2015
To Dr Brendan Tindall, my supervisor, for all the years of knowledge, sarcastic humor
and unforgettable wild adventures in South Africa that I will always remember. You are an
inspiration for everyone who has goals as huge as mine. Thank you for believing in me and
in my ambitions from the beginning, for sharing the same passion that I have for wildlife and
for all the good stories you told me in the last years, usually around the fire of a braai and
under a sky full of stars. You gave me wings to fly.
Thank you to the entire team of Robberg Veterinary Clinic in Plettenberg Bay, who allow
me to be part of the family during the 5 months I spent there, particularly to Kerry who never
left me without work to do; Nix and Viv for always having a smile during chaos and, of course
Yvonne 'Pink Lady' Funk for the friendship, the laugh, the x-rays and the coffee. Also to Dr
Stephan Nell, for teaching me more than I ever expected. Your faith in my skills made me
believe that my life aims are feasible and I can dare to dream even bigger. Baie dankie!
Thank you Wild Spirit, because without your existence, my dream would be just that – a
dream – and it would not have become real. Thank you Mark Rutherfoord, owner of
Gondwana Game Reserve, Ian Olivier and all the wildlife rangers who provide me some of
the best moments of my life along with Brendan and Stephan, performing surgery in your
wild lions. I will never forget that day!
To Dr William Fowlds and Dr Peter Caldwell for teaching me the best of Conservation
Medicine in South Africa and to Dr Jorge Soares, a mentor and a friend over the past years,
for all the memorable captures performed in Mafra.
To Chester Zoo, especially to Dr Steve Unwin, Ian Ashpole and Javier Lopez for the
opportunity and the knowledge about Zoological Medicine.
Thank you Prof Dr Sandra Jesus for the full support and advice during the growing
process of this thesis.
To my family, not only in Portugal but also in Canada. Obrigada a todos, em particular à
minha mãe e ao meu avô, que acompanhou de perto as minhas aventuras em África, Índia,
Canadá e Europa, sempre com histórias suas para contar algures no mundo, levando-me a
sonhar com as mais distantes terras e as mais exóticas culturas do planeta. To Guilherme,
for all the constant jokes about my work and for sharing the same wanderlust disease I have.
Carolina, Diana and Lília for craziness! Wild 'Faunas' for passion and ambition.
To my "zoological" family of Lisbon Zoo, particularly to Miguel Lajas, Rita Ramos, Ana
Poças, Ana Lemos, Inês Guerra and Marta Reis. You all always gave me the support
wherever I was in the world. Thank you for sharing the same level of nerding that I have for
wildlife and conservation, and for never let me give up of my goals.
And last but not the least, my thanks to a challenging hardcore cat with a very special wild
attitude.
A medicina veterinária associada à vida selvagem têm vindo a evoluir nas
últimas décadas graças às constantes mudanças nas técnicas de imobilização,
equipamentos e até fármacos utilizados. Estes progressos, aliados à experiência dos
veterinários, têm vindo a contribuir para uma prática médica mais segura tanto para os
animais como para as equipas envolvidas nos procedimentos.
Durante o estágio na África do Sul, os protocolos anestésicos usados nas
imobilizações químicas (184 dos 245 indivíduos imobilizados) foram analisados para as
diferentes espécies manipuladas, tendo em conta o equipamento de disparo utilizado, o
ambiente envolvente e o propósito das intervenções praticadas. As diferentes imobilizações
químicas foram classificadas como bem-sucedidas (176) ou não (8), tendo sido referidas as
principais complicações que afectaram os procedimentos. Nos casos sem o sucesso
anestésico esperado, recorrendo a um segundo dardo (13 casos) ou culminando na morte
dos individuos (3 casos), as razões para o insucesso foram discutidas e algumas medidas
preventivas para o futuro foram propostas.
É importante que este tipo de informação seja sempre analisado após a
execução de uma imobilização, divulgando as conclusões dessa análise e respectivas
experiências pessoais dos casos, de modo a poderem ser exploradas pelos médicos-
veterinários de vida selvagem para evitar complicações futuras.
Palavras-chave: Anestesia, Medicina de Vida Selvagem, Princípios de Imobilização
Química, Medicina de Conservação, África do Sul.
Due to constant changes in restraint techniques, equipment and even
immobilization drugs, wildlife veterinary practices have improved over the past decades. This
broad progress coupled with the experience of practitioners contributes towards a safer
practice for both the animals and people involved in the procedures.
Anesthetic protocols used for chemical immobilizations performed during an
internship in South Africa (184 of 245 restrained individuals) were analyzed for the various
species approached, taking into account the darting equipment involved, the surrounding
environment and some of the main purposes of each intervention. The different chemical
immobilizations performed were classified as successful (176) or unsuccessful (8), and the
most common complications that affected the procedures were documented. The reasons for
the failings that in cases led to a second darting (13 cases) or to the death of animals (3
cases) are also addressed and preventive measures to avoid them were put forward.
It is important to analyze the information logged after every immobilization
procedure, including personal experiences from each clinical case, and present and
disseminate the conclusions, to help wildlife practitioners worldwide avoid future
complications.
Keywords: Anesthesia, Wildlife Practice, Chemical Immobilization Principles, Conservation
Medicine, South Africa.
ACKOWLEDGEMENTS II
RESUMO III
ABSTACT IV
TABLE OF CONTENTS V
LIST OF FIGURES IX
LIST OF GRAPHICS IX
LIST OF TABLES X
LIST OF ABBREVIATIONS AND SYMBOLS XII
CHAPTER 1. INTRODUCTION 1
1.1 INTERNSHIP TRAINING ACTIVITIES 1
CHAPTER 2. LITERATURE REVIEW 4
2.1 ANESTHESIOLOGY IN WILDLIFE 4
2.2 IMMOBILIZATION TECHNIQUES 5
2.2.1 PHYSICAL RESTRAINT 6
a) Physical Force 6
b) Passive Capture 6
c) Bomas 7
d) Nets 7
e) Net Gun 8
2.2.2 CHEMICAL RESTRAINT 8
a) Oral 8
b) Hand-held Injection 8
c) Pole Syringe 9
d) Blow Dart (Blow Pipe) 9
e) Darts 9
2.3 TYPES OF DARTS AND DART GUNS 11
2.3.1 TWO-CHAMBERED COMPRESSED GAS DARTS (PRESSURIZED) 11
2.3.2 GUNPOWDER EXPLOSIVE DARTS 12
2.3.3 REMOTE DELIVERY SYSTEMS 13
2.4 DARTING SITES 14
2.5 NEEDLE LENGTHS, GAUGES AND CHARACTERISTICS 17
2.6 FACTORS THAT INFLUENCE A CAPTURE 17
2.7 PHASES OF IMMOBILIZATION 19
2.7.1 PLANNING AND PREPARATION 19
2.7.2 APPROACH 19
2.7.3 INDUCTIN 20
2.7.4 HANDLING AND MONITORING 21
2.7.5 REVERSAL/RECOVERY 22
2.7.6 REFLECTION 22
2.8 WILDLIFE PRACTICE APPLIED PHARMACOLOGY 22
2.8.1 INTRODUCTION 22
2.8.2 IMMOBILIZATION AGENTS 23
2.8.2.1 OPIOIDS 23
2.8.2.1.1 Etorphine hydrochloride 24
2.8.2.1.2 Thiafentanil oxalate 25
2.8.2.1.3 Carfentanil 25
2.8.2.2 OPIOID ANTAGONISTS 26
2.8.2.2.1 Butorphanol tartrate 26
2.8.2.2.2 Diprenorphine 26
2.8.2.2.3 Nalbuphine & Nalophine 27
2.8.2.2.4 Naltrexone 27
2.8.2.2.5 Naloxone 27
2.8.2.3 CYCLOHEXYLAMINES 27
2.8.2.3.1 Ketamine 28
2.8.2.3.2 Tiletamine (/Zolazepam) 29
2.8.2.4 TRANQUILIZERS 29
2.8.2.4.1 Acepromazine 30
2.8.2.4.2 Azaperone 30
2.8.2.4.3 Haloperidol 31
2.8.2.4.4 Long-Acting Tranquilizers 31
2.8.2.4.4.1 Zuclopenthixol acetate 31
2.8.2.4.4.2 Perphenazine enanthate 32
2.8.2.5 SEDATIVES 32
2.8.2.5.1 BENZODIAZEPINES 32
2.8.2.5.1.1 Diazepam 33
2.8.2.5.1.2 Midazolam 33
2.8.2.5.2 BENZODIAZEPINE ANTAGONIST: Flumazenil 33
2.8.2.6 α – 2 – ADRENERGIC AGONISTS 33
2.8.2.6.1 Xylazine 34
2.8.2.6.2 Medetomidine 34
2.8.2.6.3 Detomidine 34
2.8.2.7 α – 2 – ADRENERGIC ANTAGONISTS 35
2.8.2.7.1 Yohimbine 35
2.8.2.7.2 Atipamezole 35
2.8.2.8 OTHER DRUGS USED IN WILDLIFE PRACTICE 35
2.8.2.8.1 Doxapram 35
2.8.2.8.2 Hyaluronidase 36
2.9 COMPLICATIONS IN WILDLIFE IMMOBILIZATIONS 36
2.9.1 STRESS 36
2.9.2 CAPTURE MYOPATHY 37
2.9.2.1 Hyperacute or Capture Shock Syndrome 38
2.9.2.2 Ataxic Myoglobinuric Syndrome 39
2.9.2.3 Ruptured Muscle Syndrome 39
2.9.2.4 Delayed-Peracute Syndrome 39
2.9.3 RENARCOTIZATION 40
2.9.4 PHYSICAL TRAUMA 40
2.9.5 HIPOXEMIA AND HIPOXIA 41
2.9.6.HYPERTERMIA/HIPOTERMIA 41
2.9.7 BLOAT 42
2.9.8 VOMIT/REGURGITATION AND ASPIRATION PNEUMONIA 42
2.9.9 EQUIPMENT FAILURE 42
CHAPTER 3. PRACTICAL CASES 44
3.2 METHODS, MATERIALS AND RESULTS 47
3.2.1 AFRICAN UNGULATES 47
3.2.1.1 AFRICAN BUFFALO 47
3.2.1.2 SABLE ANTELOPE 48
3.2.1.3 ROAN ANTELOPE 49
3.2.1.4 SPRINGBOK 49
3.2.1.5 BONTEBOK 50
3.2.1.6 GIRAFFE 50
3.2.1.7 AFRICAN ELEPHANT 51
3.2.1.8 WHITE RHINOCEROS 52
3.2.1.9 GOLDEN WILDEBEEST (Blue wildebeest) 52
3.2.1.10 PLAINS ZEBRA 53
3.2.1.11 BUSHBUCK 53
3.2.2 AFRICAN CARNIVORES 53
3.2.2.1 AFRICAN LION 53
3.2.2.2 CHEETAH 54
3.2.2.3 LEOPARD 55
3.2.2.4 AFRICAN WILD DOG (Painted hunting dog) 55
CHAPTER 4. DISCUSSION 57
CHAPTER 5. CONCLUSION 63
CHAPTER 6. BIBLIOGRAPHIC REFERENCES 66
APPENDIX 1. 84
Figure 1 – Boma used to capture a herd of zebras in 2012 (Original). 7
Figure 2 - Pneu-Dart® dart projectile components (Original). 10
Figure 3 and 4 – Best darting sites in some of the most common African large 16
ungulates (Adapted from Kock & Burroughs, 2012).
Figure 5 and 6 – Best darting sites in some of the most common African wildlife 16
species (Adapted from Kock & Burroughs, 2012).
Figure 7 and 8 – Best darting sites in some of the most common African wild 16
carnivores (Adapted from Kock & Burroughs, 2012).
Figure 9 – Helicopter chasing a white rhino in an inaccessible area to improve the 20
success of the darting. The aim of the procedure was to trim the rhino's horn due to
poaching attempts in the reserve in 2014 (Original).
Figure 10 – Herd of buffalos in a Game Farm (semi-controlled environment) 47
(Original).
Figure 11 – Free-ranging bull elephant sedated for blood collection and GnRH 51
administration (Original).
Figure 12 – Anesthetized African lion in Gondwana Game Reserve (Original). 53
Figure 13 – Castration performed by Dr Brendan Tindall on a male black leopard 55
Figure 14 – Female giraffe guided by rangers into a transportation vehicle, after the 61
reversal agent has been administered (Original).
Figure 15 and 16 – African lioness being prepared for surgery (left) and hemi- 62
hysterectomy (right) in Gondwana Game Reserve (Original).
LIST OF GRAPHICS
Graphic 1 – Percentage of individuals from different species examined during the 45
internship for veterinary procedures.
Table 1 – List of the species immobilized during the internship and their classification 2-3
by IUCN Red List of Threatened Species (Bauer, Packer, Funston, Henschel & Nowell,
2015; Blanc, 2008; Durant, Mitchell, Ipavec & Groom, 2015; Emslie, 2012; Fennessy &
Brown, 2010; Hack & Lorenzen, 2008; Henschel et al., 2008; IUCN SSC Antelope
Specialist Group, 2008a; IUCN SSC Antelope Specialist Group, 2008b; IUCN SSC
Antelope Specialist Group, 2008c; IUCN SSC Antelope Specialist Group, 2008d; IUCN
SSC Antelope Specialist Group, 2008e; IUCN SSC Antelope Specialist Group, 2008f;
IUCN SSC Antelope Specialist Group, 2008g; IUCN SSC Antelope Specialist Group,
2008h; Orrell, 2016; Woodroffe & Sillero-Zubiri, 2012; IUCN/SSC Cat Specialist Group,
Table 2 – Number of animals, divided by species, immobilized during the internship 45
(total of 245 animals and 184 chemical immobilizations).
Table 3 – Different activities performed during the internship divided according to the 46
species and the procedure.
Table 4 – African buffalo's anesthetic protocols and results. 48
Table 5 – Sable antelope's anesthetic protocols and results. 49
Table 6 – Roan antelope's anesthetic protocols and results. 49
Table 7 – Springbok's anesthetic protocols and results. 50
Table 8 – Bontebok's anesthetic protocols and results. 50
Table 9 – Giraffe's anesthetic protocols and results. 51
Table 10 – African elephant's anesthetic protocols and results. 52
Table 11 – White rhino's anesthetic protocol and results. 52
Table 12 – Golden wildebeest's anesthetic protocol and results. 52
Table 13 – Plains zebra's anesthetic protocols and results. 53
Table 14 – Bushbuck's anesthetic protocol and results. 53
Table 15 – African lion's anesthetic protocols and results. 54
Table 16 – Cheetah's anesthetic protocols and results. 54
Table 17 – Male leopard's anesthetic protocol and results. 55
Table 18 – African wild dog's anesthetic protocols and results. 56
Table 19 – Characteristics of the dart needles for the most commonly immobilized 84
African species, based on personal experience plus Dr Brendan Tindall's years of
Table 20 – Different anesthetic protocols for the African buffalo. 85
Table 21 – Different anesthetic protocols for the Sable antelope. 86
Table 22 – Different anesthetic protocols for the Roan antelope. 86
Table 23 – Different anesthetic protocols for the Springbok. 86
Table 24 – Different anesthetic protocols for the Bontebok. 87
Table 25 – Different anesthetic protocols for the Giraffe. 87
Table 26 – Different anesthetic protocols for a standing sedation of African elephants in 87
controlled environments.
Table 27 – Different anesthetic protocols for the White rhino. 88
Table 28 – Different anesthetic protocols for the Golden wildebeest (Blue wildebeest). 88
Table 29 – Different anesthetic protocols for the Plains zebra. 88
Table 30 – Different anesthetic protocols for the Bushbuck. 88
Table 31 – Different anesthetic protocols for the African lion. 89
Table 32 – Different anesthetic protocols for the Cheetah. 89
Table 33 – Different anesthetic protocols for the Leopard. 90
Table 34 – Different anesthetic protocols for the African wild dog. 90
LIST OF ABBREVIATIONS AND SYMBOLS
% Percentage
½ Half
¼ One quarter
¾ Three quarters
'' Inches
C Celsius degrees
& And
µ Mu (Receptors)
δ Delta (Receptors)
κ Kappa (Receptors)
® Registered Trade-mark
ACTH Adrenocorticotropic Hormone
NSAIDs Non-steroidal anti-inflammatory drugs
ANS Autonomic Nervous System
AST Aspartate Aminotransferase
ATP Adenosine Triphosphate
cc Cubic centimeter
CM Capture Myopathy
CNS Central Nervous System
CO2 Carbon dioxide
CR Critically Endangered
CRT Capillary Refill Time
CK Creatinine Phosphokinase
DD Data Deficient
DNA Deoxyribonucleic Acid
DOP Delta Opioid Peptide
Dr Doctor
e.g. Exempli gratia
EN Endangered
EX Extinct
EW Extinct in the Wild
FMD Foot and Mouth Disease
g Gram
GC Gel Collar
GIT Gastrointestinal Tract
GPS Global Positioning System
GnRH Gonadotropin-Releasing Hormone
HPA Hypothalamic-pituitary-adrenal
IM Intramuscular
IU International Units
IUCN International Union for Conservation of Nature
IV Intravenous
Kg Kilogram
KOP Kappa Opioid Peptide
L Liter
LAN Long-Acting Neuroleptics
LC Least Concern
LDH Lactate Dehydrogenase
m Meter
mg Milligram
mL Milliliter
mm Millimeter
MOP Mu Opioid Peptide
NE Not Evaluated
NMDA N-Metil-D-Aspartate
NT Near Threatened
PNS Peripheral Nervous System
Prof Professor
SA South Africa
SECONDS Species, Environment, Capture related factors, Other diseases, Nutrition,
Drugs, Signalment
SNS Sympathetic Nervous System
SSC Species Survival Commission
SQ Subcutaneous
TB Bovine Tuberculosis
TZ Tiletamine/Zolazepam
US Ultra-sound
VU Vulnerable
WB Wire Barb
1.1 INTERNSHIP TRAINING ACTIVITIES
This academic thesis aims to provide a short review of immobilization principles
in the wild by documenting examples of clinical cases based on data gathered during an
internship in South Africa (SA). The training period (over 1100 hours) between July 27th and
December 21st of 2015 was supervised by Dr Brendan Tindall of the Robberg Veterinary
Clinic in Plettenberg Bay. The vast majority of the training consisted of improving practical
skills in wildlife and conservation, and applying knowledge mostly acquired on previous
South African wildlife courses undertaken in recent years. However, different fields of
veterinary medicine, such as zoological medicine, exotic pets, equines, small and farm
animals also made up part of the work carried out.
SA is one of the countries where the practice of veterinary medicine in the wildlife
field is more advanced. National parks, private reserves and game farms teaming with
wildlife allow the veterinarians to develop and improve techniques used to immobilize free-
ranging animals. The veterinary practitioners involved in these procedures require a
particular knowledge and experience due to the existing biodiversity of species and the
variety of conditions to which the animals are exposed. Also, it is important to bear in mind
that most of the wild species usually subject to veterinary procedures have an important
conservational value, which raises the responsibility of the veterinarian.
Table 1 shows the different species immobilized during the internship period and
their classification according to the IUCN (International Union for Conservation of Nature)
Red List of Threatened Species. This list has several categories and criteria to classify the
species according to their relative risk of extinction. All species are categorized though some
are classified as Not Evaluated (NE) or even as Data Deficient – DD (absence of adequate
data). When the information available is reliable, the species can be properly classified as:
Least Concern (LC), Near Threatened (NT), Vulnerable (VU), Endangered (EN), and
Critically Endangered (CR) – these last three categories being for threatened species - with
the final stages being: Extinct in the Wild (EW) and Extinct (EX). (http://www.iucnredlist.org)
Table 1 – List of the species immobilized during the internship and their classification
by IUCN Red List of Threatened Species (Bauer, Packer, Funston, Henschel & Nowell, 2015;
Blanc, 2008; Durant, Mitchell, Ipavec & Groom, 2015; Emslie, 2012; Fennessy & Brown,
2010; Hack & Lorenzen, 2008; Henschel et al., 2008; IUCN SSC Antelope Specialist Group,
2008a; IUCN SSC Antelope Specialist Group, 2008b; IUCN SSC Antelope Specialist Group,
2008c; IUCN SSC Antelope Specialist Group, 2008d; IUCN SSC Antelope Specialist Group,
2008e; IUCN SSC Antelope Specialist Group, 2008f; IUCN SSC Antelope Specialist Group,
2008g; IUCN SSC Antelope Specialist Group, 2008h; Orrell, 2016; Woodroffe & Sillero-
Zubiri, 2012; IUCN/SSC Cat Specialist Group, 2015).
SPECIES IUCN RED LIST
African Buffalo Syncerus caffer, Sparrman 1779 Least Concern
African Elephant Loxodonta africana, Blumenbach 1797 Vulnerable
African Lion Panthera leo, Linnaeus 1758 Vulnerable
Blue Duiker Philantomba monticola, Thunberg 1789 Least Concern
Blue Wildebeest (Golden) Connochaetes taurinus, Burchell 1823 Least Concern
Bontebok Damaliscus pygargus pygargus, Pallas 1767 Near Threatened
Bushbuck Tragelaphus scriptus, Pallas 1766 Least Concern
Cheetah Acinonyx jubatus, Schreber 1776 Vulnerable
Leopard Panthera pardus, Linnaeus 1758 Near Threatened
Gemsbok (Golden) Oryx gazella, Linnaeus 1758 Least Concern
Giraffe Giraffa camelopardalis, Linnaeus 1758 Least Concern
Plains Zebra Equus quagga, Boddaert 1785 Least Concern
Roan Antelope Hippotragus equinus, Saint-Hilaire 1803 Least Concern
Sable Antelope Hippotragus niger, Harris 1838 Least Concern
Springbok Antidorcas marsupialis, Zimmermann 1780 Least Concern
African Wild Dog Lycaon pictus, Temminck 1820 Endangered
White Rhinoceros Ceratotherium simum, Burchell 1817 Near Threatened
Wildlife practice is a very distinctive and specialized area of veterinary medicine.
Restraining an individual might require the use of anesthetic drugs to allow a safe and
efficient immobilization before the practitioner proceeds with plans for the targeted individual.
Anesthesia plays an important role in wildlife veterinary medicine as the majority of the
procedures carried out in SA involved chemical restraint techniques.
Preventive medicine, surgery, imagiology and, obviously, anesthesia were the
key areas of the interventions carried out on the wildlife. Each time an individual was
immobilized various procedures were performed on the animal to take advantage of the
immobilization. For example, a sable antelope, immobilized for re-location purposes, was
also bio-measured, ear-tagged, implanted with a microchip, vaccinated and de-wormed. In
certain instances blood and hair collection was also performed for deoxyribonucleic acid
(DNA) testing.
This thesis aims to present chemical immobilization principles and methods used
by the practitioners in free-ranging African species, comparing the different approaches,
protocols, doses and reversals observed during the internship.
2.1 ANESTHESIOLOGY IN WILDLIFE
Wildlife veterinary medicine is closely associated with anesthesiology because
of the non-domesticated nature of the animals and the vast majority of interventions require
the anesthetic immobilization of the individuals (Hernandez, 2014; Foggin, Masterson &
Hoare, 2012; Thurmon & Short, 2007).
Originating from the Greek anaisthaesia, the word 'anesthesia' means total or
partial 'insensibility' of the body, absence of sensations (Clarke, Trim & Hall, 2014). Most
wildlife examinations are usually performed under general anesthesia to provide a safe
procedure for the animal, the veterinarian and the personnel involved (Thurmon & Short,
2007). Analgesia, also a Greek term, is used to describe the absence of pain (Clarke et al.,
2014). This status can sometimes be mistaken for tranquilization (when the animal presents
a change in its behavior, becoming more relaxed and less anxious – but yet aware – without
drowsiness) or sedation (depression and drowsiness – not aware). Local anesthesia or
analgesia is a loss of painful sensation in a specific area of the body (but yet the animal is
still apparently aware) while regional analgesia is a loss in a larger area. General anesthesia
induces a depression of the central nervous system (CNS) (Clarke et al., 2014; Thurmon &
Short, 2007).
Clinical pharmacology – the study of drugs and respective interaction in the
organism – in wildlife practice can sometimes be neglected due to the difficulties in working
with non-domesticated animals. This oversight can increase the risk of secondary effects in
the individual and jeopardize the safety of the immobilization for the animal and the
personnel (Clarke et al., 2014; Lamont & Grimm, 2014; Thurmon & Short, 2007). For this
reason, limiting stress conditions and improving monitoring and clinical care can be
advantageous (Clarke et al., 2014; Lamont & Grimm, 2014). The drug effect in the organism
is intentional but it can have adverse effects. The relationship between the dose and the
effects is called pharmacodynamics (What the Drug Does to the Body, Clarke et al., 2014)
and it is different for each individual, as is the route of administration, its absorption, the
distribution to the tissues, biotransformation and elimination from the body
(pharmacokinetics: description of the processes associated with the drug's movement –
What the Body Does to the Drug, Clarke et al., 2014) (Clarke et al., 2014; Lamont & Grimm,
2014; Thurmon & Short, 2007).
The ideal immobilizing drug for wildlife interventions must have the following
important characteristics:
- Versatility to be used in different species,
- High potency in a small volume,
- Wide margin of safety,
- Quick action,
- Reversible effects,
- Rapid elimination from the body,
- Minimal side effects,
- Calm induction and recovery,
- Minimal handling risk,
- Stability,
- Analgesic properties (Foggin et al., 2012; Muir, 2007; Thurmon & Short, 2007).
2.2 IMMOBILIZATION TECHNIQUES
Restraint techniques to immobilize wildlife are many and varied. The different
methods can be divided in two different restraint groups: physical restraint techniques and
chemical restraint techniques. Each one is better suited to particular species or situations
(Hernandez, 2014; Atkinson, Kock & Meltzer, 2012; Shury, 2014). Most of the techniques
used in free-ranging animals were extrapolated in the late 20th century from zoos, parks and
farms. Restraint techniques have become an ingenious art thanks to the experience of the
practitioners, as well as becoming safer for the animals and the people involved (Shury,
2.2.1 PHYSICAL RESTRAINT
The physical restraint of a free-ranging animal must follow a series of basic
principles that are also applied in zoo medicine: it must be safe for the people involved in the
procedure and for the animals; mortality rate and injuries should be minimal. The technique
must be simple and easily executed by non-experienced practitioners. Equipment must be
portable and easy to set up, and the procedure has to be swift enough to allow the animal to
return to its normal physiological state (Hernandez, 2014; Shury, 2014). It should also be
usable in combination with chemical restraint to improve the immobilization and the efficiency
of the performance (Atkinson et al., 2012). Other factors such as cost, season, time of the
day, species, behavior, sex and age of the animal, number of people involved and
environmental conditions must also be considered (Hernandez, 2014; Shury, 2014; Atkinson
et al., 2012; La Grange, 2012).
Free-ranging animals can be physically restrained individually, using box/cage
traps (passive capture) and net guns (using a helicopter), or en-masse by using mass
capture techniques like bomas or nets (drop net, rocket net, drive net) (Hernandez, 2014;
Shury, 2014; La Grange, 2012).
a) Physical force
Using physical restraint to immobilize wildlife is simple but can only be used on
limited species such as small mammals, birds and reptiles (Atkinson et al., 2012).
b) Passive capture
Animals are caught and trapped in an enclosure – box, cage, crate – using water
or bait to lure them. The enclosure entrance is closed manually or automatically once the
animal is inside. It is a selective method, relatively stress-free for the animal and turns out
low mortality rates. It is reusable and requires minimum staff. However, it can be very time
consuming (Hernandez, 2014; Shury, 2014; La Grange, 2012).
c) Bomas
Bomas are one of the oldest and most-used techniques in mass captures for
ungulates, especially in re-
location procedures and for
testing and research purposes
(Shury, 2014, Atkinson et al.,
2012). A physical and visual
barrier is used to drive the
animals into a desired direction,
usually into a vehicle (Atkinson
et al., 2012). The principle of this
technique is simple: the herd of
animals is driven into a funnel-
shaped enclosure guided by a
helicopter while a series of
curtains gradually draw to
behind the animals as they Figure 1 – Boma used to capture a herd of zebras in
move forward. The final curtain 2012 (Original).
corrals the herd and forces the animals directly into a transport truck using a ramp (Figure 1).
Once inside the truck, the herd is usually pole-syringed with long-acting neuroleptics (LANs).
Another option is to dart the herd in the final enclosure, load it by hand and reverse the
animals once inside (this second option is mostly performed on aggressive and valuable
animals like sable and roan antelope). This method has low mortality rates and allows for a
large number of animals to be caught in a short period with very little handling. On the down
side, this technique involves expensive equipment, is labor intensive and requires
experience. It is also weather dependent and time consuming (Fivaz & Ebedes, 2012; La
Grange, 2012).
d) Nets
Animals usually need to be driven into the nets (net-boma) by the practitioners on
foot, using horses, vehicles or aircraft like helicopters or fixed wings (Shury, 2014). Once this
has been achieved the nets collapse and entangle the animals. They are hand-restrained
and loaded with or without immobilization and/or tranquilization. This method is appropriate
for certain species (e. g. nyala - Tragelaphus angasii, or bushbuck) that inhabit thick
vegetation and often difficult terrain. The equipment is cost-efficient, easy to set up and is
less weather dependent than the boma. Nevertheless, it is labor-intensive and requires an
experienced team to manage the animals that can easily be injured and stressed. This
procedure can also be hazardous to the staff (Goodman, Hedley & Meredith, 2013; La
Grange, 2012, Fivaz & Ebedes, 2012; Bothma & Van Rooyen, 2005).
e) Net-Gun
A more recent technique, the net-gun, was developed in the 1970s in New
Zealand to capture red deer (Cervus elaphus). It soon spread to North America where it was
used to capture ungulates without using immobilizing drugs, although the technique can be
combined with chemical immobilization to extend the restraint in certain situations (Shury,
2014; Jacques et al., 2009). In cases where chemical immobilization cannot be performed
this particular gun is limited by the species to be captured and environmental conditions.
(Walzer & Gerritsmann, 2015; Goodman et al., 2013).
2.2.2 CHEMICAL RESTRAINT
Wildlife medicine veterinarians need to administer anesthetic drugs effectively
using different types of remote delivery systems (Hernandez, 2014; Arnemo, Evans,
Fahlman & Caulkett, 2014; Isaza, 2014). The practitioner selects the best system for each
situation according to the behavior and cooperation of the animal (Isaza, 2014; Atkinson et
al., 2012). The procedure chosen by the veterinarian for trained animals in zoos (controlled
environment) will differ from the procedure chosen for a free-ranging unpredictable animal
(Hernandez, 2014; Isaza, 2014, Fahlman, 2008).
Chemical restraint is used mainly for large ungulates and carnivores when
physical restraint is impractical. This chemical method is not usually appropriate for
immobilizing large numbers of animals but it is the primary choice if an individual requires
medical assistance. It is expensive and its success could depend on the experience of the
veterinarian in charge (Hernandez, 2014).
There are different approaches to administering different drugs in a chemical
restraint procedure: oral, hand-held injection, pole syringe, and using darts (Hernandez,
2014; Isaza, 2014). In cooperative animals, hand-held injections or the pole-syringed
administration are the delivery routes of choice. If the animal is uncooperative, remote
delivery systems using blow darts, gunpowder explosive darts or compressed gas projectors
are the most suitable choice for the practitioner (Isaza, 2014).
a) Oral
Oral administration is inevitably dependent on the ability to ensure the animal
accepts the drug. For this reason, its absorption and effect make it an unreliable method.
Oral administration is used mainly in carnivores or primates for sedation before darting and
requires high doses of immobilization drugs. In ruminants, because of slow absorption rates
and large mass of ingesta, this method is not commonly used (Atkinson et al., 2012;
Burroughs, Meltzer & Morkel, 2012a).
b) Hand-held Injection
This administration route is only used in animals that are already physically
restrained or cooperative in behavior. The practitioner must administer an effective rapid
intramuscular (IM) injection but must be careful to prevent accidental self-injection. The drug
is pulled into a plastic syringe with a specific needle attached (Isaza, 2014; Goodman et al.,
2013; Atkinson et al., 2012).
c) Pole Syringe
This method is used as an extension of the hand and it allows for the injection of
drugs in animals already in enclosures (crates, transport vehicles from the roof-top) or even
trees (Hernandez, 2014; Goodman et al., 2013; Kock, Jessup & Burroughs, 2012a). It follows
the same principles as the hand-held injection but has a long pole attached to the syringe,
providing a safer procedure than the hand-held option. The drug is pulled into the syringe,
which usually has a lower gauge needle. The syringe is embedded in the pole and it is ready
to use (Hernandez, 2014; Isaza, 2014; Atkinson et al., 2012). The veterinarian must
approach the animal as quietly as possible and then inject it, fast and forcefully, through the
skin into the muscle. It should always be borne in mind that the animal could still react and
cause injuries and it is therefore advisable for the injector to wait until he has the best shot
possible, because if a first attempt is missed the animal can become agitated, making it more
difficult to obtain a nice and clear second attempt (Kock et al., 2012a).
d) Blow Dart (Blow Pipe)
As its effectiveness is limited to short distances (15-25 meters [m], being more
accurate at 8-10m) the blow dart is mainly used in controlled environments like zoos, bomas,
trees or crates (Hernandez, 2014; Goodman et al., 2013; Atkinson et al., 2012; Kock et al.,
2012a) This system has a silent projection and a lower dart impact energy which results in a
softer impact to the animal, preventing possible injuries caused by the dart. It can be used on
small and large animals by blowing in one firm exhalation to propel the dart, which requires a
lot of practice (Hernandez, 2014; Atkinson et al., 2012).
e) Darts
The most common technique used for a chemical restraint in free-ranging
animals is darts fired from dart guns. The darts are very versatile and offer many varied
remote delivery systems, giving the practitioner a choice of different equipment to be used
according to the situation (darts can be used in bomas, zoos, from vehicles, from helicopters,
from horses or even by foot) from a distance of between 2m and 100m (Isaza, 2014;
Goodman et al., 2013; Atkinson et al., 2012; Kock, Jessup & Burroughs, 2012b). The first
semblance of a dart was developed in 1958, and the authors described it as a 'flying syringe'
that used an acid-base reaction to administer the drug(s) into the animal. Due to its reliability
and efficiency the remote delivery system remained unchanged for the 55 years subsequent
to its development (Walzer & Gerritsmann, 2015).
For maximum accuracy, dart guns require an experienced practitioner who is
familiar with the equipment, skilled at estimating the distance of the shot (or in using a
rangefinder), and able to calibrate the charge according to that distance (Walzer &
Gerritsmann, 2015; Hernandez, 2014).
Those adjustments need to be made quickly and quietly before the animal
moves, avoiding excessive impact to prevent accidents such as tissue damage, bone
fractures or body cavity penetrations in the darted animal (Walzer & Gerritsmann, 2015;
Hernandez, 2014; Arnemo et al., 2014; Atkinson et al., 2012). Aside a proper knowledge of
wild animals' anatomy, physiology and their behavior, the practitioner must also have enough
field experience to avoid the dangerous pitfalls associated with wildlife work (the danger of
working with carnivores is, normally, well judged but ungulates can also be potentially lethal
due to bites, kicks and injuries caused by the horns) (Hernandez, 2014).
Impact energy is the amount of
energy generated when the dart strikes the
animal – ½ dart mass x velocity2. A dart,
ideally with the minimum drug volume in the
smallest dart possible, traces an arc through
the air and drops over a distance, hitting the
animal, and hopefully providing a good IM
injection (Atkinson et al., 2012). Darts can be NEEDLE
use in animals that otherwise would be
impossible to immobilize and are less stressful DRUG
for them than the physical capture. Darts have STORAGE
low mortality rates and are safer for the
personnel involved. However, it is a time
consuming exercise (only a few animals at a
time can be targeted) that is also expensive,
and requires a very experienced veterinarian
(Isaza, 2014; Atkinson et al., 2012).
A dart can be divided in four
components: the drug(s) storage chamber, the Figure 2 - Pneu-Dart® dart projectile
Components (Original).
discharge method, the needle required to
penetrate the animal and a stabilizer for balance during the flight (Isaza, 2014). Figure 2
represents the different components of a Pneu-Dart® (Pneu-Dart, Inc., Philadelphia, USA)
dart projectile with a stabilizer colored tail.
Transmitter darts are a specific type of dart mainly used in situations where the
vegetation is dense (for nyala or a bushbuck), with the potential of the darted animal getting
lost. They can be supplied by Tele-Dart® (Tele-Dart, Westheim, Germany), Cap-Chur®
(Palmer Cap-Chur Equipment Inc., Georgia, USA), Pneu-Dart® and Dan-Inject® (Dan-Inject
Equipment, Borkop, Denmark) (Kock et al., 2012b).
2.3 TYPES OF DARTS AND DART GUNS
The chemical immobilization of wildlife usually requires remote drug delivery
systems (Hernandez, 2014). Due to the difficulties in approaching a free-ranging wild animal,
an effective and reliable remote injection system that is able to hit the animal over long
distances without causing unnecessary complications or injuries is required. The
performance of different systems is evaluated by their range and accuracy (Walzer &
Gerritsmann, 2015; Hernandez, 2014). These remote delivery systems consist of a dart and
a projector, which can be a blowpipe (blow dart), a compressed air projector or a gunpowder
cartridge rifle (powder charges) (Hernandez, 2014; Isaza, 2014; Kock et al., 2012b; Kock,
Jessup & Morkel, 2012c).
2.3.1 TWO-CHAMBERED COMPRESSED GAS DARTS (PRESSURIZED)
Two-chambered compressed gas darts (pressurized) are usually made of plastic,
with the two chambers being divided by a plunger. The front chamber is for the drug solution
and the back chamber for pressurized gas (Hernandez, 2014). Firstly, before the practitioner
even starts to fill up the dart, the system must be tested by pressurizing and depressurizing
the dart. Only then it is ready for the veterinarian to load the drug(s) into the anterior
chamber. The needle is attached, taking care to cover the needle's side hole with a silicone
sleeve (using a Leatherman® [Leatherman Tool Group Inc., Oregon, USA] to make the
needle tightly attached). Then, given that the needle's side-point is covered with a sleeve, the
operator can pressurize the posterior chamber with compressed gas/air. This is introduced
into the chamber manually by using an adapter attached to a syringe (usually a 20 ml
syringe) (Hernandez, 2014; Isaza, 2014; Kock et al., 2012b). Finally, the stabilizer – the
tailpiece – is attached to the back of the dart. When the dart hits the skin of the animal, the
silicone sleeve on the needle slides down allowing the drug to be injected into the animal
(Isaza, 2014). The pressurized gas moves the plunger forward and pushes the drug(s) into
the anterior chamber, and out through the needle hole (Hernandez, 2014). These darts are
lightweight, quiet and less traumatic for the tissues thanks to a slower injection (Hernandez,
2014; Isaza, 2014). By choosing the right needle-lengths, gauges and characteristics, this
type of dart can be used in almost every species. They are usually used for short to medium
distances with clear trajectories, in good weather conditions, from the ground or air, and
particularly in zoos or controlled environments (Isaza, 2014; Kock et al., 2012b). As these
darts are usually transparent, the veterinarian can verify that the entire volume of the dart
was injected (which make it safer when removed). However, compressed gas darts are not
recommended for all species (e.g. they are not advised for carnivores as they can easily
break the plastic dart with their teeth). Telinject® (Telinject Remote Injection Equipment,
California, USA) and Dan-Inject® are the major manufacturers of these types of darts/guns
offering a large variety of darts (volume, needle sizes, tail pieces) that can be reused after
proper cleaning and storage (Hernandez, 2014; Isaza, 2014). As the darts are reusable there
is a risk that they might fail after repeated use due to pressurizing problems or damage of the
plastic (Kock et al., 2012b).
The pressurized systems and non-pressurized explosive systems are both very
reliable, but with the former, the veterinarian requires experience and practice to guarantee a
correct pressurization of the dart (Kock et al., 2012b).
2.3.2 GUNPOWDER EXPLOSIVE DARTS
Gunpowder Explosive Darts have a gunpowder cap as a mechanism to
discharge the drug(s). A plunger is inserted to divide the anterior chamber with the drug(s)
solution from the posterior chamber with the explosive cap, a spring and a weighted firing
pin. A tailpiece is attached to the back of the dart as a stabilizer (Hernandez, 2014; Isaza,
2014). To prepare this type of dart, the practitioner has to fill the anterior chamber with the
drug(s) he is planning to use, through the needle. Needles can vary greatly in length, gauge
and characteristics. When the dart hits the animal the firing pin collides with the cap, which
results in detonation. The expanding gas moves the plunger forward and the drug(s) inject
into the animal through the needle. This is a faster mechanism in comparison to the
pressurized darts with greater chances of causing muscle trauma, particularly in small
species (Hernandez, 2014; Isaza, 2014; Kock et al., 2012b). This method can use modular
gunpowder explosive-powered reusable darts (e.g. Palmer Cap-Chur®) – one of the oldest
models and the first efficient dart manufactured in the 1960s – or prefabricated gunpowder
explosive-powered disposable darts (Pneu-Dart®) (Hernandez, 2014; Kock et al., 2012b).
The practitioner should be particularly careful when using Cap-Chur® darts as
if the charge is accidentally attached backwards it will detonate on firing and spray the drugs
out of the dart gun barrel and some of the drugs commonly used in wildlife can be lethal to
humans (Kock et al., 2012b).
The other disadvantages of these darts are the cost and the opaque plastic or
aluminum material of the darts, which prevents the practitioner from seeing if the plunger has
moved forward (Hernandez, 2014). The main difference between them is that with the
prefabricated dart model, the dart is a complete unit that cannot be dismantled. The
explosive mechanism has to be replaced by the company and cannot be reused (Hernandez,
2014; Isaza, 2014; Kock et al., 2012b). Pneu-Dart® has two different types of darts: a 'P'
type with a yellow tail (and no ridge on the tail) on the back for blowpipes, carbon dioxide
(CO2) powered projectors or air-pumped guns; and the type 'C' with an orange tail on the
back (larger than the yellow of type 'P' and with a ridge) for .22 blank-powered cartridge
projectors (Isaza, 2014; Kock et al., 2012b). In both types, the veterinarian has to select the
appropriate dart for the procedure, according to the species, choosing the length, gauge and
characteristics of the needles (Isaza, 2014). The dart must be inspected and loaded with the
drug(s). If the drug(s) does not occupy the total volume of the chamber, the practitioner must
top up the dart with sterile water or 0.9% sodium chloride (NaCl) solution to avoid a loss of
balance during the trajectory (Hernandez, 2014; Isaza, 2014). The tailpieces attached to the
back of the darts are colored to make it easier to find if the dart gets lost on the ground (Kock
et al., 2012b).
Besides the darts mentioned, there are more types, associated with a large
variety of different dart guns: the Aluminum Two-Chambered Compressed Gas Darts for
Palmer Cap-Chur®, the Chemical-Powered Darts (with and acid-base reaction which results
in a gas production to expel the dart's content), the Spring-Powered Darts or even Solid Drug
Darts (Isaza, 2014).
Additionally to the type of dart or remote delivery system used, it is always good
practice to mark the syringes during the procedures (Kock et al., 2012b).
2.3.3 REMOTE DELIVERY SYSTEMS
Usually used in controlled environments such as zoos or on confined animals,
blowpipes and gauged blowgun projectors are two of the most common compressed gas
projectors that allow a short distance shot. The blowgun system is very similar to the
blowpipe but it has an external compressed gas source, eliminating the need for the
practitioner to exhale through the pipe. One of the most commonly-used blowguns is
manufactured by Dan-Inject® (Isaza, 2014).
Air or CO2 rifles and pistols are commonly used in free-ranging wildlife. The
compressed gas such as CO2, comes in cylinders or in air pumps and the veterinarian has to
select the amount of gas to use each time he wants to dart an animal. Palmer Cap-Chur®
and Pneu-Dart® type 'P' are the leading manufactures of these projectors. These darts allow
for very accurate long-distance shots. However, the dart can cause a significant impact on
the animal with traumatic consequences (Isaza, 2014; Kock et al., 2012c).
The Pneu-Dart® type 'P' uses a CO2 cartridge or a reusable CO2 cylinder and the
practitioner can easily control the amount of gas he is using according to the distance
between him and the animal. The latest model of Pneu-Dart® type 'P' is simple, durable and
relatively inexpensive, allowing the user to dart accurately, quietly and quickly. The weight of
this heavy gun can be a problem in the field. However, the Dan-Inject® model, also available
with CO2 cylinders, is lighter and yet robust; the downside to this gun is the unreliability on
long distance shots (optimal range between 10-40m) or when the veterinarian is darting from
a helicopter (Kock et al., 2012c).
Gunpowder cartridge-powered rifles are supplied with .22 caliber blanks of
different strengths worked according to color. They are highly accurate and can be used by
the practitioner over long distances even in windy conditions. The most common
manufacturers are Palmer Cap-Chur® and Pneu-Dart® type 'C' (Isaza, 2014).
The Pneu-Dart® type 'C' models are supplied with powder charges of different
strengths, represented by a different color. Brown charges are the lightest, followed by
green, yellow and finally red, the strongest one. After loading the charges in the gun extra
distance control can be achieved by altering the port adapter in the dart discharge system
(Kock et al., 2012c).
Additional dart equipment includes laser range finders, scopes, sights and
binoculars. These are vital in calculating the appropriate pressure and ensuring accurate dart
placement (Isaza, 2014; Kock et al., 2012c). The sights are particularly useful when darting
from the ground in thick bush areas or from a helicopter, whereas telescopic sights are used
most in accurate long-distance shots. Some practitioners prefer the red dot point scope,
where both eyes are kept open during darting (which is advantageous in situations where
dangerous animals are in the area) for both short and long distances, either darting from a
helicopter or even at night (commonly when carnivores are being baited). A laser sight can
also be an option for highly accurate night darting (Kock et al., 2012c).
Regardless the type of remote darting equipment used, the practitioner must
always be familiar with the equipment and practice with it. Experience and practice are the
key to a good immobilization procedure (Kock et al., 2012b).
2.4 DARTING SITES
Large muscle masses are the ideal darting sites to allow a correct IM
administration. The hindquarters are, in most of the species, the best place to target.
However, because of adipose tissue, in some species the base of the neck or the triceps
muscle are more suitable (Hernandez, 2014).
Base of the neck region: This area is suitable for animals with heavy necks such
as the rhino, the hippopotamus (Hippopotamus amphibious) or large antelopes like the kudu
(Tragelaphus strepsiceros) causing little muscle trauma. The veterinarian must be aware that
the dart could strike a cervical vertebra, vital blood vessels or the nuchal ligament. It can also
be quite irritating for the animal (Hernandez, 2014; Arnemo et al., 2014; Burroughs et al.,
2012b).
Triceps muscle region: This is a suitable area for most species. In thin animals
there is a risk of the dart striking the scapula cartilage, the scapula spine or vital blood
vessels in the neck. The biceps and triceps are easy to dart from the ground, particularly in
animals like the rhino or the elephant. If the animal is too thin, its hindquarters are the
preferable area for darting (Hernandez, 2014; Atkinson et al., 2012).
Hindquarters: With the most muscle mass for a large target area, the hindquarters
is the most common and desirable darting site (Hernandez, 2014). It has only a few vital
structures (although care must be taken to avoid penetrating the femur and pelvis if the
animal is struck on the medial side of the hind leg) and it can be accessed either from the
ground or a helicopter. The veterinarian must dart the animal perpendicularly to the surface
to ensure a deep IM injection. If the practitioner is darting the animal from behind, he must
take care to avoid the soft tissue area of the perineum, which can cause problems,
particularly in equines (Atkinson et al., 2012).
Withers and rump: These spots can be a small target when the practitioner aims
from the side or from above (using a helicopter) but usually represent a good muscle mass,
especially in the eland - Taurotragus oryx and the rhino (Atkinson et al., 2012; McTaggart,
Kock & Hofmeyr, 2012).
Chest: Due to the risk of hitting the head or penetrating the thoracic cavity,
causing pneumothorax, the chest is not a common place to dart an animal. The pectoral area
is an option for darting if no alternative is available. Rhino, buffalo, giraffe or eland are
suitable for pectoral darting (Atkinson et al., 2012; Kock, La Grange & du Toit, 1990).
The following figures (3,4,5,6,7 and 8) represent the different darting sites in
some of the most common African wildlife species.
Figure 3 and 4 – Best darting sites in some of the most common African large
Figure 5 and 6 – Best darting sites in some of the most common African
wildlife species (Adapted from Kock & Burroughs, 2012).
Figure 7 and 8 – Best darting sites in some of the most common African wild
2.5 NEEDLE LENGHTS, GAUGES AND CHARACTERISTICS
There are numerous types of darts, each one with a specific volume (cubic centimeter, cc),
needle length (inches, ''), needle gauge and with characteristics that match their purpose.
The length of a needle is very important to a successful immobilization and avoiding
unnecessary injury or pain to the animal. The practitioner also needs to bear in mind the
gauge of the needle and characteristics such as collars or barbs (Hernandez, 2014; Atkinson
et al., 2012). However, the practitioner's decision will also be influenced by the choice of
drug(s). If the dart is to be filled with antibiotics (which have a high viscosity), a lower gauge
needle is required (Atkinson et al., 2012).
The length of the needle is usually associated with the size of the animal to be
darted, the thickness of the skin and the depth of the muscles. For an effective darting the
drug(s) must be injected in the deep muscles, which requires an accurate dart placement
(Hernandez, 2014). Logically, longer needles will provide a better dart attachment within the
animal and, if the practitioner wants to make sure that the dart is going to stay firmly
embedded in the animal, he can opt to use barbed needles. A barbed dart can stay attached
to the animal long enough for the practitioner to recover it while the animal is immobilized.
However, the barb can cause more muscle damage than plain or collared darts. The
practitioner has to extract the barbs correctly to avoid further trauma (Atkinson et al., 2012).
These types of darts can be used in antelopes, buffalos or rhinos. A drop-out dart, with
collared needles, preferably the gelatinous absorbable type, is usually used by the
practitioner when darting an animal for vaccination (Atkinson et al., 2012).
Table 19 (Appendix 1.) shows the best dart characteristics according to the
information collected from the clinical cases during the internship and based on the
experience of Dr Brendan Tindall (unpublished data).
2.6 FACTORS THAT INFLUENCE A CAPTURE
Most of the free-ranging wildlife in SA has to be immobilized in a capture
procedure. Various factors can affect that immobilization: species, environment, capture-
related factors, other diseases, nutritional status, drugs used to immobilize the animal and
administration route, and signalment. The veterinarian in charge must evaluate these seven
categories of predisposing factors with the mnemonic SECONDS on every procedure
(Arnemo et al., 2014; Hernandez, 2014; Shury, 2014; Atkinson et al., 2012; La Grange,
Different species have different responses to the same drug and/or dose. Tamed
individuals or individuals in a controlled environment may have a better response to a lower
dose than free-ranging animals (Atkinson et al., 2012). Environmental factors (e.g. humidity,
high temperatures, terrain) are also an important consideration. During the induction phase
animals in areas inaccessible to vehicles can become lost or injured (e.g. falling from a cliff
or drowning) and should not be immobilized (Arnemo et al., 2014; Ko & Krimins, 2014;
Shury, 2014; Atkinson et al., 2012). Weather conditions (e.g. wind, temperature and time of
day) must also be considered when planning an immobilization (Arnemo et al., 2014;
Hernandez, 2014; Shury, 2014). Crosswinds might change the dart's trajectory; procedures
during the late afternoon could result in an animal becoming lost in the dark; high
temperatures could cause hyperthermia in animals, and low temperatures could result in
hypothermia, which can both be fatal. The habitat must also be evaluated. Areas where the
bush is too thick might be a problem to dart and find the animal (Arnemo et al., 2014;
Hernandez, 2014; Ko & Krimins, 2014; Atkinson et al., 2012; Meltzer & Kock, 2012).
Capture-related factors such as the techniques, handling methods, transportation, position of
the animal or injuries during the procedure can play a role, as well as other diseases
(underlying infectious diseases, parasites, anemia or organ damage), which can predispose
the individual to clinical problems during the immobilization. The nutritional status of the
animals can be related to dehydration, pre-existent vitamin E and selenium deficiency or
other minerals and vitamins or poor body condition (e.g. season related). These different
conditions can interfere with the planned anesthetic protocol. The drugs used depend on the
species to immobilize and it falls to the veterinarian to choose the most adequate
combination suitable for the situation. An opioid combination with tranquilizers or sedatives is
usually used in ungulates, while in carnivores it is preferable to use combinations of
cyclohexylamine and sedatives. These combinations have different effects on each species
and each individual (Burroughs et al., 2012a; Foggin et al., 2012). The route of administration
must follow manufacturers' pharmacological instructions and recommendations to improve
the immobilization and avoid problems (e. g. drugs formulated with oils being administered
intravenously) (Atkinson et al., 2012; Burroughs et al., 2012b). When the veterinarian uses a
pole syringe or dart to immobilize the animal, the best route of administration is IM, although
in cases the dart or syringe can embed subcutaneously. The best IM administration sites are
in the hindquarters, shoulder or neck and are dependent on the species and the line of sight.
Depending on the drug of choice the intravenous (IV) route can be preferable for topping up
an animal that is starting to wake from anesthesia, as well as for the administration of a fast
reversal drug (Atkinson et al., 2012; Burroughs et al., 2012a; Burroughs et al., 2012b; Kock
et al., 2012a). Finally, signalment has to be taken into account (e.g. sex – some studies
affirm that males have a higher risk of developing severe complications such as capture
myopathy, and estrogens have a protective effect in females; age; pregnancy) (Blumstein et
al., 2015; Wolfe, 2015; Hernandez, 2014; Pas, 2014; Paterson, 2014; Hofmeyr, Fivaz &
Meltzer, 2012; Atkinson et al., 2012; Sanchez, 2011).
2.7 PHASES OF IMMOBILIZATION
To avoid complications, the practitioner must follow this sequence of events:
Planning and preparation phase, Approach phase, Induction phase, Handling and monitoring
phase, Reversal/Recovery phase and the Reflection phase (Arnemo et al, 2014; Shury,
2.7.1 PLANNING AND PREPARATION
The main goal of this phase is to organize the actions that the operation requires,
according to the purpose of the immobilization. An adequate plan of action must be
established and the entire team informed (Arnemo et al., 2014; Hernandez, 2014). The
veterinarian in charge must prepare the drugs, the equipment and give instructions to the
personnel, including human safety issues to avoid major complications during the
immobilization. He has an ethical responsibility towards the animal and the choice of what is
best must be elaborated also in accordance with the animal's behavior and its health
status/body condition, always bearing in mind the specific individual's sensitivity (Hernandez,
2014; Atkinson et al., 2012; La Grange, 2012).
As previously mentioned, the terrain is an important factor in capturing free-
ranging species. The practitioner or a member of the team must be familiar with the terrain
and its conditions. Flat or open areas might lead the veterinarian to plan a slower induction
whereas thick wooded areas must be associated with a faster induction or the use of
telemetry/drones to avoid losing the animal. The same can occur if the weather is
excessively hot and humid, which can force the procedure to be rescheduled for the benefit
of the animal. The veterinarian must be familiar with the drugs available and abide by the
principle that it is preferable to overdose than to underdose (Arnemo et al., 2014; Ko &
Krimins, 2014; Burroughs et al., 2012a; La Grange, 2012).
A crucial part of the planning phase is the choice of equipment (Hernandez,
2014). Besides the drugs' delivery systems, drugs and eventual medication, blindfolds,
earplugs, water, ropes, stretchers and other equipment must be readily available during the
immobilization procedure (Arnemo et al., 2014; Goodman et al., 2013; Atkinson et al., 2012;
Kock, 2012).
2.7.2 APPROACH
The approach and the darting are the two most important actions during the
immobilization process. Although a successful approach and a successful darting improve
the procedure, its success is not guaranteed (Atkinson et al., 2012). This phase is more
important in free-ranging wildlife than in zoo immobilizations as some of the wild species can
become excited and run, sometimes for long distances before the intended effect of the
drugs kicks in (Arnemo et al., 2014; Atkinson et al., 2012). Whenever possible the
veterinarian should avoid darting on foot or using vehicles that the animals are not familiar
with. He should stay on the road; avoid disturbing the bushes or changing the shape of the
vehicles well as using the minimum people required. Patience is essential and chasing
animals will increase their stress levels as well as the difficulty of the approach and the
darting (Atkinson et al., 2012). The darting must be done when the animal is not moving, at a
maximum range of 40-50m (except if the darting is taking place from an helicopter, where the
ideal scenario is to have the helicopter and the animal moving at the same speed) (Atkinson
et al., 2012; McTaggart et al., 2012). If there is a high risk of losing the animal, the use of a
helicopter could be the best choice for the darting. Helicopters are a very efficient form of
approach, particularly if the terrain does not allow a ground approach (Figure 9); although
transmitter darts might be an option if no helicopter is available (Atkinson et al., 2012; Kock
et al., 2012b; McTaggart et al., 2012). The best approach after the darting is to leave the
animal undisturbed, if possible, otherwise, if it runs, a chase will be necessary to keep it in
sight (Atkinson et al., 2012).
Figure 9 – Helicopter chasing a white rhino in an inaccessible area to improve the success
of darting. The aim of the procedure was to trim the rhino's horn due to poaching atempts in
the reserve in 2014 (Original).
2.7.3 INDUCTION
The induction time is defined as the time between the injection/darting and the
effective immobilization of the animal (Atkinson et al., 2012). The veterinarian has to be
aware of the variable factors (drugs, specie, stress, dart, dosage) that can influence the
induction phase, which usually takes between 3-15 minutes (Hernandez, 2014). A shorter
induction time might indicate that the animal was overdosed, whereas a longer induction time
can suggest an underdosage and the animal might require extra immobilization drugs
(always bear in mind that it is better go higher with the dose, especially with opioids –
Burroughs et al., 2012a). First, the animal will usually experience a stage of alarm, followed
by a wariness, then disorientation/agitation and possible behavior changes, mainly in
locomotion such as running and ataxia. This could cause the animal to stumble or move
away from the herd, possibly towards vehicles or trees, and loss of balance. An
inexperienced veterinarian might find it harder to notice such changes. This phase is followed
by a phase of excitement ("hackney gait") and then, if the quantity of the dose suffices, the
animal loses coordination and falls into recumbency. If the dosage is insufficient the animal
will take longer to reach recumbency, and might in fact never reach this stage, experiencing
instead a prolonged stage of excitement which increases the risk of injury and hyperthermia,
exhaustion and, consequently, can lead to death (Ko & Krimins, 2014; Radcliffe & Morkel,
2014; Atkinson et al., 2012). The veterinarian must be particularly aware of the animals'
recumbency as for example, ruminants normally have to be in a sternal position to avoid
bloat, and species like carnivores must have their eyes covered with blindfolds (Arnemo et
al., 2014; Hernandez, 2014). Additionally, the practitioner and the team involved must be
acquainted with the terrain and able to anticipate the behavior of the animal and the potential
risks (Hernandez, 2014). The use of vehicles and helicopters to guide the animal to a safe
area and monitoring it with binoculars without getting close might be an option (Arnemo et
al., 2014; Hernandez, 2014; Ozeki & Caulkett, 2014; McTaggart et al., 2012).
2.7.4 HANDLING AND MONITORING
Anesthesia affects all animals differently: they can be found on their feet but
handleable; lying down, although they still get up when approached; down with spontaneous
movement of the head; down with no voluntary movement (caution); down with poor
breathing (danger).
The veterinarian must take care when approaching the animal and should move it
into a correct position if necessary (e.g. herbivores in a sternal position with the nose down;
elephants in a lateral position) as well as evaluate the depth of the anesthesia. If necessary
he should cover the animals' eyes with blindfolds and use earplugs to block the noise.
Carnivores may need their eyes lubricated before being blindfolded (Hernandez, 2014;
Atkinson et al., 2012; Hofmeyr et al., 2012). Monitoring is essential and basic references
such as body temperature, respiratory rate, heart rate, and capillary refill time (CRF) should
be recorded at least every 5-10 minutes. Major anesthetic complications can be anticipated
by clinical signs such as depression, weakness, abnormal body temperature, cold
extremities, abnormal heart rate and/or respiratory rate, cyanotic mucous membranes, slow
capillary refill time, tremors, neurological signs, pain, and lameness. A pulse oximeter might
also be a useful tool to have in the field (Hernandez, 2014; Ozeki & Caulkett, 2014; Atkinson
et al., 2012; Hofmeyr et al., 2012). At the same time, the dart must be collected and safely
discarded to avoid accidents. The dart wound must be inspected and treated for trauma
injuries (Hernandez, 2014; Atkinson et al., 2012). Some of the most common complications
that can occur during wildlife capture procedures will be explained in detail later on
(Hernandez, 2014; Ozeki & Caulkett, 2014). The veterinarian can also administer: wound
treatment to the dart injury (e.g. penicillin), other antibiotics, vitamins, anti-inflammatory
drugs, eye ointments, analgesics, cleaning solutions and, if necessary, initiate
emergency/reversal (or partial reversal) drugs (Atkinson et al., 2012; Hofmeyr et al., 2012;
Kock et al., 2012a). If necessary, the team must be prepared to cool the animal down to
avoid any possible consequences of hyperthermia, and if the temperature does not go down
despite that cooling, the practitioner might need to administer antidotes to antagonize the
effects of the anesthesia (Hernandez, 2014; Ko & Krimins, 2014).
2.7.5 REVERSAL/RECOVERY
The recovery of an animal will depend on the procedure, the anesthetic protocol
and the species. The reversal administered by the veterinarian might be a total or a partial
antagonist of the main drug(s) used (Arnemo et al., 2014; Hernandez, 2014). For example,
Ketamine is a dissociative that cannot be reversed but, because it is usually combined with
α-2-agonists like Medetomidine, the practitioner can use an antidote to reverse the sedative
(Hernandez, 2014; Burroughs et al., 2012a). A partial reversal might be necessary to load
the animal in some cases, or to improve safety conditions for the people involved, who must
be distanced from the animal when the practitioner completely reverses the anesthesia
(Hernandez, 2014; Atkinson et al., 2012). The reversal(s) should be given intravenously, but
in some cases, such as with carnivores, an IM injection can be a better option. Then the
blindfold and earplugs must be removed and all the equipment must be relocated to a safe
area (Arnemo et al., 2014; Hernandez, 2014; Atkinson et al., 2012; Burroughs, Hofmeyr et
al., 2012b). The recovery environment has to be assessed: to ensure a calm recovery, the
spot cannot be exposed to sunlight, to wind or other animals, and, if possible, the practitioner
must observe the animal from a safe distance until it is fully recovered, able to walk, protect
itself from other animals, and avoid hazards (Arnemo et al., 2014; Hernandez, 2014).
2.7.6 REFLECTION
After every immobilization procedure the practitioner should take some 'reflection'
time to log all the doses and drugs used, the methods he decided to use, any mistakes made
and any improvements he could adopt in future immobilization processes. Some practitioners
use data capture sheets to ensure all details are recorded properly, mainly for research
purposes (Goodman et al., 2013).
2.8 WILDLIFE PRACTICE APPLIED PHARMACOLOGY
Different wildlife species have different reactions to the restraint method chosen
by the practitioner (Hernandez, 2014; Fahlman, 2008). Consequently, these methods have to
be adapted to the species and its behavior in order to cause as little stress as possible and
minimal physiological disturbance, safeguarding the welfare of the immobilized animal
(Hernandez, 2014; Fowler, 1995). Most of the procedures in wildlife practice require the
individuals to be chemically immobilized, which in itself represents challenges and risks for
the practitioner involved (Hernandez, 2014; Schumacher, 2008). Over the years the drugs
available on the market have improved, allowing the practitioners to adopt chemical restraint
methods over physical, or in association with the latter (Swan, 1993). Unfortunately, there is
no perfect drug available with all the characteristics of an ideal anesthetic and the
veterinarian has to choose the best drug(s) according to the situation with which he is faced
with (Muir, 2007). For example, the drug of choice for one particular species might be
inappropriate for another. Opioids, frequently used to immobilize ungulates, can cause
several problems, such as respiratory depression in primates, or excitement in felids
(Fahlman, 2008). The practitioner has to be familiar with the characteristics of every drug
available and be versed in using it, not forgetting that free-ranging animals usually require
higher doses than individuals in a controlled environment, as mentioned earlier (Atkinson et
al., 2012; Fahlman, 2008; Muir, 2007). Initially, in the earliest chemical immobilizations,
practitioners took advantage of the properties of the neuromuscular blockers and used it in
the restraints by paralyzing the skeletal muscle, but the animals were still aware and
conscious, increasing stress levels and resulting in high mortality rates (Caulkett & Arnemo,
2007). Nowadays, thanks to improved knowledge, veterinarians are using drugs that cause
total or partial depression of the CNS, combining different drugs with synergetic effects in
order to decrease doses and reduce side effects on the animal (Fahlman, 2008;
Schumacher, 2008). Because of all the variables involved in free-ranging wildlife practice
there is no ideal protocol for immobilization procedures. Nowadays, however, the main aim of
these operations is to minimize the morbidity and mortality of the individuals by decreasing
the excitement-free induction period, ensuring proper muscle relaxation, adequate analgesia,
decreasing the cardiopulmonary depression effects and ending the immobilization with a
rapid and smooth recovery after the administration of the reversal drug(s) (Schumacher,
2.8.2 IMMOBILIZATION AGENTS
2.8.2.1 OPIOIDS
This group of drug is probably the most important in the chemical immobilization
of ungulates. Nevertheless, the opioids family also confers analgesic effects on the
management of pain in many different species through specific opioid receptors (Lamont &
Grimm, 2014; Ramsay, 2008; Schumacher, 2008). There are different types of opioid
receptors: mu (µ) receptors – mu opioid peptide (MOP), delta (δ) receptors – delta opioid
peptide (DOP) and kappa (κ) receptors – kappa opioid peptide (KOP) (Lamont & Grimm,
2014; Ramsay, 2008). The µ receptors are associated with analgesic properties and some
side effects associated with opioid administration. On the other hand, δ receptors have minor
analgesic effects but they are able to modify other receptors; the κ receptors are associated
with the analgesia in some specific areas on the central and peripheral nervous system
(PNS). Some natural molecules produced by the organism in the CNS, in the adrenal gland
or even in the hypophysis are endogenous opioid peptides, which couple in the endogenous
opioid receptor and mediate some analgesia. Exogenous systemic administration of opioids
(IV, IM or SQ) and endogenous opioids act on the CNS receptors. Opioids can cause
depression of the CNS, leading to sedation, or stimulation of the CNS and, consequently,
excitement, depending on the species and the dose that is administered (Lamont & Grimm,
2014). Opioids can also affect the thermoregulation center, causing hypothermia (or
hyperthermia in some species); the emetic center, by causing nausea and vomiting because
of the stimulation of the chemoreceptor trigger zone; they can also cause suppression of the
cough center and changes in the pupillary diameter – mydriasis in case of excitement. The
respiratory system is also affected by a dose-dependent depression of ventilation
(bradypnea), and at a cardiovascular level, bradycardia can occur as well as arterial
hypertension (or hypotension in some cases). Ruminants might be predisposed to
gastrointestinal complications associated with the administration of opioids (e.g. bloat) or
problems often associated with myopathies (Wolfe, 2015; Lamont & Grimm, 2014;
Schumacher, 2008). These complications and the undesirable effects are often observed
when the opioids are administered alone, without any synergetic drugs associated to allow
for a lower dose of the opioid agent and, consequently, fewer side effects on the animal
(Schumacher, 2008).
The opioids used in veterinary practice are usually morphine derivatives which
react with endorphin receptors such as µ, δ and κ receptors in the brain, spinal cord,
autonomic nervous system (ANS), mesenteric plexus of the gastrointestinal tract (GIT),
cardiovascular system and kidneys, affecting mainly pain, behavior, and voluntary muscle
and GIT motility (Lamont & Grimm, 2014). The most common effects are sedation,
excitement, ataxia, analgesia, respiratory depression, hyper or hypotension, hypothermia
and inhibition of the GIT motility. Drugs available on the market can act as opioid agonists,
antagonists or partial agonists/partial antagonists and they can have a large range of effects
on the animal (Lamont & Grimm, 2014; Ramsay, 2008).
2.8.2.1.1 Etorphine hydrochloride
This drug is probably the most widely used immobilization drug for African
ungulates, preferably reversed with diprenorphine hydrochloride (Janssen & Allen, 2015;
Burroughs et al., 2012a; Schumacher, 2008). Due to its sedative properties etorphine is
mainly used to restrain wildlife (it is rarely used as an analgesic) (Lamont & Grimm, 2014). It
has a quick onset, starting with a slight ataxia, which progresses to an evident ataxic event,
high stepping gait can be visualized and some of the animals might run while others might
stand until falling into a recumbency position – sternal or lateral (Wolfe, 2015; Lamont &
Grimm, 2014; Burroughs et al., 2012a). The doses are calculated according to the species
not the body mass of the animal (Wolfe, 2015; Burroughs et al., 2012a; Atkinson et al., 2012;
La Grange, 2012). To reduce side effects, improve the induction time and avoid hypertonicity
it is mainly used in combination with a tranquilizer (e.g. butyrophenone) or a sedative (e.g. α-
2-adrenergic-agonists) (Lamont & Grimm, 2014; Burroughs et al., 2012a; Fowler, 2008;
Schumacher, 2008). The reversal drug – diprenorphine, naltrexone, naloxone – must be
given as quickly as possible to avoid the side effects of the etorphine: respiratory depression,
which can lead to severe hypoxia, acidosis and death; cardiovascular changes (initially it
starts with hypertension and changes to hypotension); excitement; hypertonicity;
hyperthermia; convulsions; GIT stasis, which can result in bloat, regurgitation, aspiration
pneumonia (ruminants require a sternal recumbency, head up with the nose down)
(Burroughs et al., 2012a; Schumacher, 2008; Caulkett & Arnemo, 2007; Nielsen, 1999). The
induction time of Etorphine is longer when compared to other opioid agents. However, the
recovery takes about 1-3 minutes after IV injection of the reversal agent (or 5-10 minutes
when the practitioner opts for an IM administration) (Schumacher, 2008; Caulkett & Arnemo,
2.8.2.1.2 Thiafentanil oxalate
Thiafentanil is a synthetic opioid like etorphine (Burroughs et al., 2012a). It has a
similar potency and similar properties when compared to carfentanil or even etorphine, but a
shorter induction time, allowing a faster immobilization of herbivores, fewer cardiopulmonary
depression effects and a shorter half-life (Burroughs et al., 2012a; Schumacher, 2008).
When the practitioner chooses to use thiafentanil, the chances of a complication called
renarcotization are lower than with etorphine or carfentanil because of the short time of
action of these drugs, allowing for a proper anesthetic reversal with an antidote (usually
naltrexone) (Burroughs et al., 2012a; Schumacher, 2008; Allen, 1996). It is used in species
with a higher sensitivity to etorphine (e.g. sable antelope) or even as a combination of both
opioids (e.g. giraffe) (Lamont & Grimm, 2014; Lance & Kenny, 2012; Burroughs et al., 2012a;
Burroughs et al., 2012b; Janssen et al., 1991).
2.8.2.1.3 Carfentanil
Carfentanil is no longer used in SA as a registered product but it is still available
in the United States of America (USA) (Burroughs et al., 2012a; Caulkett & Arnemo, 2007). It
is a derivative of fentanyl and it is more potent than etorphine, with a faster onset of action
(induction time around 2-5 minutes) but a longer duration of action (Lamont & Grimm, 2014;
Fowler, 2008; Schumacher, 2008). It is not completely reversible with diprenorphine, not
even the side effects on the CNS. Naltrexone is the most efficient reversal, in a ratio dose of
carfentanil 90:1 or 100:1 (Lamont & Grimm, 2014; Fowler, 2008; Caulkett & Arnemo, 2007).
2.8.2.2 OPIOID ANTAGONISTS
The biggest advantage of using opioids in wildlife practice lies in its reversible
properties with the proper antagonist (Ramsay, 2008; Caulkett & Arnemo, 2007; Nielsen,
1999). Opioid antagonists are used to reverse the effects of opioids by binding with opioid
receptors in the body. When administered in high doses, or because they have a better
affinity to the receptors, these antagonists compete with the opioids and replace them at the
receptors level. Some antagonists can also be used to not completely reverse the animal but,
for example, to lighten the immobilization and reduce the respiratory depression or other side
effects. They can be divided in two groups: the mixed antagonists, with a slight agonistic
effect (diprenorphine, butorphanol, nalbuphine, nalorphine) also known as partial
antagonists/agonists or agonist-antagonists; and the pure antagonists (naltrexone,
naloxone), depending on the relative concentration between agonistic and antagonistic
effects (Lamont & Grimm, 2014; Burroughs et al., 2012a). Partial antagonists compete for the
µ receptors, where they have an antagonistic effect, but they act as agonists for the κ
receptors. These drugs were developed for acting as analgesics with fewer respiratory
depression effects. Contrarily, pure opioid antagonists have an affinity for both µ and κ
receptors (Lamont & Grimm, 2014).
2.8.2.2.1 Butorphanol tartrate
This synthetic partial antagonist interacts with the κ receptors as an agonist
opioid, inducing analgesic effects, and as an antagonist on the µ receptors, reversing the
effect of some of the most potent opioids (Lamont & Grimm, 2014; Burroughs et al., 2012a;
Ramsay, 2008). It is commonly administered in combination with other drugs (e.g. azaperone
or medetomidine) and its properties allow it to be used in standing sedations, for
walking/transporting rhinos and to improve breathing in deep immobilization cases with
opioids by manipulating the level of anesthesia (Miller & Buss, 2015; Radcliffe & Morkel,
2014; Burroughs et al., 2012a).
2.8.2.2.2 Diprenorphine
This partial antagonist is the most commonly used antidote for etorphine (can be
used to reverse thiafentanil but it is less effective) (Burroughs et al., 2012a). At lower doses,
it has antagonistic effects in the animal (e.g. improving the respiration after the IV
administration) but in higher doses (ratio of diprenorphine 10:1) or if repeated, it has
agonistic effects. The doses are calculated according to the opioid used for the
immobilization: 3 times (x) the dose of the etorphine used for large animals or 2x the dose for
the rest of the ungulates, and it can be administered by IV or IM injection (Lamont & Grimm,
2.8.2.2.3 Nalbuphine & Nalophine
The characteristics of these two drugs are very similar to butorphanol.
Nalbuphine is considered a very similar version of the previous drug used by veterinarians,
nalophine (Lamont & Grimm, 2014; Burroughs et al., 2012a). They are used to improve
animals' breathing in a deep anesthetic stage and in procedures that require walking rhinos
(Burroughs et al., 2012a).
2.8.2.2.4 Naltrexone
Naltrexone is used to antagonize the action of the opioids on the µ receptors.
Contrary to the other previously mentioned antagonists, this is a pure antagonist, with a long
half-life (no/lower renarcotization risk) and it can be used in human opioid overdoses
(Burroughs et al., 2012a). Naltrexone is able to act as an antidote to all the effects of the
different opioids (including the renarcotization with carfentanil), improving the animal's
response, the perception of pain and awareness. Once administered, the animal cannot be
re-immobilized in the next 24 hours (Lamont & Grimm, 2014; Burroughs et al., 2012a;
Caulkett & Arnemo, 2007; Allen, 1996). It can be administered by IM or IV injections
(Caulkett & Arnemo, 2007). The ratio to be used is 10:1 although some authors refer 20 mg
of naltrexone per mg of etorphine (Burroughs et al., 2012a) or 100mg per mg of carfentanil
(Fowler, 2008; Nielsen, 1999). In white rhinos, naltrexone must be always given when the
animal is released into the field (Burroughs et al., 2012a).
2.8.2.2.5 Naloxone
This drug is also a pure antagonist and it has been the first choice for human
intoxications with opioids (Naltrexone can also be used). Because of its short action, the
opioid used might still have an effect on the animal immobilized (Caulkett & Shury, 2014;
Morkel & Kock, 2012; Burroughs et al., 2012a). Consequently, animals immobilized with
opioids and reversed with naloxone must be monitored to ensure renarcotization does not
occur. Naloxone can reverse the effects of all opioids and increase the sense of reaction,
awareness, perception and pain (Lamont & Grimm, 2014; Caulkett & Shury, 2014).
2.8.2.3 CYCLOHEXYLAMINES
The cyclohexylamine group is also known as the dissociative anesthetics used as
knock-down drugs in wildlife practice. There are no antidotes to reverse the effect of the
dissociative agents and usually an animal immobilized with a drug of this category requires a
longer recovery. Like opioids, cyclohexylamine can be combined with a tranquilizer or a
sedative (which can be reversible) to attenuate its side effects, such as the hypertonia of the
skeletal muscle (Burroughs et al., 2012a; Thurmon & Short, 2007). It has a cataleptic effect,
analgesic properties (short duration) and cause immobility, loss of consciousness and
amnesia (Lamont & Grimm, 2014; Burroughs et al., 2012a; Nielsen, 1999). Even when
knocked-down, the animals still maintain the pharyngeal and laryngeal reflexes, as well as
the palpebral and corneal reflexes, the tongue might move and the eyelids remain open (it is
advised to protect the cornea from the light with eye drops or ophthalmic gel), spontaneous
movements of the animals, mydriasis or nystagmus occur (Burroughs et al., 2012a; Thurmon
& Short, 2007). Dissociative agents can be used to immobilize carnivores, primates, reptiles
or birds without severe effects on the respiratory or cardiovascular systems, even in high
doses (Burroughs et al., 2012a; Caulkett & Arnemo, 2007; Nielsen, 1999). Ketamine and
tiletamine are the most common types of cyclohexylamine used in wildlife practice, which can
be administered as an IM or an IV injection (Lamont & Grimm, 2014; Burroughs et al.,
2012a). The induction takes about 5-10 minutes and some of the side effects may include
convulsions (mainly if overdosed and, for that reason, it is advised to combine a muscle
relaxant and decrease the dose of the cyclohexylamine), hyper-salivation and hyperthermia
(which can be the result of the hypertonicity and convulsions) (Burroughs et al., 2012a;
Caulkett & Arnemo, 2007). These drugs have an unpredictable effect on the excitement
phase (particularly if the veterinarian uses a lower dose), but the animals do not tend to run
or 'hackney gait' like the effect of the opioids on the ungulates. The use of α-2-agonists with
the cyclohexylamine might cause vomiting in some animals (Radcliffe & Morkel, 2014;
Burroughs et al., 2012a).
2.8.2.3.1 Ketamine
Ketamine can be found in different formulations from a large variety of
manufacturers and can be used in a broad spectrum of species, including wildlife. Although
its various administration routes, in wildlife practice, Ketamine is usually administered by IM
injection (usually painful), orally (as bait in higher doses) or by IV injection with an onset of
action between 3-5 minutes (complete immobilization in 5-10 minutes), with a smooth
induction (Lamont & Grimm, 2014; Burroughs et al., 2012a). Ketamine inhibits GABA
(Gamma-AminoButyric Acid), might suppress serotonin, norepinephrine and dopamine in the
CNS and activates the limbic system (Plumb, 2008). The effects are dose-dependent with 2-
3 hours of duration and might cause convulsions (reason why ketamine is often combined
with other drugs to reduce this effect) and consequently hyperthermia (Burroughs et al.,
2012a; Caulkett & Arnemo, 2007). It induces anesthesia, catalepsy and amnesia (an
advantage if the animal has to be immobilized more than once), but also increases the heart
rate and the blood pressure and suppresses the respiratory rate (Lamont & Grimm, 2014;
Plumb, 2008). Although ketamine is mainly used in the immobilization of carnivores, it can be
given to ruminants as a top up drug, usually intravenously (Caulkett & Arnemo, 2007;
Nielsen, 1999). It is a safe drug with a margin up to 10x the recommended dose (Swan,
1993). Hallucinations are a side effect in humans but it is difficult to evaluate in animals,
although some primates and some felids present some abnormal behaviors and vocalization
during recovery (Lamont & Grimm, 2014; Fowler, 2008).
2.8.2.3.2 Tiletamine (/Zolazepam)
Tiletamine is available in combination with zolazepam as an injectable anesthetic
with a short induction of 5-8 minutes (Lamont & Grimm, 2014; Ramsay, 2014; Burroughs et
al., 2012a; Walzer & Huber, 2002). It induces a dissociative anesthesia similar to the one
with ketamine but it has a potency 3-4x higher and it is mainly used in carnivores by IM
injection (but also possible IV), that need to be immobilized for 2-4 hours or longer (Lamont &
Grimm, 2014; Burroughs et al., 2012a; Plumb, 2008; Walzer & Huber, 2002). It can cause
muscular hypertonicity but because of the combination with the zolazepam, it is an unusual
event. This combination reduces convulsions, induces muscle relaxation and improves the
recovery of the animal (Lamont & Grimm, 2014; Burroughs et al., 2012a; Swan, 1993).
However, some of the side effects of Tiletamine might include hyperthermia, cyanosis,
vomiting and abnormal vocalization (Nielsen, 1999). Some long-action effects on the CNS,
which can last 24-48 hours after administration, might include convulsions, weakness and
anorexia (Fowler, 2008). Most of the times the Tiletamine/Zolazepam (TZ) combination is
used with an α-2-adrenergic-agonist like medetomidine, which is reversible in felids allowing
for a reduction of the dose of the dissociative and, consequently, fewer side-effects (Ramsay,
2.8.2.4 TRANQUILIZERS
Tranquilizers can be often confused with sedatives because both reduce the
motor activity of the animal by acting at the adrenergic receptors level in the CNS and in the
PNS (Burroughs et al., 2012a). However, tranquilizers have a more selective action,
suppressing the behavior response and have potent effects on the autonomic and the
endocrine systems (Lamont & Grimm, 2014). The effects are irreversible and are not dose-
dependent, which means that if the practitioner increases the dose of the tranquilizer, the
effects are prolonged (including the side effects) but the level of tranquilization does not
increase, contrary to sedatives, which have a dose-dependent effect (if the veterinarian
administers a higher dose than recommended, the effects will increase as well) (Lamont &
Grimm, 2014; Burroughs et al., 2012a). The animal shows signs of reduced fear, anxiety and
aggressive behavior (Burroughs et al., 2012a). The 2 main groups of tranquilizers used
nowadays are the phenothiazine derivatives, like acepromazine (ACP) and the
butyrophenone derivatives (e.g. haloperidol) (Lamont & Grimm, 2014; Burroughs et al.,
2012a). Wildlife practice has taken advantage of both groups by associating them with
opioids for immobilizations or as long-acting tranquilizers to benefit the translocation of
ungulates by promoting effects that can last hours, days or even weeks. Consequently,
stress levels decrease and the welfare of the animals improves by reducing trauma injuries
and by upgrading adaptation to new environments (Burroughs et al., 2012a; Caulkett &
Arnemo, 2007). The action of the tranquilizers blocks dopamine receptors (located in the
cerebral cortex, basal ganglia and limbic system) as well as the α - and β - adrenoceptors,
with antagonistic effects. Blood pressure decreases, anorexia and convulsions might occur
and even inhibition of the thermoregulatory center, but some anti-emetic properties might be
favorable. Usually tranquilizers do not have any analgesic effect and they do not have
antidotes, in the event the practitioner should decide to reverse their effect, while sedatives
often do (Lamont & Grimm, 2014; Burroughs et al., 2012a).
2.8.2.4.1 Acepromazine
ACP is probably the most used phenothiazine derivative in veterinary practice,
combined or not with an opioid – historically named neuroleptanalgesia (Lamont & Grimm,
2014; Montané et al., 2003). In general, phenothiazines block the dopamine receptors at
post-synaptic level in the CNS and might inhibit the release of dopamine (Montané et al.,
2003). Side effects such as bradycardia, hypotension and a low hematocrit are often present.
ACP has also an antiemetic effect, reduces the GIT motility (mainly in equines), causes
relaxation, hyperthermia, protrusion of the third eyelid and it has some extrapyramidal effects
(e.g. convulsions, circling and chewing) (Lamont & Grimm, 2014; Burroughs et al., 2012a). It
lasts for 6-8 hours and, in specific cases of immobilizations for semen collection by electro-
ejaculation, ACP is not recommended because it can cause an inhibition of the ejaculation
(Burroughs et al., 2012a; Plumb, 2008).
2.8.2.4.2 Azaperone
Azaperone reduces the motor activity and inhibits the CNS catecholamines,
dopamine and norepinephrine (Burroughs et al., 2012a; Plumb, 2008). It causes sedation,
vasodilation and bradycardia (but the pulse remains strong), has minimal effects on
respiration (might improve the respiration rate) and has antiemetic properties (Plumb, 2008).
However, it does not have antagonist or analgesic effects and it can cause chewing,
torticollis, catalepsy and aggressive behavior in some species (e.g. the gemsbok) (Burroughs
et al., 2012a; Plumb, 2008). Its vasodilation properties can block the vasoconstriction effect
of the opioids, which is extremely important in species such as the elephant and the white
rhino. It can be administered by IV, IM or SQ injections with short-acting effects and it lasts
for 2-4 hours (with a smooth recovery) (Wolfe, 2015; Burroughs et al., 2012a). It is usually
combined with opioids like etorphine in immobilization cases (Lamont & Grimm, 2014) or
used on its own as a tranquilizer to improve the welfare in transportation or the adaptation to
new environments (Burroughs et al., 2012a).
2.8.2.4.3 Haloperidol
Haloperidol is a butyrophenone derivative that blocks the action of
catecholamines like dopamine, causing sedation and it is mainly used on its own for
transportation of antelopes as a medium-acting tranquilizer with an onset action between 5-
10 minutes up to 8-12 hours (Wolfe, 2015; Burroughs et al., 2012a; Hofmeyr, 1981).
Haloperidol does not cause ataxia or sleepiness but, in some antelopes (e.g. kudu, blesbok -
Damaliscus pygargus phillipsi, or red hartebeest - Alcelaphus buselaphus caama) it can
promote aggressive behavior. It cannot be mixed with opioids, hence why the veterinarian
must not use it in immobilization cocktails (Burroughs et al., 2012a; Hofmeyr, 1981).
2.8.2.4.4 Long-Acting Tranquilizers
This particular group of tranquilizers (phenothiazine derivatives) is known as
LANs – long-acting neuroleptics – and it is extremely useful in wildlife practice because of its
effects: the drugs reduce anxiety, excitement and motor activity (Burroughs et al., 2012a;
Fick, Mitchell & Fuller, 2007; Read, Caulkett & McCallister, 2000). Knowing that wildlife
immobilizations are performed on a daily basis in SA, the use of LANs by the practitioner
decreased the mortality rates in certain species (Burroughs et al., 2012a). Because of their
oil compounds, LANs must not be administered as an IV injection and IM/SQ injections
promote a slow absorption by the muscle tissue, reducing stress levels, trauma injuries and
improving translocations and adaptation to new environments for several days with a single
administration. It can also be used to control animals with aggressive behavior – particularly
males in certain species – or to calm animals that are hospitalized or in rehabilitation
(Burroughs et al., 2012a; Fick et al., 2007; Read et al., 2000; Diverio, Goddard & Gordon,
1996). The main problem of LANs is overdosing, which can induce side effects (e.g.
chewing, torticollis, anorexia, shivering, tremors, star-gazing and hyper/hypothermia). The
practitioner must administer the drugs by IM injection, preferably, using a pole syringe or a
projectile dart with a lower gauge needle on (Wolfe, 2015; Burroughs et al., 2012a; Kock et
al., 2012a; Fick et al, 2007). In felids, contrary to the ungulates, LANs must be carefully used
for behavioral changes, mainly in cheetahs (Ramsay, 2014). This group of drugs does not
have any specific antidote available but some of the side effects might be reversed with
biperiden or diazepam (Burroughs et al., 2012a).
2.8.2.4.4.1 Zuclopenthixol acetate
By promoting a decrease in stress levels, zuclopenthixol is administered to
improve adaptation to new environments (or confined environments for rehabilitation, e.g.)
and it can also influence the behavior of the individuals, particularly aggressive behavior
(Wolfe, 2015; Read et al., 2000). It is a medium-acting tranquilizer in herbivore species such
as the rhino and it must be carefully used in carnivores (Miller & Buss, 2015; Ramsay, 2014;
Burroughs et al., 2012a). The effects begin 1-2 hours after administration and it can last for
3-4 days (peak of action at 36 hours after administration) with possible side effects including
convulsions (Wolfe, 2015; Burroughs et al., 2012a).
2.8.2.4.4.2 Perphenazine enanthate
Due to its ester in sesame oil vehicle, the deposit of perphenazine in the
muscular tissue is slowly released after 12-18 hours after administration (peak of action at 36
hours) and it can last for 7-10 days (Wolfe, 2015; Burroughs et al., 2012a). Like
zuclopenthixol, perphenazine is used to reduce anxiety and stress in ungulates and to control
aggressive behavior (Wolfe, 2015; Burroughs et al., 2012a; Read et al., 2000). The animals
do not get drowsy and can eat/drink normally.
Sometimes, the veterinarian opts to use more than one tranquilizer to cover the
entire adaptation process of the animal, for example, by combining haloperidol and
perphenazine to facilitate the translocation of an ungulate and its adaptation to a new
environment (Burroughs et al., 2012a).
2.8.2.5 SEDATIVES
Sedatives are commonly used in veterinary practice. Their effect depresses the
CNS (not the PNS) causing drowsiness and they decrease the locomotor activity. These
effects are dose dependent and the side effects are minimal (Lamont & Grimm, 2014;
Burroughs et al., 2012a). These drugs are less selective than tranquilizers, promote sleep
and are reversible (Wolfe, 2015; Burroughs et al., 2012a). The two main classes of sedatives
are the benzodiazepines (e.g. diazepam, midazolam or zolazepam) and the α-2-adrenergic-
agonists (e.g. xylazine, medetomidine and detomidine) (Lamont & Grimm, 2014; Burroughs
et al, 2012a).
2.8.2.5.1 BENZODIAZEPINES
The benzodiazepines group is one of the primary sedatives used in veterinary
medicine, including wildlife practice. As mentioned before, zolazepam is combined with
tiletamine but other benzodiazepines such as diazepam or midazolam are also available and
commonly used in wildlife immobilizations (Burroughs et al., 2012a). They promote the action
of GABA, which reduces the release or the turnover of acetylcholine in the CNS. It induces
depression of the CNS and promotes sedation, skeletal muscle relaxation, anxiolytic and
anticonvulsive effects (Plumb, 2008). These drugs have amnesic properties, which are very
useful when the veterinarian has to re-immobilize the animal in a short period of time.
However, although the properties of sedatives result in amnesic effects in human beings, in
felids there is still some reluctance about these effects of benzodiazepines (Ramsay, 2014).
2.8.2.5.1.1 Diazepam
Diazepam was the first sedative to be used in the very first wildlife immobilization
procedures (Burroughs et al., 2012a). It is a muscle relaxant, anxiolytic, anticonvulsant and
hypnotic drug, also used as an appetite-stimulant (mainly IV) (Burroughs et al., 2012a;
Plumb, 2008). In some species it can cause ataxia, increase the CNS excitement and be
responsible for behavioral changes (Plumb, 2008). When mixed with other drugs, diazepam
can precipitate and so it cannot be included in the cocktail dart (Burroughs et al., 2012a;
Plumb, 2008). Oral diazepam, commonly used in felids, can be given as premedication with
bait, 1-3 hours before the immobilization (Ramsay, 2014).
2.8.2.5.1.2 Midazolam
Midazolam is a relatively recent drug when compared with diazepam and it is
more potent and effective (Burroughs et al., 2012a). It has replaced diazepam because it has
a more predictable IM absorption and it can be used in wildlife practice as a top-up drug, a
short-acting sedative in rhino re-location, or even in bait (Miller & Buss, 2015; Ramsay, 2014;
2.8.2.5.2 BENZODIAZEPINE ANTAGONIST: Flumazenil
Benzodiazepines antagonists compete with benzodiazepines at the receptors
level and reverse its effect in the CNS by antagonizing the sedation either in cases of
therapeutic use or in overdose cases (Plumb, 2008; Walzer & Huber, 2002). Flumazenil is
the most used antidote to reverse the zolazepam and promote a smooth recovery in 1-2
minutes after an IV administration (Ramsay, 2014; Burroughs et al., 2012a, Plumb, 2008).
2.8.2.6 α – 2 - ADRENERGIC AGONISTS
Alpha-2-agonists and opioids usually have a synergistic and addictive effect. At
the presynaptic level of the noradrenergic neurons, α-2-agonists inhibit the norepinephrine
release by bounding with the α-2-adrenoreceptors. Activity in the Sympathetic Nervous
System (SNS) is reduced and it results in a decreased heart rate and blood pressure
(Lamont & Grimm, 2014). It induces muscle relaxation, sedation and analgesia, and reduces
the stress response. In higher doses, it can induce vomiting because of the activation of the
chemoreceptor trigger zone, hypothermia (most of the time, but temperature rises can be
observed in animals in warm environments), miosis and hypoxemia. By inhibition of the
antidiuretic hormone, the animal usually has an increase of the urine production and a
decrease of gastrointestinal motility might result in bloat and colic problems, mainly in
herbivores (Wolfe, 2015; Lamont & Grimm, 2014).
2.8.2.6.1 Xylazine
Xylazine was probably the first α-2-agonist to be used in veterinary practice. It
can be administered in many species and is easily available at a low price (Lamont & Grimm,
2014; Burroughs et al., 2012a). It promotes good muscle relaxation, sedation and a short
period of analgesia (Burroughs et al., 2012a; Plumb, 2008). However, it can cause hyper
salivation, muscle tremors in some species, GIT motility suppression (leading to ruminal
atony and bloat), vomiting in big cats, hypertension followed by hypotension and bradycardia,
decreased heart contraction, respiratory depression, hyper or hypothermia according to the
environment temperature, and even abortion in the last trimester. It can be reversible with
yohimbine or atipamezole (Lamont & Grimm, 2014; Burroughs et al., 2012a; Plumb, 2008).
The depression on the CNS is caused by a decreasing of the release of the norepinephrine
and dopamine, and by the inhibition of norepinephrine release in the adrenoceptors at a
presynaptic level and at a postsynaptic level in the receptors located on the peripheral
vascular smooth muscle (Lamont & Grimm, 2014; Plumb, 2008). Xylazine can be combined
with opioids or cyclohexylamines to improve the immobilization by reducing the doses and
the induction time, and by promoting a better muscle relaxation (Lamont & Grimm, 2014;
2.8.2.6.2 Medetomidine
Medetomidine is about 10x more powerful than Xylazine (Burroughs et al.,
2012a). It is a sedative with analgesic properties commonly used in veterinary practice,
preferably in carnivores (Ramsay, 2014). Because of its selective bounding with the α-2-
receptors, the antidote available, atipamezole, is specifically used to reverse medetomidine
(Lamont & Grimm, 2014). It can cause bradycardia, bradypnea, hypothermia, vomiting and
hypotension (Lamont & Grimm, 2014; Burroughs et al., 2012a; Plumb, 2008). Medetomidine
is usually combined with opioids, ketamine or TZ and administered intramuscularly, although
it can be painful (Lamont & Grimm, 2014; Ramsay, 2014; Plumb, 2008).
2.8.2.6.3 Detomidine
Detomidine is a sedative also used because of its analgesic properties
(Burroughs et al., 2012a; Plumb, 2008). As a long-acting sedative, it is relatively safe to be
used in pregnant females and, like medetomidine, it is 10x more potent than xylazine
(Lamont & Grimm, 2014; Burroughs et al., 2012a). It is less used than medetomidine and it is
usually mixed with other drugs for the immobilization of specific species (e.g. zebra and white
rhino) (Miller & Buss, 2015; Lamont & Grimm, 2014; Burroughs et al., 2012a).
2.8.2.7 α – 2 - ADRENERGIC ANTAGONISTS
Probably the biggest advantage of using α-2-adrenergic-agonists lies in the fact
that these drugs have antidotes which can easily reverse its effects (Lamont & Grimm, 2014;
Ramsay, 2014; Plumb, 2008). The veterinarian has to be aware that the analgesic effect of
the α-2-agonists previously administered is also reversed with the antidote and the animal
might need other extra analgesic drug after the administration of the reversal. Vasodilation
and tachycardia as result of the use of reversal drugs need to be monitored (Lamont &
Grimm, 2014; Plumb, 2008). In some species, the practitioner may not administer the
antagonist to promote a relaxed recovery avoiding self-induced trauma lesions (Lamont &
Grimm, 2014).
2.8.2.7.1 Yohimbine
Yohimbine is administered intravenously, slowly, to reverse the effects of
xylazine, but it has no effect against medetomidine (Lamont & Grimm, 2014; Burroughs et
al., 2012a; Plumb, 2008). It can cause muscle tremors, hyper-salivation, tachypnea and
excitement of the CNS (Plumb, 2008).
2.8.2.7.2 Atipamezole
It is the first choice antidote to reverse medetomidine, due to its selectivity to α-2-
adrenergic receptors (Ramsay, 2014; Plumb, 2008). It is administered as an IM (or IV)
injection – the recommendations in carnivores are 2.5-5x the mg equivalent of α-2-agonits -
and might cause some excitement in some cases and in ruminants, 5x the mg equivalent of
α-2-adrenergic-agonits (Lamont & Grimm, 2014; Ramsay, 2014; Burroughs et al., 2012a).
However, it is an expensive antidote and, occasionally, it might induce vomiting, diarrhea,
hyper-salivation, tremors and aggressive behavior. In painful procedures, the practitioner
must considerer the administration of additional analgesia (e.g. butorphanol) to improve the
recovery because atipamezole can reverse the main analgesic properties of the α-2-
adrenergic-agonists (Plumb, 2008; Lamont & Grimm, 2014).
2.8.2.8 OTHER DRUGS USED IN WILDLIFE PRACTICE
2.8.2.8.1 Doxapram
Doxapram is a temporary CNS stimulant and is used by practitioners to improve
respiration by stimulation of the medullary respiratory center and through the reflex activation
of the peripheral aortic and carotid sinus chemoreceptors (Burroughs et al., 2012a; Plumb,
2008). Although it's fast-acting, its effect is very short after IM or IV administration. The
veterinarian must replace its action by administering butorphanol or naltrexone when the
animals are still hypoxic (Burroughs et al., 2012a). In white rhinos, it must be used with
caution because it can cause excitement of the CNS and increase muscle tremors (Radcliffe
& Morkel, 2014).
2.8.2.8.2 Hyaluronidase
Hyaluronidase is an enzyme that liquefies hyaluronic acid, decreases the
viscosity of the connective tissue and promotes a better diffusion of the injectable drugs
(Burroughs et al., 2012a; Cattet & Obbard, 2010). In wildlife practice it has been used to
improve the efficacy and safety of the chemical immobilization of free-ranging animals.
Hyaluronidase is added as a powder to the drugs in the dart to promote the drug(s)
absorption by the muscle and reduce the induction time in several species (e.g. giraffe)
(Cattet & Obbard, 2010).
2.9 COMPLICATIONS IN WILDLIFE IMMOBILIZATIONS
2.9.1 STRESS
Stressful procedures such as a chemical immobilization, physical restraint or fear
of humans can lead to an acute stress response that will cause distress and influence the
homeostasis of the wild animal (Arnemo et al., 2014). The two most important responses to
stress are mediated by the hypothalamic-pituitary-adrenal (HPA) axis (long-term response)
and by the SNS, which stimulates the adrenal glands to release catecholamines (adrenaline)
to improve and prepare the body's response for the threatening situation (short-term
response) (Hernandez, 2014; Meltzer & Kock, 2012; Sheriff, Dantzer, Delehanty, Palme &
Boonstra, 2011; Macbeth, Cattet, Stenhouse, Gibeau & Janz, 2010; Terio, Marker & Munson,
2004). Endogenous or exogenous stressful events stimulate the HPA, which leads to a
disturbance of the animal's homeostasis. As a response, the pituitary gland releases ACTH,
which stimulates the zona fasciculata cells of the adrenal gland, resulting in an increase of
glucocorticoids levels. The high levels of cortisol, the main glucocorticoid, have a long-term
catabolic effect, reducing fertility, leading to stunted growth and suppression of the immune
system (Hernandez, 2014; Macbeth et al., 2010; Terio et al., 2004). An increase of
glucocorticoids causes a short-term 'fight or flight' stress response (adaptive) with the
production of adrenaline by the adrenal medulla, which is a part of the ANS. Adrenaline
increases heart rate, pulse and blood pressure, and decreases digestion and other
parasympathetic nervous system responses, which will only cease when the threat is over
(Meltzer & Kock, 2012; Sheriff et al., 2011; Macbeth et al., 2010). These stress-related
changes can lead to major complications such as physical trauma, hyperthermia, exhaustion
(capture myopathy), acidosis, shock and, ultimately, death (Arnemo et al., 2014; Hofmeyr et
al., 2012).
Stress can be classified in three different phases: alarm, resistance or adaptation
and exhaustion. The first stage will induce a 'fight or flight' response; at the second stage, the
stress factor is still increasing the levels of glucocorticoids, catabolic effects are present and
the immune system is suppressed. Finally, at the exhaustion stage, catabolism and immune
system suppression make the animal extremely vulnerable to succumb to other diseases,
increasing the chances of gastric ulcers and cellular degeneration scenarios (Hofmeyr et al.,
2012; Meltzer & Kock, 2012).
Maladaptation is another complication, particularly in recently re-located animals.
It can be associated with stress and usually means the unsuitability of an animal to a new
environment which is sometimes related with inappropriate nutrition. For example,
springboks are often moved to bushveld wet areas where they succumb to problems such as
heartwater because of the stress of the transport to a new habitat with a new climate (Meltzer
& Kock, 2012).
2.9.2 CAPTURE MYOPATHY
Capture Myopathy (CM) is probably the most common complication in free-
ranging wildlife (Wolfe, 2015; Arnemo et al., 2014; Spraker, 1993). It is a noninfectious,
metabolic disease found in domesticated and non-domesticated animals which can result in
death (Paterson, 2014) It has been termed in literature as: Capture Stress, Stress Myopathy,
Disease Muscle Dystrophy, Exertional Myopathy, Exertional Rhabdomyolysis, Overstraining
Disease, Degenerative Polymyopathy, Leg Paralysis, White Muscle Disease, Idiopathic
Muscle Necrosis, Muscle Necrosis, Diffuse Muscular Degeneration or even March
Myoglobinuria/Exertional Rhabdomyolysis syndrome in human beings, especially in
untrained athletes following hard exercise (Blumstein et al., 2015; Paterson, 2014; Sanchez,
2011; Cattet, Stenhouse & Bollinger, 2008b; Hartup, Kollias, Jacobsen, Valentine & Kimber,
1999; Spraker, 1993). This life-threatening syndrome was described decades ago in
herbivores but it is now reported in several different mammals, reptiles, fish, amphibians and
avian species associated with extended pursuits, restraint techniques, re-location and other
stress-inducing factors (Blumstein et al., 2015; Wolfe, 2015, Hernandez, 2014; Napier &
Armstrong, 2014; Paterson, 2014; Kohn, 2013; Sanchez, 2011; Cattet, Boulanger,
Stenhouse, Powell & Reynolds-Hogland, 2008a; Cattet et al., 2008b; Hartup et al., 1999;
Spraker, 1993). For example the stress of being chased by a helicopter, stimulates the
animal to run faster and longer, resulting in an intensive muscular activity and exhaustion
which causes CM (Arnemo et al., 2014; Arnemo et al., 2006). An alteration of the blood flow
to the tissues caused by a stress-inducing factor will exhaust the skeletal and cardiac
muscular adenosine triphosphate (ATP – aerobic energy), reducing the delivery of oxygen to
the different tissues in the body and increasing the production of lactic acid (responsible for
the metabolic acidosis). This phenomenon will cause muscle necrosis by an incorrect
removal of cellular waste and, consequently, myoglobin in circulation increases, leading to
myoglobinuria and inappropriate renal function (Wolfe, 2015; Paterson, 2014; Meltzer &
Kock, 2012; Spraker, 1993). Other signs of CM often identified in wildlife practice include
prostration, ataxia, inability to stand, hyperthermia, tachypnea, tachycardia, muscle stiffness,
tremors and torticollis, paralysis, muscle pain and acute renal failure (Blumstein et al., 2015;
Wolfe, 2015; Hernandez, 2014; Paterson, 2014; Kohn, 2013; Hofmeyr et al., 2012; Sanchez,
2011). Typically, animals become unresponsive, anorectic and death can occur within
minutes to hours, days, or even weeks after the immobilization event (Blumstein et al., 2015;
Paterson, 2014). The word rhabdomyolysis describes the pathophysiology behind the CM
syndrome. It means 'dissolution of skeletal muscle', which results in a rupture of the skeletal
muscle fibers with a leakage of intracellular creatinine phosphokinase (CK) and myoglobin
into the blood (Paterson, 2014).
CM can be prevented by decreasing stress levels during the immobilization,
keeping handling, noise, restraint, external stimulation and danger perception to a minimum
especially if predisposing factors are present (SECONDS) (Arnemo et al., 2014; Hernandez,
2014; Pas, 2014; Sanchez, 2011).
To avoid CM, the veterinarian can limit the pursuit to 3 minutes (ideally), wait for
a temperature below 20ºC, reduce visual and auditory stimuli and decrease the induction and
immobilization time, minimizing the excitement and avoiding underdoses (Wolf, 2015;
Sanchez, 2011). However, when CM is detected, any treatment that the practitioner might
administer to avoid a fatal myopathy is often insufficient to reverse the situation, hence being
associated with a low success rate (Wolfe, 2015; Paterson, 2014). The animal has to be
cooled down and the veterinarian must start an aggressive fluid therapy, oxygen therapy,
metabolic acidosis correction with sodium bicarbonate, analgesics and muscle relaxants
(benzodiazepines such as midazolam or central-acting muscle relaxants like
methocarbamol), and vitamins (vitamin E and selenium, mainly but others might also be
administer). Some associations of corticosteroids and non-steroidal anti-inflammatory drugs
(NSAIDs) can be administered to protect vascular integrity (Wolfe, 2015; Hernandez, 2014;
Paterson, 2014; Hofmeyr et al., 2012; Sanchez, 2011). A number of authors recommend the
administration of dantrolene sodium, a lipid soluble hydantoin analog that prevents malignant
hyperthermia in humans and the exertional rhabdomyolysis in horses by suppressing the
calcium release from the sarcoplasmic reticulum. However, there are no reports of its effects
in wildlife yet (Paterson, 2014).
CM can be classified in four different categories: (Blumstein et al., 2015; Wolfe,
2015; Hernandez, 2014; Paterson, 2014; Sanchez, 2011; Spraker, 1993)
2.9.2.1 Hyperacute or Capture Shock Syndrome
Shock is the term used by the veterinary practitioners to describe a terminal
stage of organs' failure, such as a drop in blood pressure as a result of blood volume loss,
severe dehydration or high levels of parasitism (Meltzer & Kock, 2012). During the
immobilization, an acute death syndrome known as capture shock syndrome might occur
within 1-6 hours postcapture. The animal shows signs of tachypnea, tachycardia,
hyperthermia, hypotension, depression and ultimately death (Wolfe, 2015; Hernandez, 2014;
Paterson, 2014; Sanchez, 2011). By analyzing the serum enzymes the practitioner will detect
an increase of the aspartate aminotransferase (AST), of the CK and an increase of the
lactate dehydrogenase (LDH) (Paterson, 2014). Postmortem findings might include hepatic
congestion, pulmonary congestion and edema, small intestine congestion with blood-tinged
contents in the lumen. Histologically, areas of necrosis will occur in the skeletal muscle,
brain, liver, heart, adrenal glands, spleen, pancreas, kidneys and lymph nodes (Wolfe, 2015;
Hernandez, 2014; Paterson, 2014).
2.9.2.2 Ataxic Myoglobinuric Syndrome
Is the most commonly observed of the four syndromes and it can occur hours or
days after the immobilization (Paterson, 2014). Severe clinical signs of ataxia, torticollis and
myoglobinuria might be visible (Wolfe, 2015; Hernandez, 2014; Paterson, 2014; Sanchez,
2011; Spraker, 1993). Serum enzymes AST, CK, LDH and BUN (blood urea nitrogen) levels
are abnormally increased. The kidneys are dark and swollen, with dark content, tubular
necrosis and myoglobin casts; and the skeletal muscle has multifocal, pale, soft, dry areas,
associated with hypoxia and ATP deficiency, which results from an exhaustion of the aerobic
glycolysis (Wolfe, 2015; Hernandez, 2014; Paterson, 2014; Spraker, 1993). Animals with
moderate to severe signs have a higher chance of dying (Paterson, 2014).
2.9.2.3 Ruptured Muscle Syndrome
Occurs within 24-48 hours postcapture. Clinical signs might include a drop of the
hindquarters and uni or bilateral hyperflexion of the hock, caused by the rupture of
gastrocnemius muscle, middle and deep gluteal, semitendinosus or semimembranosus
muscles (Wolfe, 2015; Hernandez, 2014; Paterson, 2014; Sanchez, 2011; Spraker, 1993). In
the serum, AST, CK and LDH are extremely increased but BUN is often normal or slightly
increased. The animal shows signs of SQ hemorrhage of the rear limbs, muscle necrosis and
multifocal pale lesions on the forelimb, hind limb, diaphragm, cervical and lumbar muscles
(Paterson, 2014). The type of lesions found in animals with this syndrome is also compatible
with vitamin E and selenium deficiency, which can be related to CM. Individuals with this
form of CM might resist and survive for some weeks but most of them will die (Hernandez,
2014; Spraker, 1993).
2.9.2.4 Delayed-Peracute Syndrome
This is a rare syndrome that could appear more than 24 hours after the
immobilization procedure (Paterson, 2014). Although the animals seem to be normal, sudden
death occurs due to a ventricular fibrillation and high levels of AST, CK and LDH (Wolfe,
2015; Hernandez, 2014; Paterson, 2014; Sanchez, 2011; Spraker, 1993). There are no
visible lesions but sometimes small pale foci on the skeletal muscle are detected,
characterized by a mild to moderate rhabdomyolysis throughout the skeletal muscle,
particularly in the hind limbs (Wolfe, 2015; Hernandez, 2014; Paterson, 2014; Spraker,
Differential diagnosis for CM in wildlife might include plant toxicity, malignant
hyperthermia, early tetanus, hypocalcemia or myositis (Paterson, 2014).
2.9.3 RENARCOTIZATION
Renarcotization is a particular complication in wildlife medicine, particularly in
herbivores immobilized with opioids (Napier & Armstrong, 2014; Pas, 2014). Animals that
have already been immobilized and reversed with an antidote will show signs of re-sedation
hours after the reversal has been administered (Pas, 2014). Antagonists like diprenorphine to
reverse carfentanil or thiafentanil might cause re-narcotization. Because of its long half-life,
naltrexone is a much safer choice if the practitioner intends to re-immobilize the animal
without the chance of the animal being re-sedated after the antidote has been administered
(Napier & Armstrong, 2014; Pas, 2014). But the veterinarian must use the correct doses of
antidote to reverse the opioid otherwise an underdose might cause renarcotization (Napier &
Armstrong, 2014).
Opioid levels in the blood will maintain the animal's clinical signs for 12-24 hours
after the administration of the reversal. Some of the signs might include excitement, circling,
high stepping, recumbency paddling and, consequently, hyperthermia, acidosis and
exhaustion. In these situations, the veterinarian needs to repeat the antidote by IM
administration of half of the dose. Sometimes, if the second dose of antidote does not
reverse the animal completely, the practitioner must repeat the dose until the clinical signs of
renarcotization cease (Napier & Armstrong, 2014).
2.9.4 PHYSICAL TRAUMA
An immobilization procedure, with physical or chemical restraint, can be traumatic
for the animal involved. Injuries such as lacerations, fractures or abrasions can be
accidentally inflicted on the animal, by itself, by other animals or by the team in charge. A
proper handling and a correct prediction of the animal's behavior as well as previous
knowledge of the environmental hazards might prevent several physical traumas (Arnemo et
al., 2014; Arnemo et al., 2006). Trauma can even lead to death in capture procedures and a
loss of 10 % of the animals in an immobilization is expected. However, when immobilization
involves endangered species, a loss of a single individual can be dramatic and this
percentage must not be accepted (Hernandez, 2014; Meltzer & Kock, 2012).
2.9.5 HIPOXEMIA AND HYPOXIA
One of the most common complications that can occur during immobilization is
hypoxemia (low arterial oxygen tension) brought on by hypoventilation, which can be a result
of respiratory depression caused by some immobilization drugs. It may also be a
consequence of loss of lung volume, possibly resulting from an incorrect recumbency
position. In herbivores, for example, the GIT content presses the diaphragm forward,
decreasing the volume of the lungs and compromising the expansion of the thorax on
inspiration (Arnemo et al., 2014; Fahlman, 2014; Lamont & Grimm, 2014; Napier &
Armstrong, 2014; Pas, 2014; Cracknell, 2014; Meltzer & Kock, 2012; Bush, Raath, Grobler &
Klein, 2004). An unusual cause of hypoxemia is pneumothorax, which can happen after a
dart penetration into the thoracic cavity or a puncture of a horn (Arnemo et al., 2014; Arnemo
et al., 2006). Hypoxia, the inadequate oxygen level in the body tissues, is a consequence of
hypoxemia when the oxygen delivery is not improved (by increasing the cardiac output or by
decreasing the oxygen consumption in the tissues). This status results in rapid cellular
degeneration in organs like the brain, heart, kidneys and liver. A lower perfusion of the
tissues with low levels of hemoglobin can also be responsible for hypoxic status because the
tissues might not be able to use oxygen properly or because the hemoglobin is not able to
transport the oxygen to the tissues (Fahlman, 2014; Cracknell, 2014).
2.9.6 HYPERTERMIA/HYPOTHERMIA
Overheating during an immobilization procedure in wildlife practice is an
expected complication. Hyperthermia can result from high environmental temperatures,
especially during summer or during the middle of the day. It occurs due to excessive
muscular exercise and muscular contraction, when the animals are being chased or
frightened, or from the drugs used, which interfere with the thermoregulatory center (Arnemo
et al., 2014; Schumacher, 2008; Burroughs & McKenzie, 1993). The ideal temperature for
immobilizations should be around 25ºC (Meltzer & Kock, 2012). When its temperature rises
above 43ºC the animal might show signs of panting, weakness, arrhythmia, shallow
breathing, convulsions or even die. A rectal temperature of more than 41ºC is considered an
emergency and, in these cases, the practitioner has to act (Arnemo et al., 2014; Meltzer &
Kock, 2012; Schumacher, 2008). To avoid this situation or to cool down the temperature, the
animal has to be moved to a shaded area and sprayed with cold water (or cooled in a river,
with snow, fluids or enemas), supplied with oxygen (because hyperthermia increases the
oxygen demand) or keep stress to a minimum by avoiding an intense chase or stressful
restraint methods (Arnemo et al., 2014; Hofmeyr et al., 2012; Schumacher, 2008). The
practitioner must always evaluate the conditions and decide if the immobilization will go
ahead or not (Arnemo et al., 2014; Hofmeyr et al., 2012; Meltzer & Kock, 2012).
Hypothermia is characterized in mammals by a temperature below 35ºC and
while not as common as hyperthermia, can also occur, particularly if the immobilization is
taking place in areas with low temperatures, wet, rainy environments or when the animals
immobilized are young, small or in poor body condition (Arnemo et al., 2014; Schumacher,
2008). This condition can be reversed by warming the animal up with hot water bottles or by
drying the animal. To prevent hypothermia, weather conditions such as exposure to wind or
low temperatures must be avoided (Arnemo et al., 2014).
2.9.7 BLOAT
In ruminants, a correct recumbency position (sternal) during an immobilization
procedure is vital, otherwise in a lateral position bloat can occur. Another cause of bloat is
the use of immobilization drugs such as the α-2-agonists, which will result in a ruminal atony
and, consequently, bloat. That tympanic stage happens due to the inability to expel gases
through eructation. If the immobilized animal starts to bloat, it must be re-positioned into a
sternal recumbency with the neck extended and the head with the nose pointing down to
drain the saliva. Intubating the animal to relieve the gases inside might be an option, or even
a trocharization of the rumen. If the problem is caused by α-2-adrenergic-agonists, the
veterinarian might use the antidotes available to antagonize their effect (Arnemo et al., 2014;
Napier & Armstrong, 2014; Hofmeyr et al., 2012; Arnemo et al., 2006).
2.9.8 VOMIT/REGURGITATION AND ASPIRATION PNEUMONIA
Vomiting and regurgitation are considered emergencies since they might entail
the possibility of aspiration pneumonia. Vomiting could be a side effect of opioids or α-2-
adrenergic-agonists administration. As mentioned before, the correct recumbency position is
an important aspect in wildlife practice. While ruminants are usually in a sternal position with
the neck straight and the nose down, carnivores are positioned laterally. If the practitioner
notices that the animal has aspirated the stomach/rumen content and might develop
pneumonia, he will need to administer a broad spectrum long-acting antibiotic (Arnemo et al.,
2014; Pas, 2014; Hofmeyr et al., 2012; Arnemo et al., 2006).
2.9.9 EQUIPMENT FAILURE
Notwithstanding complications that can stem from an inadequate planning of the
immobilization, selecting the right drug(s) and dose, or even complications associated with
inexperienced personnel, equipment failure is always a factor that still can occur during a
procedure (Arnemo et al., 2014; Atkinson et al., 2012). Darts form part of basic equipment for
wildlife immobilizations but can still suffer a number of failures and problems during
procedures (Hernandez, 2014; Shury, 2014). When the needle impacts on the animal it can
bend or even break, meaning the drug(s) might be inadequately administered. An
inappropriate loading of the dart in the gun might cause an incorrect projection, while a loss
of the tailpiece can cause a loss of balance. Power charges might not work properly because
of the weather conditions and the carbon dioxide or the compressed-air cylinders may leak.
The practitioner might miss the target or hit the animal in the wrong place (SQ tissue, fat
deposit, chest or abdomen, bone or vital organs).The strength of impact/velocity of impact
can influence the injection of the content into the animal. The dart gun and the dart might be
rusty or dirty; the dart wound might cause infections or a misplaced dart might even cause a
pneumothorax (Arnemo et al., 2014; Hernandez, 2014; Atkinson et al., 2012; Arnemo et al.,
PRACTICAL CASES
Given that the goal of this thesis is to review the main principles of wildlife
immobilization, the anesthetic protocols used on the various African species immobilized are
presented in this chapter, along with the doses and the characteristics of the darts/ dart guns
involved in each capture procedure.
Graphic 1 shows the percentage of the different species examined during the
internship. Most of the species were ungulates (85.32%) and only 14.68% were carnivores.
The African buffalo was the most immobilized species, followed by the sable antelope, the
blue duiker and the African lion. At the other end of the spectrum, species such as the
leopard, the white rhino or the bushbuck were only immobilized once. Table 2 represents the
exact number of individuals restrained and the respective number of chemical immobilization
done for each species approached.
Graphic 1 – Percentage of individuals from different species examined during the
0.82% 10.20%
Table 2 – Number of animals, divided by species, immobilized during the
internship (total of 245 animals and 184 chemical immobilizations).
Golden Gemsbok
African Elephant
Sable Antelope
African Buffalo
Plains Zebra
Blue Duiker
Bushbuck
99 42 8 11 1 1 25 6 6 1 5 2 2 25 1 5 5
chemical 98 31 8 3 1 0 0 6 5 1 5 2 2 14 1 2 5
immobilizations
This large diversity of species is representative of the various procedures
performed. All the procedures were grouped in fourteen categories according to the species
immobilized (Table 3).
Table 3 – Different activities performed during the internship divided according to the species and the procedure.
Transport, re- Tests: TB, X-rays, US, Wound
Total of vitamins, Ear GPS Necropsies,
SPECIE location, off- Microchip Measurements Implants Surgery Vaccination GnRH, DNA, Pregnancy Certifications treatments, TOTAL
individuals de- Tag Collar Eutanasia
load or boma coprology exams check ups
African Buffalo 99 16 99 22 61 22 0 0 0 95 21 54 1 0 1 392
Sable Antelope 42 28 35 19 13 3 0 0 0 25 16 5 0 0 2 146
Roan Antelope 8 2 8 4 5 2 0 0 0 4 4 0 0 0 0 29
11 0 1 0 0 0 0 0 2 6 4 0 0 1 1 15
Bushbuck 1 1 1 0 0 0 0 0 0 0 0 0 0 0 1 3
Blue Duiker 25 0 25 25 0 25 0 0 0 0 25 0 0 0 1 101
Bontebok 6 0 6 6 0 6 0 0 0 0 6 0 0 0 0 24
Springbok 6 5 5 0 0 0 0 0 1 0 0 0 0 0 0 11
White Rhino 1 0 0 0 0 0 0 0 1 0 0 0 1 0 0 2
Giraffe 5 5 4 0 0 0 0 0 0 0 0 0 1 0 0 10
Plains Zebra 2 0 2 0 0 0 0 0 3 0 0 0 0 0 0 5
African Lion 25 3 16 0 0 0 5 3 5 13 0 0 0 0 3 48
Leopard 1 0 1 0 0 0 0 0 2 0 0 0 0 0 0 3
Cheetah 5 1 2 1 1 0 0 0 0 4 1 0 0 0 0 9
African wild
5 5 5 4 1 0 0 0 0 4 0 4 0 0 0 23
TOTAL 245 69 212 82 82 58 5 3 14 151 77 63 3 1 9 829
GPS (Global Positioning System), TB (Bovine Tuberculosis), GnRH (Gonadotropin-Releasing Hormone), US (Ultra-Sound)
3.2 METHODS, MATERIALS AND RESULTS
The tables presented later on explain the different anesthetic protocols used on
the various African species immobilized during the internship. The doses of the drugs (used
for immobilization, used as reversals and as additional administrations) and the
characteristics of the darts/ dart guns involved in each procedure are tabled along with the
results (immobilization succeeded or not succeeded) for each protocol performed. Additional
comments are also provided about the results obtained.
The weight of the species was established according to the references from Miller &
Fowler (2015) except for the male bushbuck. Its weight is based on a reference by Kock &
Burroughs, 2012. As can be seen below, for ungulates, the doses are tabled as net doses in
mg and not associated with the weight of the animal (mg/kg) because the various protocols
published by the main wildlife practitioners are also in net doses references (see Appendix 2.
Table 20-34, where the different protocols suggested by the main wildlife practitioners are
tabled).
3.2.1 AFRICAN UNGULATES
3.2.1.1 AFRICAN BUFFALO
The African buffalo is one of the most commonly immobilized species in SA
because of its impact on livestock (and vice-versa). In terms of diseases legally requiring
testing, bovine tuberculosis (TB), brucellosis, foot and mouth disease (FMD) and corridor
disease (Theileriosis by Theileria parva) are the most important ones (Tanner et al., 2015;
Anderson, Ezenwa & Jolles, 2013; Beechler, Broughton, Bell, Ezenwa & Jolles, 2012;
Burroughs et al., 2012b; Pienaar, Potgieter, Latif, Thekisoe & Mans, 2011; Chaisi, Sibeko,
Collins, Potgieter & Oosthuizen, 2011; Ayebazibwe et al., 2010; Sibeko et al., 2008; De Klerk
et al., 2006; Oosthuizen, 2006; Jolles, Cooper & Levin, 2005; Kalema-Zikusoka, Bengis,
Michel & Woodford, 2005; Anderson, Foggin et al., 1993; Anderson, Doughty, Anderson &
Paling, 1979; Hedge, 1972).
Figure 10 – Herd of buffalos in a Game Farm (semi-controlled environment) (Original).
During the internship, most of the buffalos examined were in large parks/semi-
controlled environments, sometimes in bomas (Figure 10) and any time an animal did not go
down after the first dart, a second dart with the same dose or with half of the dose was given
(Table 4).
Table 4 – African buffalo's anesthetic protocols and results.
DART DART
SAMPLE OF PROTOCOL REVERSAL RESULTS COMMENTS
GUN TYPE
IV administration of 7.5mg of
Midazolam if needed;
Thiafentanil:4mg +
IV administration of
Bull (300- Etorphine:4mg
22 Well succeeded Medetomidine 1mg if needed
900kg) (+ Medetomidine:1-2mg)
Diprenorphine:3 (and reversed with Atipamezole
+ Azaperone: 40-60mg
x Etorphine + 5mg);
2cc darts
Pneu Naltrexone: 10x 1 individual required a 2nd dart
1.5'' WB
Dart gun Thiafentanil:3mg + Thiafentanil
Cow (300- type C Etorphine:3mg (+ Or
55 Well succeeded Midazolam if needed;
900kg) Medetomidine:1mg) Naltrexone: 10x
2 individuals required a 2nd dart
+ Azaperone: 40-60mg Thiafentanil
Thiafentanil: 2.5-3.5mg +
Juvenile 5 Well succeeded -
Azaperone:40mg
Calf 16 Thiafentanil: 1.5mg Well succeeded -
3/4'' WB
Drugs with ( ) were not administered to all animals
Thiafentanil (Thianil®, 10mg/m, Wildlife Pharmaceuticals, Mpumalanga, SA)
Etorphine (M99®, 9.8mg/mL, Novartis, Vorna Valley, SA)
Naltrexone (Trexonil®, 50mg/mL, Wildlife Pharmaceuticals, Mpumalanga, SA; Naltrexone®, 50mg/mL, Kyron
Labs, Johannesburg, SA)
Diprenorphine (M5050®, 12mg/mL, Novartis, Vorna Valley, SA)
Medetomidine (Medetomidine®, 20mg/mL, V-Tech, Pretoria, SA; or Domitor®, 1mg/mL, Pfizer, Port Elizabeth,
SA)
Atipamezole (Antisedan®, 5mg/mL, Pfizer, Port Elizabeth, SA)
Azaperone (Stresnil®, 40mg/mL, Bayer, Isando, SA)
Midazolam (Midazolam®, 15mg/3mL, Roche Products, Illovo, SA)
3.2.1.2 SABLE ANTELOPE
Most of the sables were chemically immobilized in large parks but the smallest
ones were inside bomas. Contrary to others captures, sable and roan antelopes were not
blindfolded but, because of their horns, they required horn protections (plastic pipes) to
improve the safety of the procedure for the personnel. Zuclopenthixol (Clopixol-Acuphase®,
50mg/mL, Lundbeck Limited, North Riding, SA) or haloperidol (Haloperidol®, 20mg/mL, V-
Tech, Pretoria, SA) IM were also administered for re-location purposes, allowing an
improvement during the transportation and a better adaptation to a new environment (Table
Table 5 – Sable antelope's anesthetic protocols and results.
IV administration of 5-15mg of
Bull (190- Thiafentanil:6-7mg IV administration of
19 Well succeeded
300Kg) + Azaperone:40-60mg Medetomidine 1mg if needed
(and reversed with Atipamezole
5mg)
2cc darts IV administration of 5-15mg of
3/4'' WB Naltrexone 10x Midazolam if needed;
Cow (190- Thiafentanil:6mg Thiafentanil IV administration of
6 type P Well succeeded
Thiafentanil: 3.5mg +
Azaperone:40-60mg
1cc darts Thiafentanil:1.5mg +
Calf 2 Well succeeded -
3/4'' WB Azaperone: 40-60mg
3.2.1.3 ROAN ANTELOPE
All the roan antelopes were chemically immobilized in large parks and
approached by foot or by vehicle (Table 6). The animals immobilized were adult males for
DNA testing or bio-measurements.
Table 6 – Roan antelope's anesthetic protocols and results.
Naltrexone 10x Midazolam if needed;
Pneu 2cc darts Thiafentanil:7mg +
Bull (190- Thiafentanil + And/or
8 Dart gun 1+1/4'' (Medetomidine:1-2mg) + Well succeeded
300Kg) Atipamezole 1 mg of Medetomidine
type P WB Azaperone: 40-60mg
5xMedetomidine (reversal:Atipamezole 5mg);
3.2.1.4 SPRINGBOK
The springboks immobilized were darted by foot and by vehicle in non-controlled
environments. They were captured for re-location purposes (only one was physically
restrained and it is not part of the samples in Table 7), and haloperidol was given IM to the
individuals (Table 7).
Table 7 – Springbok's anesthetic protocols and results.
Ram (30- Thiafentanil:1.5-1.7mg+
1 Well succeeded -
45Kg) Azaperone:20mg
1cc darts Naltrexone 10x
Dart gun One death: not related to the
1'' GC Thiafentanil
Ewe (30- type P Thiafentanil:1.5mg+ ¼ not succeeded; anesthetic protocol – Drowned
45Kg) Azaperone:20mg ¾ well succeeded in a lake during the excitement
3.2.1.5 BONTEBOK
Due to the slight visual differences between a bontebok and a blesbok (Damaliscus
pygargus phillipsi), many owners in SA want their animals tested to make sure the individuals
are not a result of an unwanted cross-breeding (van Wyk, Kotzé, Randi & Dalton, 2013; Van
der Walt, Nel & Hoelzel, 2001). The free-ranging bonteboks examined were approached by
vehicle and chemically immobilized for DNA testing (Table 8).
Table 8 – Bontebok's anesthetic protocols and results.
SAMPLE DART GUN DART TYPE PROTOCOL REVERSAL RESULTS
Ram (55-
3 Well succeeded
80Kg)
Pneu Dart gun 1cc darts 1'' Thiafentanil:3mg+ Naltrexone 10x
type P GC Azaperone:20-40mg Thiafentanil
Ewe (55-
3.2.1.6 GIRAFFE
Giraffe capture is a complex and dangerous procedure. Usually, a free-ranging
giraffe is chemically immobilized mainly for re-location purposes because of the high risk of
complications during restraints that could result in death. The goal of this restraint relies on a
fast and efficient darting, preferably by helicopter, with a very high dose of opioid (Table 9).
Excessive running must also be avoided (Bertelsen, 2015; Citino & Bush, 2014; Citino, Bush,
Lance, Hofmeyer & Grobler, 2006; Bush, 2003; Bush et al., 2001; Bush, Raath, Phillips &
Lance, 1997; Bush, 1993; Morkel, 1993; Morkel, 1992; Bush & de Vos, 1987; Bush, 1976;
Bush, Ensley, Mehren & Rapley, 1976). During captures, the giraffes were always
approached by helicopter.
Table 9 – Giraffe's anesthetic protocols and results.
Etorphine:10mg +
Cow (700- 2cc darts
1 Butorphanol: 10mg + Well Succeeded -
1200Kg) 2''WB
Hyaluronidase: 1500-3000 IU
Pneu Etorphine:4-6mg +
Dart gun Butorphanol: 5mg/ 10x opioid One death: not related with the
¾ Well
type C 2cc Etorphine:4-5mg + anesthetic protocol – Fall into a
Juvenile 4 succeeded; ¼ Not
1.5''WB Thiafentanil:2mg + mud pit during the excitement
Butorphanol:15mg + phase
Butorphanol (Butorphanol® 50mg/mL, Kyron Labs, Johannesburg, SA or Butorphanol® 10mg/mL, Kyron Labs,
Johannesburg, SA)
Hyaluronidase (Hyalase®, 5000 IU, Kyron Labs, Johannesburg, SA)
3.2.1.7 AFRICAN ELEPHANT
Although there is no published protocol for a standing sedation on a free-ranging
African elephant, in controlled environments the standing sedation is a common procedure
(Wiedner, 2015; Horne, Loomis,
2014; Fowler & Mikota, 2006;
Neiffer et al., 2005; Du Toit, 2001;
Ramsay, 2000; Raath, 1993). The
standing sedations were carried out
in a semi-controlled environment, to
perform surgery on the tusks and
blood collection for testosterone
levels testing (Table 10).
The free-ranging bull
elephant was darted with an
experimental cocktail for standing Figure 11 – Free-ranging bull elephant sedated for
sedations (Figure 11). blood collection and GnRH administration (Original).
Table 10 – African elephant's anesthetic protocols and results.
NUMBER OF DART DART
SAMPLE PROTOCOL RESULTS COMMENTS REVERSAL
ANIMALS GUN TYPE
For the not succeeded
Etorphine:3mg + Naltrexone
One succeeded immobilization: 2nd dart
Dan Medetomidine:40mg + 150mg (IV)
(1st protocol); with 30mg Medetomidine
2 (controlled Inject 5cc darts Butorphanol:150mg +
one not + 75mg Butorphanol
environment) pump 2'' and Atipamezole
Bull succeeded (2nd and top up (IV): 10mg
pistol Medetomidine:40mg 25mg (2/3
(4000- protocol) Medetomidine + 25mg
+Butorphanol:150mg IM + 1/3 IV)
7000Kg) Butorphanol
Etorphine:4mg +
Pneu 2nd dart: 14mg Etorphine
1 (free- 5cc darts Medetomidine:50mg Naltrexone
Dart gun Not succeeded 100 mg Azaperone + 5000
ranging) 2'' WB +Butorphanol:150mg + 5000 IU 140mg
type C IU Hyaluronidase
3.2.1.8 WHITE RHINOCEROS
The female white rhino darted for horn trimming was in a game reserve where the
white rhinos had had anesthetic complications in the past. For this reason, additional
measures were taken into account, particularly the choice of the protocol and respective
doses (Table 11). The animal was darted from a vehicle and followed carefully after that.
Because no evidence of sedation was noticed, a second dart was administered following the
same protocol.
Table 11 – White rhino's anesthetic protocol and results.
SAMPLE DART GUN DART TYPE PROTOCOL RESULTS COMMENTS
2nd dart with same
Etorphine:3.5mg +
Cow (1800- Not protocol (effective) but the
1 Pneu Dart gun type C 2cc dart 2'' WB Azaperone: 20mg +
2200Kg) succeeded animal died during
Hyalurinidase: 2500 IU
3.2.1.9 GOLDEN WILDEBEEST (Blue Wildebeest)
The Golden wildebeest is a color variation of the blue wildebeest species (van
Hoven, 2015). The two individuals darted were males, both immobilized for re-location
purposes (Table 12). A drone followed one of them during the entire procedure to guarantee
that the animal would not get lost after darting.
Table 12 – Golden wildebeest's anesthetic protocol and results.
NUMBER OF DART
SAMPLE DART TYPE PROTOCOL REVERSAL RESULTS COMMENTS
ANIMALS GUN
Bull (110- Pneu Dart 1.5cc darts ¾'' Thiafentanil: 3.5-4mg + Naltrexone Zuclopenthixol: 100mg
180kg) gun type P WB Azaperone: 40mg 10xThiafentanil was given IM
3.2.1.10 PLAINS ZEBRA
Two Plains zebra (a stallion and a mare) were darted on foot and immobilized in
order to perform hooves trimming and castration on the stallion (Table 13).
Table 13 –Plains zebra's anesthetic protocols and results.
SAMPLE DART TYPE PROTOCOL REVERSAL RESULTS
Etorphine: 6mg +
Stallion (175-275kg) 1 Well succeeded
Pneu Dart 2cc darts ¾'' Azaperone: 60mg Diprenorphine
gun type P WB Etorphine: 5.5mg + 3xEtorphine
Mare (175-275 kg) 1 Well succeeded
Azaperone: 60mg
3.2.1.11 BUSHBUCK
The male bushbuck was immobilized for re-location purposes after being found
stuck in a private backyard (Table 14).
Table 14 – Bushbuck's anesthetic protocol and results.
Pneu Dart Thiafentanil: 1mg +
Naltrexone 10xThiafentanil +
Ram (40-50Kg) 1 gun type 1cc dart 1'' GC Medetomidine Well succeeded
Atipamezole 5x Medetomidine
C (Domitor®): 1mg
3.2.2 AFRICAN CARNIVORES
3.2.2.1 AFRICAN LION
The African lions immobilized
during the internship were restrained for
different reasons and in different
environments (Figure 12). The approach was
done on foot or by vehicle, and most of them
were darted after a carcass (bait) had been
offered to improve the accuracy of darting
(Table 15).
Figure 12 – Anesthetized African lion in
Gondwana Game Reserve (Original).
Table 15 – African lion's anesthetic protocol and results.
SAMPLE DART GUN DART TYPE PROTOCOL REVERSAL RESULTS COMMENTS
1.5cc darts TZ: 0.4- The animals that went to surgery (2)
Lion Atipamezole
1''WB; 2cc 0.5mg/kg + required top-ups of Medetomidine
(120- 5 3x Well succeeded
¾'' WB; 2cc Medetomidine: 0.01-0.015 mg/kg and/or TZ 0.15-
250Kg) Medetomidine
1+1/4'' WB 0.03-0.04mg/kg 0.2mg/kg (IV or IM)
Pneu Dart
The animals that went to surgery (3)
gun type C
1.5cc darts TZ: 0.4- required top-ups of Medetomidine
Lioness or type P Atipamezole
1''WB; 2cc 0.7mg/kg + 0.01-0.015 mg/kg and/or TZ 0.15-
¾'' WB; 2cc Medetomidine: 0.2mg/kg (IV or IM) *;
1+1/4'' WB 0.04-0.07mg/kg 2 individuals required a 2nd dart with
half of the dose
TZ : Zoletil 100®, 100mg/mL, Virbac, Halfway House, SA (Tiletamine 50% + Zolazepam 50%)
* Two of the three surgeries performed on lionesses were hemi-hysterectomies in 2 members of the same pride
and, for that reason they were both immobilized at the same time along with the male. While the first surgery
was being performed, the second lioness was under isoflurane maintenance + TZ + Medetomidine.
3.2.2.2 CHEETAH
Cheetahs were immobilized in controlled environments: boma or enclosures
(Table 16). Even on free-ranging animals this method is always good practice to improve
darting opportunities. Other advice for adequate immobilization of carnivores might include a
top-up to maintain the desired level of anesthesia, guarantee the safety of the personnel. Eye
lubricant should be applied before the animal is blindfolded, and the practitioner must keep
an eye on the animal until it gets up, recovers from anesthesia and, if possible, keep
monitoring it for the next 24 hours (Lamberski, 2015; Ramsay, 2014; Crosier et al., 2007;
Stegmann & Jago, 2006; Wack, 2003).
Table 16 – Cheetah's anesthetic protocols and results.
SAMPLE OF DART GUN PROTOCOL REVERSAL RESULTS COMMENTS
The individual
required a 2nd dart
+ Top-up
TZ : 0.6mg/kg + Atipamezole 3x
Male (35-72Kg) 1 1cc Not succeeded administrations
Pneu Dart gun Medetomidine:0.05mg/kg Medetomidine
darts with
1''GC Medetomidine:
TZ: 0.5mg/kg + Atipamezole 3x
Female (35-72Kg) 1 Well succeeded -
Medetomidine:0.05mg/kg Medetomidine
3.2.2.3 LEOPARD
The male black leopard immobilized was in a controlled environment and surgery
was performed on this individual (Figure 13) as well as nail clipping due to excessive claw
growing (Table 17).
Figure 13 – Castration performed by Dr Brendan Tindall on a male black leopard (Original).
Table 17 – Male leopard's anesthetic protocol and results.
Top-ups required: 1st top up: TZ:
0.15mg/kg +
TZ: 0.5mg/kg Medetomidine:0.01mg/kg (2nd
Male (23- Pneu Dart + Atipamezole 3x Not dart); 2nd top up: TZ: 0.3mg/kg +
1 1cc darts 1''GC
91kg) gun type P Medetomidine: Medetomidine Succeeded Medetomidine:0.01mg/kg (Pole
0.05mg/kg syringe); 3rd top up: TZ: 0.3mg/kg
+ Medetomidine: 0.005mg/kg
(hand-held injection)
3.2.2.4 AFRICAN WILD DOG (Painted hunting dog)
The painted dogs are usually found in packs (Scott & Kreeger, 2014; Larsen,
Kreeger, West, Heard & Caulkett, 2007; Courchamp, Rasmussen & Macdonald, 2002; de
Villiers, van Jaarveld, Meltzer & Richardson, 1997) which might require a complete restraint
of all the individuals of the group at the same time to ensure the safety of the personnel and
the animals. As an endangered species in SA, a safe chemical restraint of all the immobilized
canids is critically important (Courchamp et al., 2002; de Villiers et al., 1997; van Heerden,
Burroughs, Dauth & Dreyer, 1991). Four of the five immobilizations performed started with
display of bait (carcass) to improve the accuracy of the darting (Table 18).
Table 18 – African wild dog's anesthetic protocols and results.
Male (19- Well
35Kg) succeeded
Administration of a 2nd dart on
the unsuccessful
TZ: 1mg/Kg + Atipamezole
Pneu Dart ¼ Not immobilization with 1/2 of the
Female 1cc darts ¾'' WB Medetomidine: 3x
gun type P succeeded; dose after some signs had
(19- 4 0.06mg/kg Medetomidine
¾ Well been detected but the animal
succeeded was not sedated enough to
tolerate an approach +
Medetomidine:0.02mg/kg
As evident in the results from Table 2 (Chapter 3), some of the restraints
performed did not require any chemical immobilization, such as the ones performed on blue
duikers. A physical restraint after a net capture was the preferable method used for
immobilization of these individuals. This technique is also adequate for other species, as long
as an experienced team is restraining the animals to avoid injuries, excessive stress levels
and other complications (Hernandez, 2014; Shury, 2014; Goodman et al., 2013; Atkinson et
al., 2012; La Grange, 2012; Fivaz & Ebedes, 2012; Bothma & Van Rooyen, 2005).
However, most of the procedures carried out required a chemical immobilization.
Each individual protocol and respective doses administered were mainly based on the
personal experience of the veterinarian as opposed to data from academic references. The
differences can be seen in the comparisons of the protocols and respective doses with the
references on Appendix 2. (Table 20-34).
By analyzing the results from the chemical immobilizations performed (184), it is
possible to affirm that most of the clinical cases were well succeeded, without any
requirement of additional darts (171/184: 92.93%). Only a small percentage, 7.07% (13/184)
of the procedures needed a second dart dose.
A higher administration of top-up anesthetics to improve the depth of the
anesthesia was required in some cases. However, as was already expected, during surgery,
particularly in surgical cases involving felids, the administration of drugs to maintain the
anesthesia was common to ensure a safer procedure for the animals and for the team
involved. Additional drugs were also provided in specific cases after a first cocktail of drugs
were darted (e.g. administration of midazolam in some of the African buffalos), due to
individual sensitivity to the drugs, environmental stimulation or inadequate IM administration
through the dart. Some of the darts might have been SQ, hence interfering with the
appropriate absorption of the drugs by the organism by slowing down the induction and
decreasing the depth of the anesthesia. For these reasons, although a second dart dose was
required, the immobilizations of 3 African buffalos, 2 roan antelopes and 2 lionesses were not
classified as unsuccessful because the issue was easily identified and corrected without any
associated complications. With that in mind, there were 176 cases from the 184 chemical
immobilizations classified as 'well succeeded' (95.65%) and only 8 (4.35%) 'not succeeded'
(individuals who required a second darting plus several top-up administrations or individuals
that died during the procedures).
Most authors of scientific publications describe the doses of the drugs for
ungulates as a net dosage for the species and only specify the dose according to the
sex/approximate age of the animals (e.g. males, females, juveniles, calves). The doses for
ungulates are not measured according to body weight, like, for example, with carnivores
(Miller & Fowler, 2015; West, Heard & Caulkett, 2014; Kock & Burroughs, 2012 – Appendix
2. Table 20-34). Therefore the results presented on Chapter 3 tables for ungulates are also
described as net doses not weight dependent.
In certain cases, such as the African buffalos, the protocol performed was based
entirely on the experience of the veterinarian in charge and, as far as the author of this thesis
is aware, the combination of etorphine with thiafentanil is not referred to in the literature for
this particular species. The practitioner's years of experience allowed an improvement on the
doses applied to the individuals of this species. The choice of the dart gun device (Pneu-Dart
type C) was also based on the experience of the operator, plus the environmental conditions
in the field. The characteristics of the darts were based on the volume of the drugs and
thickness of the animal's skin (IM injection is preferable). WB needles allowed the dart to
stay in place to facilitate the administration of the total volume of drugs.
The administration of medetomidine on some bulls and cows is associated not
only with the size of the animal but also with the difficult conditions of the field (mountains,
water courses, thick bush) which required a faster immobilization of the animals to avoid
future complications after darting. However, the administration of atipazemole IM is always
recommended to reverse the effects of medetomidine, which can also be used as a top-up
drug when the animal does not show adequate signs of sedation. Midazolam, rather than
medetomidine, was also often used to increase the muscle relaxation and improve the
immobilization of buffalos. This particular drug is the top-up drug of choice by the veterinarian
in charge of these procedures instead of ketamine, which has been commonly used in the
past and well referred to in publications (Wolfe, 2015; Ball & Hormeyr, 2014; Lamont &
Grimm, 2014; Burroughs et al., 2012b). As a dissociative agent, ketamine does not have any
reversal drug and it presents more side-effects than midazolam, which can last up to 2-3
hours after administration and is excessively long for most of the usual procedures
performed (Lamont & Grimm, 2014; Plumb, 2008; Caulkett & Arnemo, 2007).
The immobilizations of sable and roan antelopes also required administration of
midazolam through the auricular vein to improve muscle relaxation. Medetomidine, also used
in these antelope species, is referenced by a few practitioners (Wolfe, 2015; Ball & Hormeyr,
2014; Citino, Bush, Grobler & Lance, 2001), and it was always added to the darting cocktail
for the roan antelopes. The characteristics of the darts used were also based on veterinarian
experience. Larger antelope species required longer lengths when compared to smaller
antelopes immobilized, such as bonteboks or springboks, which are targetable with darts of
small volumes and lengths. It would be possible to use WB instead of the GC darts on the
bontebok, but, for the volume of drugs required, the only available darts at the time were the
GC.
The 2 male golden wildebeests restrained were administered with LAN –
zuclopenthixol – to improve the tranquilization during the transportation of the animals. After
the effect of the azaperone subsided, the animals required additional tranquilizers to improve
not only the transportation but also the adaptation to the new environment (Wolfe, 2015;
Read et al., 2000).
The plains zebras were successfully immobilized with etorphine (and azaperone),
which is the best opioid for this particular species according to the references from the main
wildlife veterinarians (Janssen & Allen, 2015; West et al., 2014; Walzer, 2014; Kock &
Burroughs, 2012).
Although the immobilization of the ram bushbuck was an emergency, the protocol
chosen (thiafentanil and medetomidine), and the dart gun/darts (Pneu Dart type C, 1cc
1''GC) worked successfully and restraint was accomplished. However, to guarantee a more
efficient drug delivery to the animal, a WB dart would have been a safer choice. However, at
the time a dart with a low volume, short length and WB was not available.
The delivery system manufacturer used for the darting was mainly the Pneu Dart,
with some variations between type P and type C. Type C was used almost exclusively for
African buffalos because of the veterinarian's experience with that type of gun/darts in that
particular species, or for animals shot from a helicopter, such as the giraffes or the free-
ranging bull elephant. The other manufacturer, Dan Inject, was only used for the
immobilization of elephants in controlled environments to perform a standing sedation of the
individuals. They were darted from the ground and, because of the quieter sound of the Dan
Inject gun, in comparison to the Pneu Dart, the animals were less stressed after the shooting,
allowing a smoother standing sedation.
Results obtained from the different immobilizations performed during the
internship show that only 3 of the 245 animals immobilized died: 1 female springbok (which
ran into a lake and drowned during the excitement phase); 1 female giraffe (which fell into a
mud pit and was euthanized after a few hours of unsuccessful attempts by personnel to pull it
out, and after showing signs of inappropriate muscular function on its posterior legs); and 1
female white rhino (which was extremely sensitive to the immobilization drugs). This female
rhino was accurately darted with a rigorous cocktail of drugs because of historical problems
with anesthetic drugs in its ancestors. However, the first dart was not efficient and the drugs
were not administered on the animal. In order to immobilize the female to perform horn
trimming, a preventive method for poaching issues in the country, a second dart with the
same combination of drugs and doses was fired. Unfortunately, and despite all attempts to
reverse the anesthesia, the rhino died.
The female giraffe was not using her posterior legs and, although no bone
fracture was suspected, the muscles were too weak for the giraffe to hoist herself out of the
pit. The most probable diagnostic was myopathy after a few hours of constant attempts by
the team to pull the giraffe out. The entire capture procedure went as normal and the
problem was a difficult one to prevent. One conclusion drawn from this incident is that it is
always good practice to have a helicopter pilot with experience on hand to help prevent
situations such as this. The other clinical cases with giraffes went faultlessly. The darts were
only filled with opioids in high doses to guarantee a faster sedation of the animals and
avoiding long excitement phases. Additionally, the darts had hyaluronidase powder to
improve the fast action of the opioids in the organism (Morkel, 1993; Morkel, 1992). These
opioids must be quickly reversed with the antidotes (naltrexone) immediately after the giraffe
falls to the ground, otherwise high concentrations of opioids can be lethal to these animals
(Bertelsen, 2015; Citino & Bush, 2014; Burroughs et al., 2012b). Then, the procedure must
be carried out with the animal still on the ground and physically controlled by the team,
which, in our study, had to walk the giraffes directly to the transport vehicle (Figure 14).
Figure 14 – Female giraffe guided by rangers into a transportation vehicle,
after the reversal agent has been administered (Original).
Potential complications during wildlife immobilizations might warrant a previous
discussion between the veterinarian in charge and the personnel involved in the restraint, to
assess whether deaths can be expected during the operation. Based on the fact that 10% is
an acceptable rate when the practice is being performed in species without a conservation
value concern, a lower percentage of losses must be expected for threatened species
(Hernandez, 2014). Out of all 245 cases, only 3 (1.2%) resulted in the loss of individuals.
Two of those individuals were classified as LC species and the white rhino as NT, which
means that they are not threatened species according to the IUCN Red List of Threatened
Species (http://www.iucnredlist.org). Thus the percentage lower than 10% referred by
Hernandez (2014) can be compared to the 1.2% of losses in our study. These results confirm
the efficiency and professionalism during the operations. However, comparing the chemical
restraints performed in the 245 clinical cases examined, 184 were chemical immobilizations,
3 of which resulted in deaths. This equals 1.6%, which is still a much lower percentage,
reinforcing the high standards of the methods, protocols and doses chosen.
Besides the 3 death cases, an attempt to immobilize a free-ranging bull elephant
for a standing sedation was also unsuccessful although it was the first standing sedation
attempt on a free-ranging bull African elephant. In this case, the drug cocktail in the dart was
not strong enough to restrain the bull and a second dart with a new cocktail of drugs was
given based on drugs and respective doses for a free-ranging bull knockdown
immobilization. For the standing sedation of bull elephants in a controlled environment, one
of the immobilizations required an additional administration of drugs to improve the depth of
the anesthesia during the restraint as no pure opioids like etorphine were administered.
In carnivores, most of the procedures were well succeeded, except the male
leopard, the male cheetah and one of the female painted dogs. The black leopard case was
considered unsuccessful due to the second darting and the various top-ups required before
performing surgery. All the surgeries in felids (hemi-hysterectomies, ophthalmological
procedures, castrations and epididymectomies) were performed without severe
complications. The only requirement was the administration of medetomidine to improve the
depth of the anesthesia, which was always reversed with atipamezole, and/or TZ, as is
recommended by wildlife practitioners (Lamberski, 2015; Ramsay, 2014; Burroughs et al.,
2012b; Jacquier, Aarhaug, Arnemo, Bauer & Enriquez, 2006; Fahlman et al., 2005). No
further problems arose after these procedures, not even in the free-ranging individuals
released into a non-controlled environment. The castration was performed on a male black
leopard and the epididymectomies were carried out on male African lions because of the
inherent difficulties of performing a vasectomy. The hemi-hysterectomies, performed on the
lionesses (Figure 15 and 16) allow the free-ranging females to give birth every year, which is
a very attractive feature for tourists and a significant help to the worldwide survival of the
species without overloading a wildlife eco-reserve with too many predators. In the controlled
facilities, the reproductive issues were managed with SQ GnRH implants.
Figure 15 and 16 – African lioness being prepared for surgery (left) and hemi-hysterectomy
(right) in Gondwana Game Reserve (Original).
Although the male cheetah and one of the female painted dogs immobilized were
classified as 'not succeeded' cases because of the requirement of a second dart dose plus
the administration of top-up drugs to improve the depth of anesthesia (possible incorrect
evaluation of body weight), all re-locations were well accomplished. The individuals were
supplemented and regularly checked during transportation to prevent severe dehydration
and ensure their welfare. Given that these two species in particular are classified as
Vulnerable (Cheetah) and Endangered (Painted dog), they were among the threatened
individuals examined during the internship. The role of the veterinarian with individuals of
these species is vital, particularly during outbreaks of diseases like rabies, which can have a
huge impact on the free-ranging populations (Padilla & Hilton, 2015; Lamberski, 2015;
Ramsay, 2014; Good, Marobela, & Houser, 2005; Turnbull et al., 2004; Hofmeyr, Bingham,
Lane, Ide & Nel, 2000; Jäger, Booker, & Hübschle, 1990).
As protocols for each medical procedure can be influenced by SECONDS
(Blumstein et al., 2015; Wolfe, 2015; Hernandez, 2014; Pas, 2014; Paterson, 2014; Hofmeyr
et al., 2012; Sanchez, 2011), the experience of the veterinarian in charge is a crucial factor in
choosing the most appropriate combination of drugs and respective doses. It is also vital to
deciding the best delivery system according to the situation and the most efficient reversal,
preventing possible complications during the restraint and turning it into an efficient and safe
The majority of the immobilizations performed were well accomplished for both
ungulates and carnivores and did not require any additional drugs administered to the
animals. Percentage-wise the losses (1.2%) are well below the acceptable 10%, and no
Threatened species were lost. However, besides the positive balance of the restraints
performed, this was only possible thanks to the experience of the veterinarian in charge and
the experienced personnel involved.
There are many obstacles to this particular area of veterinary medicine and the
practitioners must have the necessary qualifications and experience to be able to coordinate
the team involved in field procedures and predict any possible complications during the
restraint event. The success of a clinical case relies on the fore planning of the procedure
and the prediction of potential problems. This is a transversal situation in all fields of
veterinary practice. However, in wildlife medicine, practitioners are often examining species
on the brink of extinction and dealing with conservational status concerns. Failures or
mistakes that can be prevented are inadmissible.
The chemical restraint protocols described by the different authors Miller &
Fowler, 2015; West et al., 2014; Kock & Burroughs, 2012 (see Appendix 2. Tables 20-34)
and the ones performed during the internship present several unconformities in terms of the
cocktails used for immobilization, doses and reversal options. The lack of information about
the characteristics of the darts and respective dart guns that can be used to capture the
different species is also an important aspect to be taken into consideration, especially given
the versatility of procedures and situations that can arise, and which the authors rarely
describe in scientific publications.
The veterinarian's experience in SECONDS dictates the different approaches
that will ultimately contribute to the constant evolution of veterinary practice. Most of the
academic compilations available do not clarify the factors that might influence the procedures
(SECONDS) nor provide a protocol or specific doses for the species to which they are
referring.
It is also important to note that some of the cocktail options presented in
publications are only based on one or a few immobilizations and, most of the time, involve
the same drug combinations and doses for free-ranging individuals as for individuals in a
controlled facility (e.g. zoo) (Miller & Fowler, 2015; West et al., 2014).
It is vital that the experience of the different wildlife practitioners becomes
standardized and is shared among veterinarians worldwide. Even data information about
mistakes and cases that failed, complications observed in unusual situations and erroneous
restraints performed in different scenarios, are fundamental to avoid as many losses as
possible in the future. All relevant information collected must be reported.
Various global threats compromise the perspectives of survival for certain African
species. However, it is important that all veterinarians who intervene in free-ranging animals
or in a zoological environment know exactly what they are dealing with, without contributing
towards the deterioration of the condition of the species worldwide. This new role of the
wildlife practitioner has already been referred to by Deem, 2015; Mazet, Hamilton & Dierauf,
2006 and Kock, Soorae & Mohammed, 2007, which define the jobs of future wildlife
veterinarians as being consistent not only with medicine but also with conservation. New
data information about successful or unsuccessful performances (in such as anesthetic
protocols used during the internship, surgeries performed and even restraint techniques
executed for certain species) should also be taken into account and published to promote the
growth of knowledge in this branch of veterinary medicine. Constant improvement is the key
that allows a more efficient and safer medical care for every single individual; whether free-
ranging or in a zoo/controlled environment and, ultimately, will benefit the population
worldwide.
Personally, I must reinforce the importance of the internship from which this
thesis originated. This work allowed me to understand how challenging the veterinary
practice can be, particularly working with wildlife and all the constraints of practicing it among
free-ranging populations in remote areas with unpredictable logistic complications and
environmental obstacles. It was not only vital for my academic education but was also an
enriching experience and a privilege to be able to help what are among the most incredible
species in the world, some of them almost on the brink of extinction.
Allen, J. L. (1996). A comparison of nalmefene and naltrexone for the prevention of
renarcotization following carfentanil immobilization of nondomestic ungulates. Journal
of Zoo and Wildlife Medicine, 496-500.
Anderson, E. C., Doughty, W. J., Anderson, J., Paling, R. (1979). The pathogenesis of foot-
and-mouth disease in the African buffalo (Syncerus caffer) and the role of this species
in the epidemiology of the disease in Kenya. Journal of Comparative Pathology, 89 (4),
Anderson, E. C., Foggin, C., Atkinson, M., Sorensen, K. J., Madekurozva, R. L., Nqindi, J.
(1993). The role of wild animals, other than buffalo, in the current epidemiology of foot-
and-mouth disease in Zimbabwe. Epidemiology and Infection, 111 (03), 559-564.
Anderson, K., Ezenwa, V. O., Jolles, A. E. (2013). Tick infestation patterns in free ranging
African buffalo (Syncercus caffer): effects of host innate immunity and niche
segregation among tick species. International Journal for Parasitology: Parasites and
Wildlife, 2, 1-9.
Arnemo, J. M., Ahlqvist, P., Andersen, R., Berntsen, F., Ericsson, G., Odden, J., Sven
Brunberg, S., Segerström, P., Swenson, J. E. (2006). Risk of capture-related mortality
in large free-ranging mammals: experiences from Scandinavia. Wildlife Biology, 12 (1),
Arnemo, J. M., Evans, A. L., Fahlman, A., Caulkett, N. (2014). Field emergencies and
complications In West, G., Heard, D., Caulkett, N. (eds) Zoo animal and wildlife
immobilization and anesthesia. (2nd Ed.). Ames, Iowa, USA: Wiley Blackwell
Publishing, 139-148.
Atkinson, M., Kock, M., Meltzer, D. (2012). Principles of chemical and physical restraint of
wild animals In Kock, M., Burroughs, R. (eds) Chemical and physical restraint of wild
animals – a training and field manual for african species. (2nd Ed.) International Wildlife
Veterinary Services, 103-130.
Ayebazibwe, C., Mwiine, F., Tjørnehøj, K., Balinda, S., Muwanika, V., Okurut, A., Belsham,
G., Normann, P., Siegismund, H., Alexandersen, S. (2010). The role of African buffalos
(Syncerus caffer) in the maintenance of foot-and-mouth disease in Uganda. BMC
Veterinary Research, 6 (1), 1.
Ball, R. L., Hormeyr, M. (2014). Antelope In West, G., Heard, D., Caulkett, N. (eds) Zoo
animal and wildlife immobilization and anesthesia. (2nd Ed.) Ames, Iowa, USA: Wiley
Blackwell Publishing, 831-842.
Bauer, H., Packer, C., Funston, P.F., Henschel, P., Nowell, K. (2015). Panthera leo. The
IUCN Red List of Threatened Species 2015: e.T15951A79929984.
http://dx.doi.org/10.2305/IUCN.UK.2015-4.RLTS.T15951A79929984.en. Downloaded
on 23 January 2016.
Beechler, B. R., Broughton, H., Bell, A., Ezenwa, V. O., Jolles, A. E. (2012). Innate immunity
in free-ranging African buffalo (Syncerus caffer): associations with parasite infection
and white blood cell counts. Physiological and Biochemical Zoology, 85 (3), 255-264.
Bertelsen, M. E. (2015) Giraffidae In Miller, R. E. & Fowler, M. E.; Zoo and Wild animal
medicine. (Volume 8). St Louis, Missouri: Elsevier Saunders, 602-610.
Blanc, J. (2008). Loxodonta africana. The IUCN Red List of Threatened Species 2008:
Downloaded on 23 January 2016.
Blumstein, D. T., Buckner, J., Shah, S., Patel, S., Alfaro, M. E., & Natterson-Horowitz, B.
(2015). The evolution of capture myopathy in hooved mammals: a model for human
stress cardiomyopathy? Evolution, Medicine, and Public Health, 2015(1), 195-203.
Bothma, J. D. P., Van Rooyen, N. (2005). Intensive wildlife production in Southern Africa.
Van Schaik Publishers.
Burroughs, M., Hofmeyr, M., Morkel, P., Kock, M., Kock, R., Meltzer, D. (2012b) Chemical
immobilization – individual species requirements In Kock, M., Burroughs, R. (eds)
Chemical and physical restraint of wild animals – a training and field manual for african
species. (2nd Ed.) International Wildlife Veterinary Services, 143-264.
Burroughs, R., Meltzer, D., Morkel, P. (2012a). Applied Pharmacology In Kock, M.,
Burroughs, R. Chemical and physical restraint of wild animals – a training and field
manual for african species. (2nd Ed.). International Wildlife Veterinary Services. 53-80.
Burroughs, R. E. J., & McKenzie, A. A. (1993). Handling, care, and loading of immobilized
herbivores. The capture and care manual. Wildlife Decision Support Services and the
South African Veterinary Foundation, Pretoria, 184-191.
Bush, M., de Vos, V. (1987). Observations on field immobilization of free-ranging giraffe
(Giraffa camelopardalis) using carfentanil and xylazine. The Journal of Zoo Animal
Medicine, 18 (4), 135-140.
Bush, M. (1976). Giraffe restraint and immobilization. In Proceedings of the American
Association of Zoo Veterinarians, 151-154.
Bush, M. (1993). Anesthesia of high-risk animals: giraffe. Zoo and Wild Animal Medicine,
Bush, M. (2003). Giraffidae. In Zoo and Wild Animal Medicine (5th Ed.), Fowler, M.E.; Miller,
R. E., eds. St. Louis, Saunders, 625-633.
Bush, M., Ensley, P. K., Mehren K, Rapley W. (1976). Giraffe immobilization utilizing
Xylazine and etorphione hydrochloride. Journal of American Veterinary Medical
Association, 169.
Bush, M., Grobler, D. G., Raath, J. P., Phillips Jr, L. G., Stamper, M. A., Lance, W. R. (2001).
Use of medetomidine and ketamine for immobilization of free-ranging giraffes. Journal
of the American Veterinary Medical Association, 218 (2), 245-249.
Bush, M., Raath, J. P., Grobler, D., Klein, L. (2004). Severe hypoxaemia in field-
anaesthetised white rhinoceros (Ceratotherium simum) and effects of using tracheal
insufflation of oxygen. Journal of the South African Veterinary Association, 75 (2), 79-
Bush, M., Raath, C., Phillips, L. G., Lance, W. (1997). Immobilization of free-ranging giraffe
(Giraffa camelopardalis) using medetomidine and ketamine. In Proceedings of the
American Association of Zoo Veterinarians Annual Conference, 276-7.
Cattet, M., Boulanger, J., Stenhouse, G., Powell, R. A., Reynolds-Hogland, M. J. (2008a). An
evaluation of long-term capture effects in ursids: implications for wildlife welfare and
research. Journal of Mammalogy, 89 (4), 973-990.
Cattet, M., Stenhouse, G., Bollinger, T. (2008b). Exertional myopathy in a grizzly bear (Ursus
arctos) captured by leghold snare. Journal of wildlife diseases, 44 (4), 973-978.
Cattet, M. R., Obbard, M. E. (2010). Use of hyaluronidase to improve chemical
immobilization of free-ranging polar bears (Ursus maritimus). Journal of wildlife
diseases, 46 (1), 246-250.
Caulkett, N., Shury, T. (2014). Human safety during wildlife capture In West, G., Heard, D.,
Caulkett, N. (eds) Zoo animal and wildlife immobilization and anesthesia. (2nd Ed.).
Ames, Iowa, USA: Wiley Blackwell Publishing, 181-190.
Caulkett, N. A., Arnemo, J. M. (2007). Chemical Immobilization of Free-Ranging Terrestrial
Mammals. In Tranquilli, W. J., Thurmon, J. C.,Grimm, K. A. (eds.), Lumb & Jones'
Veterinary Anesthesia and Analgesia (4th ed.). Ames, Iowa, USA: Blackwell, 807-832.
Chaisi, M. E., Sibeko, K. P., Collins, N. E., Potgieter, F. T., Oosthuizen, M. C. (2011).
Identification of Theileria parva and Theileria sp. (buffalo) 18S rRNA gene sequence
variants in the African Buffalo (Syncerus caffer) in southern Africa. Veterinary
parasitology, 182 (2), 150-162.
Clarke, K. W., Trim, C. M., Hall, L. W. (2014). Veterinary Anaesthesia. (11 th Ed.) Elsevier
Saunders: St. Louis, MO, 10-244.
Citino, S. B. & Bush, M. (2014). Giraffidae In West, G., Heard, D., Caulkett, N. (eds) Zoo
Citino, S. B., Bush, M., Grobler, D., Lance, W. (2001). Anaesthesia of roan antelope
(Hippotragus equinus) with a combination of A3080, medetomidine and ketamine.
Journal of the South African Veterinary Association, 72 (1), 29-32.
Citino, S. B., Bush, M., Lance, W., Hofmeyer, M., Grobler, D. (2006). Use of thiafentanil
(A3080), medetomidine and ketamine for anesthesia of captive and free-ranging giraffe
(Giraffe camelopardalis). In Proceedings American Association of Zoo Veterinarians,
Courchamp, F., Rasmussen, G. S., Macdonald, D. W. (2002). Small pack size imposes a
trade-off between hunting and pup-guarding in the painted hunting dog Lycaon pictus.
Behavioral Ecology, 13 (1), 20-27.
Cracknell, J. (2014). Airway management In West, G.; Heard, D.; Caulkett, N.; Zoo animal
and wildlife immobilization and anesthesia. (2nd Ed.) Ames, Iowa, USA: Wiley Blackwell
Publishing, 53-64.
Crosier, A. E., Marker, L., Howard, J., Pukazhenthi, B. S., Henghali, J. N., Wildt, D. E.
(2007). Ejaculate traits in the Namibian cheetah (Acinonyx jubatus): influence of age,
season and captivity. Reproduction, Fertility and Development, 19 (2), 370-382.
De Klerk, L., Michel, A.L., Grobler, D.G., Bengis, R., Bush, M., Kriek, N.P.J., Hofmeyr, M.S.,
Griffin, J.F.T., Mackintosh, C.G. (2006). An experimental intratonsilar infection model
for bovine tuberculosis in African buffaloes, Syncerus caffer. The Onderstepoort
Journal of Veterinary Research, 73, 293 – 303.
De Villiers, M. S., van Jaarsveld, A. S., Meltzer, D. G., Richardson, P. R. (1997). Social
dynamics and the cortisol response to immobilization stress of the African wild dog,
Lycaon pictus. Hormones and Behavior, 31(1), 3-14.
Deem, S. (2015). Conservation Medicine to One Health In Miller, R. E., Fowler, M. E. (eds)
Zoo and Wild Animal Medicine. (Volume 8) St Louis, Missouri: Elsevier Saunders, 698-
Diverio, S., Goddard, P. J., Gordon, I. J. (1996). Physiological responses of farmed red deer
to management practices and their modulation by long-acting neuroleptics. The Journal
of Agricultural Science, 126 (02), 211-220.
Du Toit, J. G. (2001). Veterinary care of African elephants. South African Veterinary
Foundation, 8-60.
Durant, S., Mitchell, N., Ipavec, A., Groom, R. (2015). Acinonyx jubatus. The IUCN Red List
of Threatened Species 2015: e.T219A50649567.
http://dx.doi.org/10.2305/IUCN.UK.2015-4.RLTS.T219A50649567.en. Downloaded on
23 January 2016.
Emslie, R. 2012. Ceratotherium simum. The IUCN Red List of Threatened Species 2012:
e.T4185A16980466.
http://dx.doi.org/10.2305/IUCN.UK.2012.RLTS.T4185A16980466.en. Downloaded on
Fahlman, A. (2008). Advances in Wildlife Immobilization and Anaesthesia: Clinical and
Physiological Evaluation in Selected Species. Doctoral Thesis. Uppsala: Swedish
University of Agricultural Sciences, 13-58.
Fahlman, A. (2014). Oxygen therapy In West, G., Heard, D., Caulkett, N. (eds) Zoo animal
and wildlife immobilization and anesthesia. (2nd Ed) Ames, Iowa, USA: Wiley Blackwell
Fahlman Å., Loveridge A., Wenham C., Foggin C., Arnemo J.M., Nyman G. (2005).
Reversible anaesthesia of free-ranging lions (Panthera leo) in Zimbabwe. Journal of
the South African Veterinary Association, 76 (4), 187–192.
Fennessy, J., Brown, D. 2010. Giraffa camelopardalis. The IUCN Red List of Threatened
Species 2010: e.T9194A12968471. http://dx.doi.org/10.2305/IUCN.UK.2010-
2.RLTS.T9194A12968471.en. Downloaded on 23 January 2016.
Fick, L., Mitchell, D., Fuller, A. (2007). Long-acting neuroleptics used in wildlife management
do not impair thermoregulation or physical activity in goats (Capra hircus). Comparative
Biochemistry and Physiology Part A: Molecular & Integrative Physiology, 147 (2), 445-
Fivaz, B., Ebedes, H. (2012). Post-capture boma management In Kock, M., Burroughs, R.
(eds) Chemical and physical restraint of wild animals – a training and field manual for
african species. (2nd Ed.). International Wildlife Veterinary Services, 331-338.
Foggin, C., Masterson, C., Hoare, R. (2012). Legal and ethical considerations in the use of
immobilizing drugs, using Zimbabwe and South Africa as examples In Kock, M.,
Burroughs, R. (eds) Chemical and physical restraint of wild animals – a training and
field manual for african species. (2nd Ed.). International Wildlife Veterinary Services, 1-
Fowler, M. E., Mikota, S. K. (2006). Frontmatter, In Biology, Medicine, and Surgery of
Elephants, Blackwell Publishing Ltd, Oxford, UK, 91-118.
Fowler, M.E. (1995). Restraint and Handling of Wild and Domestic Animals (2nd ed.). Ames,
Iowa, USA: Iowa State University Press.
Fowler, M.E. (2008). Restraint and handling of wild and domestic animals. (3th ed). Iowa:
Blackwell Publishing.
Good, K. M., Marobela, C., & Houser, A. M. (2005). A report of anthrax in cheetahs
(Acinonyx jubatus) in Botswana: to the editor. Journal of the South African Veterinary
Association, 76(4), 186.
Goodman, G., Hedley, J., Meredith, A. (2013). Field techniques in zoo and wildlife
conservation work. Journal of Exotic Pet Medicine, 22 (1), 58-64.
Hack, M.A, Lorenzen, E. (2008). Equus quagga. The IUCN Red List of Threatened Species
2008: e.T41013A10386901.
http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T41013A10386901.en. Downloaded on
Hartup, B. K., Kollias, G. V., Jacobsen, M. C., Valentine, B. A., & Kimber, K. R. (1999).
Exertional myopathy in translocated river otters from New York. Journal of Wildlife
Diseases, 35(3), 542-547.
Hedger, R. S. (1972). Foot-and-mouth disease and the African buffalo (Syncerus caffer).
Journal of Comparative Pathology, 82 (1), 19-28.
Henschel, P., Hunter, L., Breitenmoser, U., Purchase, N., Packer, C., Khorozyan, I., Bauer,
H., Marker, L., Sogbohossou, E. & Breitenmoser-Wursten, C. 2008. Panthera pardus.
The IUCN Red List of Threatened Species 2008: e.T15954A5329380.
http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T15954A5329380.en.
Downloaded on 03 June 2016.
Hernandez, S. (2014). Chemical immobilization of wild animals In Clarke, K. W., Trim, C. M.,
Hall, L. W. (eds) Veterinary Anaesthesia. (11th Ed.) Saunders Elsevier, 571-584.
Hofmeyr, J. M. (1981). The use of haloperidol as a long-acting neuroleptic in game capture
operations. Journal of the South African Veterinary Association, 52 (4), 273-282.
Hofmeyr, M., Bingham, J., Lane, E. P., Ide, A., Nel, L. (2000). Rabies in African wild dogs
(Lycaon pictus) in the Madikwe Game Reserve, South Africa. Veterinary Record, 146
(2), 50-52.
Hofmeyr, M., Fivaz, B., Meltzer, D. (2012). Ancillary treatments in wildlife capture and care In
Kock, M., Burroughs, R. (eds) Chemical and physical restraint of wild animals – a
training and field manual for african species. (2nd Ed.). International Wildlife Veterinary
Services, 311-318.
Horne, W. A., Loomis, M. R. (2014). Elephants In West, G., Heard, D., Caulkett, N. (eds) Zoo
Animal and Wildlife Immobilization and Anesthesia. (2nd Ed.). Ames, Iowa, USA: Wiley
http://www.iucnredlist.org Downloaded on 23 January 2016.
Isaza, R. (2014). Remote drug delivery In West, G., Heard, D., Caulkett, N. (eds) Zoo Animal
and Wildlife Immobilization and Anesthesia. (2nd Ed.). Ames, Iowa, USA: Wiley
IUCN SSC Antelope Specialist Group. (2008a). Antidorcas marsupialis. The IUCN Red List
of Threatened Species 2008: e.T1676A6359255.
http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T1676A6359255.en.
IUCN SSC Antelope Specialist Group. (2008b). Connochaetes taurinus. The IUCN Red List
of Threatened Species 2008: e.T5229A11120316.
IUCN SSC Antelope Specialist Group. (2008c). Hippotragus equinus. The IUCN Red List of
Threatened Species 2008: e.T10167A3178972.
http://dx.doi.org/10.2305/IUCN.UK.2008.RLTS.T10167A3178972.en. Downloaded on
IUCN SSC Antelope Specialist Group. (2008d). Hippotragus niger. The IUCN Red List of
IUCN SSC Antelope Specialist Group. (2008e). Oryx gazella. The IUCN Red List of
IUCN SSC Antelope Specialist Group. (2008f). Philantomba monticola. The IUCN Red List of
Threatened Species 2008: e.T4143A10469452.
IUCN SSC Antelope Specialist Group. (2008g). Syncerus caffer. The IUCN Red List of
IUCN SSC Antelope Specialist Group. (2008h). Tragelaphus scriptus. The IUCN Red
List of Threatened Species 2008: e.T22051A9352536.
IUCN/SSC Cat Specialist Group. (2015). Panthera leo. http://catsg.org/index.php?id=108,
Jacques, C. N., Jenks, J. A., Deperno, C. S., Sievers, J. D., Grovenburg, T. W., Brinkman, T.
J., Swanson, C. C., Stillings, B. A. (2009). Evaluating Ungulate Mortality Associated
With Helicopter Net‐Gun Captures in the Northern Great Plains. The Journal of Wildlife
Management, 73(8), 1282-1291.
Jacquier, M., Aarhaug, P., Arnemo, J. M., Bauer, H., Enriquez, B. (2006). Reversible
immobilization of free-ranging African lions (Panthera leo) with medetomidine-
tiletamine-zolazepam and atipamezole. Journal of wildlife diseases, 42 (2), 432-436.
Jäger, H. G., Booker, H. H., & Hübschle, O. J. (1990). Anthrax in cheetahs (Acinonyx
jubatus) in Namibia. Journal of wildlife diseases, 26(3), 423-424.
Janssen, D. L., Raath, J. P., De Vos, V., Swan, G. E., Jessup, D., Stanley, T. H. (1991). Field
studies with the narcotic immobilizing agent A3080. Proceeding American Association
of Zoo Veterinarians, 340–342.
Janssen, D.L., Allen, J. L. (2015). Equidae. Fowler's Zoo and Wild Animal Medicine, Volume
8, 559-567.
Jolles, A. E., Cooper, D. V., Levin, S. A. (2005). Hidden effects of chronic tuberculosis in
African buffalo. Ecology, 86 (9), 2358-2364.
Kalema-Zikusoka, G., Bengis, R. G., Michel, A. L., Woodford, M. H. (2005). A preliminary
investigation of tuberculosis and other diseases in African buffalo (Syncerus caffer) in
Queen Elizabeth National Park, Uganda. The Onderstepoort Journal of Veterinary
Research, 72 (2), 145.
Ko, J. C., Krimins, R. A. (2014). Thermoregulation In West, G., Heard, D., Caulkett, N. (eds)
Zoo animal and wildlife immobilization and anesthesia. (2nd ed). Ames, Iowa, USA:
Wiley Blackwell Publishing, 65-68.
Kock, M., Burroughs, R. (2012). Chemical and physical restraint of wild animals – a training
and field manual for african species. (2nd Ed.). International Wildlife Veterinary
Kock, M., Morkel, P. (2012). Other restraint tools In Kock, M., Burroughs, R. (eds) Chemical
and physical restraint of wild animals – a training and field manual for african species.
(2nd Ed.). International Wildlife Veterinary Services, 305-310.
Kock, M. (2012). Notes on developing and immobilizing kit for the field and the inclusion of
data sheets In Kock, M., Burroughs, R. Chemical and physical restraint of wild animals
– a training and field manual for african species. (2nd Ed.). International Wildlife
Kock, M. D., La Grange, M., du Toit, R. (1990). Chemical immobilization of free-ranging black
rhinoceros (Diceros bicornis) using combinations of etorphine (M99), fentanyl, and
xylazine. Journal of Zoo and Wildlife Medicine, 155-165.
Kock, M., Jessup, D., Burroughs, R. (2012a). Drug-injecting equipment In Kock, M.,
field manual for african species. (2nd Ed.) International Wildlife Veterinary Services,
Kock, M., Jessup, D., Burroughs, R. (2012b). Ballistics and projectile darting systems In
training and field manual for african species. (2nd Ed.) International Wildlife Veterinary
Kock, M., Jessup, D., Morkel, P. (2012c). Dart projectors In Kock, M., Burroughs, R. (eds)
species. (2nd Ed.). International Wildlife Veterinary Services, 287-304.
Kock, R. A., Soorae, P. S., & Mohammed, O. B. (2007). Role of veterinarians in
re‐introductions. International Zoo Yearbook, 41(1), 24-37.
Kohn, T. (2013). Capture Myopathy. Research Unit for Exercise Science and Sports
Medicine, University of Cape Town, WR SPRING 2013, 103-107.
La Grange, M. (2012). Overview of capture methods and transport of wild animals In Kock,
M., Burroughs, R. (eds) Chemical and physical restraint of wild animals – a training and
field manual for african species. (2nd Ed.). International Wildlife Veterinary Services,
Lamberski, N. (2015). Felidae In Miller, R. E., Fowler, M. E. (eds) Zoo and Wild Animal
Lamont, L. A., Grimm, K. A. (2014). Clinical Pharmacology In West, G., Heard, D., Caulkett,
N. (eds) Zoo animal and wildlife immobilization and anesthesia. (2nd Ed.) Ames, Iowa,
USA: Wiley Blackwell Publishing, 5-42.
Lance, W.R., Kenny, D.E. (2012). Thiafentanil Oxalate (A3080) in Nondomestic Ungulate
Species. In Miller, R.E., Fowler, M. (eds.), Fowler's Zoo and Wild Animal Medicine:
Current Therapy, Volume 7. St. Louis, Missouri, USA: Saunders, Elsevier, 589-595.
Larsen, R. S., Kreeger, T. J., West, G., Heard, D., Caulkett, N. (2007). Canids. Zoo animal
and wildlife immobilization and anesthesia, 395-407.
Macbeth, B. J., Cattet, M. R. L., Stenhouse, G. B., Gibeau, M. L., Janz, D. M. (2010). Hair
cortisol concentration as a noninvasive measure of long-term stress in free-ranging
grizzly bears (Ursus arctos): considerations with implications for other wildlife.
Canadian Journal of Zoology, 88 (10), 935-949.
Mazet, J. A., Hamilton, G. E., Dierauf, L. A. (2006). Educating veterinarians for careers in
free-ranging wildlife medicine and ecosystem health. Journal of Veterinary Medical
Education, 33(3), 352-360.
McTaggart, J., Kock, M., Hofmeyr, M. (2012). Helicopter and fixed-wing use in wildlife work
In Kock, M. & Burroughs, R. Chemical and physical restraint of wild animals – a training
Meltzer, D., Kock, N. (2012). Stress and Capture-related Death In Kock, M., Burroughs, R.
african species. (2nd Ed.). International Wildlife Veterinary Services, 81-102.
Miller, M. A., Buss, P. (2015). Rhinoceridae (Rhinoceroses) In Miller, R. E., Fowler, M. E.
(eds) Zoo and Wild animal medicine. (Volume 8). St Louis, Missouri: Elsevier
Saunders, 538-546.
Miller, R. E., Fowler, M. E. (2015). Zoo and Wild Animal Medicine. (Volume 8). St Louis,
Missouri: Elsevier Saunders.
Montané, J., Marco, I., López-Olvera, J., Perpiñán, D., Manteca, X., Lavín, S. (2003). Effects
of acepromazine on capture stress in roe deer (Capreolus capreolus). Journal of
Wildlife Diseases, 39(2), 375-386.
Morkel, P. (1992). Giraffe capture with etorphine HCl (M-99) and hyalase – a new approach.
The Use of Tranquilizers in Wildlife. Dept. of Agricultural Development, Sinoville
Printers, Pretoria, South Africa, 58-59.
Morkel, P. (1993). Chemical capture of the giraffe Giraffa camelopardalis. The Capture and
Care Manual: Capture, Care, Accommodation and Transportation of Wild African
Animals, 601-607.
Morkel, P. & Kock, M. (2012). Safety and first aid in the field – weapons, drugs and animals
In Kock, M., Burroughs, R. (eds) Chemical and physical restraint of wild animals – a
Services, 89-101.
Muir, W.W. (2007). Considerations for General Anesthesia. In Tranquilli, W. J., Thurmon, J.
C., Grimm, K. A. (eds.) Lumb & Jones' Veterinary Anesthesia and Analgesia (4th ed.).
Ames, Iowa, USA: Blackwell Publishing, 7-30.
Napier, J., Armstrong, D. L. (2014). Nondomestic cattle In West, G., Heard, D., Caulkett, N.
(eds) Zoo Animal and Wildlife Immobilization and Anesthesia. (2nd Ed.) Ames, Iowa,
USA: Wiley Blackwell Publishing, 863-872.
Neiffer, D.L., Miller, M.A., Weber, M., Stetter, M., Fontenot, D.K., Robbins, P.K., Pye, G.W.
(2005). Standing sedation in african elephants (Loxodonta africana) using detomidine–
butorphanol combinations. Journal of Zoo and Wildlife Medicine, 36 (2), 250-256.
Nielsen, L. (1999). Chemical Immobilization of Wild and Exotic Animals. Ames, Iowa, USA:
Iowa State University Press.
Oosthuizen, W. C. (2006). Chemical immobilization of African buffalo (Syncerus caffer) in
Kruger National Park: evaluating effects on survival and reproduction (Doctoral
Dissertation, University Of Pretoria, Pretoria), 1-25.
Orrell T. (custodian) (2016). ITIS Global: The Integrated Taxonomic Information System
(version Sep 2015). In Roskov Y., Abucay L., Orrell T., Nicolson D., Kunze T., Flann
C., Bailly N., Kirk P., Bourgoin T., DeWalt R.E., Decock W., De Wever A. (eds).
Species 2000 & ITIS Catalogue of Life, 23rd December 2015 Digital resource at
www.catalogueoflife.org/col. Species 2000: Naturalis, Leiden, the Netherlands. ISSN
2405-8858. Downloaded on 23 January 2016.
Ozeki, L., Caulkett, N. (2014). Monitoring In West, G., Heard, D., Caulkett, N. (eds) Zoo
Blackwell Publishing, 43-52.
Padilla, L. R., Hilton, C. D. (2015). Canidae In Miller, R. E., Fowler, M. E. (eds) Zoo and Wild
Animal Medicine. (Volume 8). St Louis, Missouri: Elsevier Saunders, 457-466.
Pas, A. (2014). Gazelle and small antelope In West, G., Heard, D., Caulkett, N. (eds) Zoo
Paterson, J. (2014). Capture Myopathy In West, G., Heard, D., Caulkett, N. (eds) Zoo Animal
Pienaar, R., Potgieter, F. T., Latif, A. A., Thekisoe, O. M., Mans, B. J. (2011). Mixed Theileria
infections in free-ranging buffalo herds: implications for diagnosing Theileria parva
infections in Cape buffalo (Syncerus caffer). Parasitology, 138 (07), 884-895.
Plumb, D. C. (2008). Plumb's Veterinary Drug Handbook, Pharma Vet. Inc., Stockholm, WI,
Raath, J. P. (1993). Chemical capture of the African elephant (Loxodonta africana) In The
Capture and Care Manual: Capture, Care, Accommodation and Transportation of Wild
African Mammals. Wildlife Decision Support Services and The South African Veterinary
Foundation Pretoria, 484-493.
Radcliffe, R. W., Morkel, P. (2014). Rhinoceroses In West, G., Heard, D., Caulkett, N. (eds)
Zoo Animal and Wildlife Immobilization and Anesthesia. (2nd Ed.). Ames, Iowa, USA:
Wiley Blackwell Publishing, 741-772.
Ramsay, E. (2000). Standing sedation and tranquilization in captive African elephants
(Loxodonta africana) In Annual Conference-American Association of Zoo Veterinarians.
American Association of Zoo Veterinarians; 1998, 111-114.
Ramsay, E. C. (2008). Use of Analgesics in Exotic Felids In Fowler, M. E., Miller, R. E. (eds).
Zoo and Wild Animal Medicine: Current Therapy (Vol. 6). Elsevier Health Sciences,
Ramsay, E. C. (2014). Felids In West, G., Heard, D., Caulkett, N. (eds) Zoo Animal and
Wildlife Immobilization and Anesthesia. (2nd Ed.). Ames, Iowa, USA: Wiley Blackwell
Read, M., Caulkett, N., McCallister, M. (2000). Evaluation of zuclopenthixol acetate to
decrease handling stress in wapiti. Journal of Wildlife Diseases, 36 (3), 450-459.
Sanchez, C. (2011). Complications during physical and chemical restraint – Capture
Myopathy. Chicago Zoological Society, Brookfield Zoo, 1-4.
Schumacher, J. (2008). Side Effects of Etorphine and Carfentanil in Nondomestic Hoofstock.
In Fowler, M.E., Miller, R.E. (eds.) Zoo and Wild Animal Medicine: Current Therapy,
Volume 6. St. Louis, Missouri, USA: Saunders, Elsevier, 455-461.
Scott, R., Kreeger, T. J. (2014). Canids In West, G., Heard, D., Caulkett, N. (eds) Zoo Animal
Sheriff, M. J., Dantzer, B., Delehanty, B., Palme, R., Boonstra, R. (2011). Measuring stress in
wildlife: techniques for quantifying glucocorticoids. Oecologia, 166 (4), 869-887.
Shury, T. (2014). Physical capture and restraint In West, G., Heard, D., Caulkett, N. (eds)
Sibeko, K. P., Oosthuizen, M. C., Collins, N. E., Geysen, D., Rambritch, N. E., Latif, A. A., ...
, Coetzer, J. A. (2008). Development and evaluation of a real-time polymerase chain
reaction test for the detection of Theileria parva infections in Cape buffalo (Syncerus
caffer) and cattle. Veterinary parasitology, 155(1), 37-48.
Spraker, T.R. (1993). Stress and capture myopathy in artiodactylids. In Fowler, M.E. (Ed.)
Zoo & Wild Animal Medicine: Current Therapy 3. Philadelphia, Pennsylvania, USA:
W.B. Saunders Company, 481-489
Stegmann, G. F., Jago, M. (2006). Cardiopulmonary effects of medetomidine or midazolam
in combination with ketamine or tiletamine/zolazepam for the immobilization of captive
cheetahs (Acinonyx jubatus). Journal of the South African Veterinary Association,
77(4), 205-209.
Swan, G.E. (1993). Drugs used for the immobilization, capture and translocation of wild
animals. In Andersen, M.D., et.al. (eds) The Capture and care Manual. Pretoria:
Wildlife Decision Support Services and The South African Veterinary Foundation, 14-
Tanner, M., Inlameia, O., Michel, A., Maxlhuza, G., Pondja, A., Fafetine, J., Marranangumbe,
A. S. (2015). Bovine tuberculosis and brucellosis in cattle and African buffalo in the
Limpopo National Park, Mozambique. Transboundary and emerging diseases, 62 (6),
Terio, K. A., Marker, L., Munson, L. (2004). Evidence for chronic stress in captive but not
free-ranging cheetahs (Acinonyx jubatus) based on adrenal morphology and function.
Journal of Wildlife Diseases, 40 (2), 259-266.
Thurmon, J. C., Short, C. E. (2007). History and overview of veterinary anesthesia. Tranquilli,
W.J.; Thurmon, J.C.; Kurt, A.G., Lumb & Jones Veterinary Anesthesia and Analgesia
(4th ed). Blackwell Publishing Professional, 3-6.
Turnbull, P. C. B., Tindall, B. W., Coetzee, J. D., Conradie, C. M., Bull, R. L., Lindeque, P.
M., & Huebschle, O. J. B. (2004). Vaccine-induced protection against anthrax in
cheetah (Acinonyx jubatus) and black rhinoceros (Diceros bicornis). Vaccine, 22(25),
Van der Walt, J. M., Nel, L. H., & Hoelzel, A. R. (2001). Characterization of major
histocompatibility complex DRB diversity in the endemic South African antelope
Damaliscus pygargus: a comparison in two subspecies with different demographic
histories. Molecular Ecology, 10(7), 1679-1688.
Van Heerden, J., Burroughs, R. E. J., Dauth, J., Dreyer, M. J. (1991). Immobilization of wild
dogs (Lycaon pictus) with a tiletamine hydrochloride/zolazepam hydrochloride
combination and subsequent evaluation of selected blood chemistry parameters.
Journal of wildlife diseases, 27(2), 225-229.
van Hoven, W. (2015). Private game reserves in southern Africa. In Institutional
Arrangements for Conservation, Development and Tourism in Eastern and Southern
Africa. Springer Netherlands, 101-118
van Wyk, A. M., Kotzé, A., Randi, E., & Dalton, D. L. (2013). A hybrid dilemma: a molecular
investigation of South African bontebok (Damaliscus pygargus pygargus) and blesbok
(Damaliscus pygargus phillipsi). Conservation Genetics, 14(3), 589-599.
Wack, R. F. (2003). Felidae. In Fowler, M. E., Miler, R. E. (eds) Zoo and Wildlife Medicine, 5th
ed. St. Louis: WB Saunders, 491- 501.
Walzer, C., Gerritsmann, H. (2015). Update on remote delivery and restraint equipment In
Miller, R. E., Fowler, M. E.; Zoo and Wild Animal Medicine. (Volume 8). St Louis,
Missouri: Elsevier Saunders, 730-732.
Walzer, C. (2014). Nondomestic equids In West, G., Heard, D., Caulkett, N. (eds) Zoo animal
Walzer, C., Huber, C. (2002). Partial antagonism of tiletamine-zolazepam anesthesia in
cheetah. Journal of wildlife diseases, 38 (2), 468-472.
West, G., Heard, D., Caulkett, N. (2014). Zoo animal and wildlife immobilization and
anesthesia. (2nd Ed.). Ames, Iowa, USA: Wiley Blackwell Publishing.
Wiedner, E. (2015). Proboscidea In Miller, R. E., Fowler, M. E. (eds) Zoo and Wild Animal
Wolfe, B. (2015). Bovidae (except sheep and goats) and antilocapridae In Miller, R. E.,
Fowler, M. E. (eds) Zoo and Wild Animal Medicine. (Volume 8). St Louis, Missouri:
Elsevier Saunders, 626-644.
Woodroffe, R., Sillero-Zubiri, C. (2012). Lycaon pictus. The IUCN Red List of Threatened
Species 2012: e.T12436A16711116.
APPENDIX 1.
Table 19 – Characteristics of the dart needles for the most commonly
immobilized African species, based on personal experience plus Dr Brendan Tindall's years
of experience.
SPECIES NEEDLE
Rhino 60mm collared/barbed; 2-3mm
Elephant diameter
Large 40-60mm collared/barbed; 2mm
Buffalo, Giraffe, Eland
herbivores diameter
Antelope (Sable, Roan, Gemsbok,
30-40 mm barbed; 2 mm diameter
Wildebeest, Kudu), Zebra
Medium Antelope (Impala, Springbok) 20-30mm collared/barbed; 1-2mm
Small 20mm collared/barbed; 1-1.5mm
Small antelopes (Duiker, Steenbok)
Large 25-35mm barbed; 1.5mm
Lion, Leopard, Cheetah, Painted dog
carnivores diameter
The next sequence of tables shows the different protocols referred to by the main
wildlife practitioners Kock & Burroughs, 2012; West et al., 2014 and Miller & Fowler, 2015 for
the chemically immobilized species during the internship. The weight of the species was
established according to the references from Miller & Fowler (2015).
Most of the doses published for ungulates are net doses in mg and not calculated
on the weight of the animal (mg/kg). This is because of the difficulties in evaluating the
weight of the animals, which can only really be achieved in a zoo environment, and
environmental conditions that might influence the procedure and, consequently, the doses.
Bearing in mind that most of the clinical cases in Chapter 3 were performed on free-ranging
ungulates, the doses were also described as a net dose for the various species.
Table 20 – Different anesthetic protocols for the African buffalo.
Kock & Burroughs, 2012 West et al., 2014 Miller & Fowler, 2015
(boma) Etorphine:5-8mg+Azaperone:40-
Bull 80mg or Xylazine:10mg/ (free-ranging) Xylazine: 0.2-0.4mg/kg / Etorphine:0.012mg/kg
(300- Etorphine:8-10mg + Azaperone:80-150mg or + Xylazine:0.1mg/kg / Etorphine:0.005-
900kg) Xylazine:70-90mg/Thiafentanil:6-10mg + 0.007mg/kg + Acepromazine:0.02-0.03mg/kg
Carfentanil: 0.005mg + Xylazine:0.05mg/kg /
Azaperone:50-100mg + Xylazine:0.08-0.18mg/kg /
Etorphine:0.015mg/kg + Xylazine:0.1-
(boma) Etorphine:5-8mg+ Azaperone:40- Etorphine:0.015mg/kg+Azaperone:0.15mg/kg /
0.15mg/kg / Thiafentanil:0.01-0.025mg/kg +
80mg or Xylazine:10mg/ Carfentanil:0.006-0.008mg/kg
Cow Medetomidine:0.05-0.1mg/kg
(free-ranging)Etorphine:6-10mg + /Carfentanil:0.002-0.005mg/kg +
(300-
Azaperone:80-100mg or Xylazine:40- Xylazine:0.028-0.05mg/kg /Thiafentanil:0.007-
900kg)
60mg/Thiafentanil:6-8mg + Azaperone:50- 0.014mg/kg + Azaperone:0.06-0.07mg/kg
Diprenorphine:16-25mg(bull)14-20mg(cow) if Naltrexone 50xEtorphine IV or
Diprenorphine:0.01-0.03mg/Kg (Etorphine) or
free-ranging or 15-20mg if in bomas / 30xThiafentanil IV or 100xCarfentanil IM or
Reversal Naltrexone: 0.07-0.8mg/Kg (Carfentanil or
Naltrexone 15xThiafentanil or 80- IV/ Atipamezole 5xMedetomidine or
Thiafentanil)
150mg(bull) 180-200mg(cow) if Thiafentanil 0.1mg/kg (Xylazine) IM
Table 21 – Different anesthetic protocols for the Sable antelope.
Etorphine:6-8mg or Thiafentanil:6-8mg + Carfentanil:0.015-0.02mg/kg +
Bull (190- Etorphine:4-5mg+ Azaperone:100mg/
Azaperone:80-100mg/ Medetomidine:2- Xylazine:0.15-0.2mg/kg/ Etorphine:0.015-
300Kg) Carfentanil:4.6mg + Xylazine:45mg
3mg/Detomidine:2-3mg/ Xylazine:10-15mg 0.025mg/kg + Xylazine:0.1-0.2mg/kg +
Cow Etorphine:4-5mg or Thiafentanil:4-6mg + Ketamine:0.15-0.2mg/kg /
Etorphine:3-4mg+ Azaperone:100mg/
(190- Azaperone:80-100mg/ Medetomidine:2- Thiafentanil:0.03mg/kg + Xylazine:0.1-
Carfentanil:2.9mg + Xylazine:28mg
300Kg) 3mg/Detomidine:2-3mg/ Xylazine:10-15mg 0.2mg/kg
Naltrexone:90-120mg(bull), 60-90mg
Naltrexone 50xEtorphine IV or
(cow) or Diprenorphine:15-20mg(bull), 10-
Diprenorphine 2.5x Etorphine or Naltrexone 30xThiafentanil IV or 100xCarfentanil IM or
Reversal 12mg(cow) (if Etorphine) + Atipamezole
15x Etorphine IV/ Atipamezole 5xMedetomidine or
5xMedetomidine or Detomidine +
0.1mg/kg (Xylazine) IM
Yohimbine: 1mL/50Kg
Table 22 – Different anesthetic protocols for the Roan antelope.
Thiafentanil:6-10mg/ Etorphine:6-8mg
Bull (190- +Azaperonel:60-100mg/ Medetomidine:5- Etorphine:4-5mg+ Azaperone:100mg/ Carfentanil:0.015-0.02mg/kg +
310Kg) 6mg/ Detomidine:3-5mg/ Xylazine:20- Carfentanil:5.6mg + Xylazine:56mg Xylazine:0.15-0.2mg/kg/
30mg Etorphine:0.025mg/kg + Xylazine:0.15-
Thiafentanil:6-8mg/ Etorphine:5-7mg 0.25mg/kg / Thiafentanil:0.01-0.02mg/kg +
Cow (190- +Azaperonel:50-80mg/ Medetomidine:3- Etorphine:3-4mg+ Azaperone:100mg/ Medetomidine:0.005-0.006mg/kg + +
300Kg) 5mg/ Detomidine:3-5mg/ Xylazine:20- Carfentanil:4.2mg + Xylazine:42mg Ketamine:0.3-0.6mg/kg
Naltrexone:75-150mg(bull), 90-120mg
(cow), 60-125mg (juvenile), 45mg (calf) or Naltrexone 50xEtorphine IV or
Diprenorphine:20mg(bull), 13-18mg(cow) Diprenorphine 2.5x Etorphine or Naltrexone 30xThiafentanil IV or 100xCarfentanil IM or
(if Etorphine) + Atipamezole 15x Etorphine IV/ Atipamezole 5xMedetomidine or
5xMedetomidine or Detomidine + 0.1mg/kg (Xylazine) IM
Table 23 – Different anesthetic protocols for the Springbok.
Thiafentanil:0.5-1mg/ Etorphine: 0.5mg/ Carfentanil:0.03mg/kg / Etorphine:0.05-
Fentanyl:10-15mg + Medetomidine:1mg/ No data 0.1mg/kg + Xylazina:0.15-0.25mg/kg /
Azaperone:10-20mg/ Xylazine:2-5mg Xylazine:0.5mg/kg + Ketamine:9mg/kg
Naltrexone 10-15mg or Diprenorphine:2-
Reversal No data 30xThiafentanil IV or 100xCarfentanil IM or
3mg (if Etorphine), 2.5mg (if Fentanyl)
IV/ Atipamezole 0.1mg/kg (Xylazine) IM
Table 24 – Different anesthetic protocols for the Bontebok.
Ram (55- Carfentanil:0.015-0.025mg/kg +
80Kg) Xylazine:0.2-0.35mg/kg + Ketamine:1.5-
Thiafentanil:3mg+ Azaperone:40mg or
2.5mg / Etorphine:0.02-0.025mg/kg +
Xylazine:5mg or Detomidine:3-5mg
No data Xylazine:0.2-0.3mg/kg + Ketamine: 0.2-
Ewe (55- /Etorphine:3mg+Azaperone:40mg
0.3mg/kg / Thiafentani:0.03mg/kg +
Azaperone:0.5mg/kg / Medetomidine:0.05-
0.09mg/kg + Ketamine:1-1.3mg/kg
Naltrexone 45-60mg or Diprenorphine 6- 30xThiafentanil IV or 100xCarfentanil IM or
Reversal No data
9mg IV/ Atipamezole 5xMedetomidine or
Table 25 – Different anesthetic protocols for the Giraffe.
Etorphine or Thiafentanil up to 18mg/
Thiafentanil up to 9mg + Etorphine up to
Bull (850- Etorphine:12-16mg / Thiafentanil:14-20mg 9mg /
1950Kg) * Thiafentanil:0.006-0.01mg/kg +
Etorphine:8-15mg / Thiafentanil:8-16mg / Medetomidine:0.01-0.014mg/kg +
Etorphine:4mg+Thiafentanil:8-16mg Ketamine:0.5mg/kg
Etorphine or Thiafentanil 14-15mg/
Cow (700- Etorphine:10-12mg / Thiafentanil:12-14mg Thiafentanil:7-7.5mg + Etorphine: 7-7.5mg/
1200Kg) * Thiafentanil:5-8.3mg + Medetomidine:8.3-
11.6mg+ Ketamine:415mg
Naltrexone 30-100x Etorphine or Naltrexone 0.3-0.4mg/Kg (if Thiafentanil and
Naltrexone10x opioid or Diprenorphine
Reversal Diprenorphine 2xEtorphine/ Naltrexone Etorphine), 0.2-0.3mg/Kg + Atipamezole
2.5x Etorphine
30xThiafentanil 0.05mg/Kg
Table 26 – Different anesthetic protocols for a standing sedation of African
elephants in controlled environments.
Azaperone: 0.06-0.15mg/kg / Butorphanol:
0.12mg/kg + Xylazine: 0.2-0.3mg/kg /
Bull (4000- Butorphanol:0.013-0.02mg/kg + Detomidine:
No data Xylazine: 0.11-0.55mg/kg / Detomidine: 0.014-
7000Kg) 0.013-0.02mg/kg
0.0162mg/kg + Butorphanol: 0.014-
0.0162mg/kg
Yohimbine: 0.5xXylazine Naltrexone 50-100x opioid/ 2-3.5mg/Kg +
Atipamezole:0.1xXylazine Atipamezole 0.1-0.16mg/Kg
Note: There is no published anesthetic protocol for a standing sedation on a free-ranging
African elephant.
Table 27 – Different anesthetic protocols for the White rhino.
Etorphine:2-3.5mg+Butorphanol:40- Carfentanil:1.2mg / Etorphine:2-3mg +
Cow Etorphine:2-4mg + Butorphanol: 30-80mg 90mg+Midazolam:25-50mg / Etorphine:3- Azaperona:20-40mg / Butorphanol:120-
(1800- + Azaperone:40-60mg or Detomidine:2- 4.5mg+Azaperone:40-60mg+Detomidine:10- 150mg + Medetomidine:5-7mg /
2200Kg) 4mg 20mg Butorphanol:70-120mg + Azaperone:100-
Naltrexone 40-100mg/mg Etorphine or 1-
Naltrexone:40-80mg or Diprenorphine: 6- Naltrexone 40x Etorphine IV or Diprenorphine
Reversal 2.5mg/mg Butorphanol (+ Atipamezole 5x
12mg 2-2.5xEtorphine IV
Medetomidine)
Table 28 – Different anesthetic protocols for the Golden wildebeest (Blue
wildebeest).
Thiafentanil:4-6mg or Etorphine: 5-7mg +
Bull (110- Etorphine:4-5mg +
Azaperone:100mg or Medetomidine: 5-6mg or Carfentanil:0.008mg/kg +
180Kg) Azaperone:100mg
Detomidine:10 mg or Xylazine: 20-40mg Xylazine:0.08mg/kg /
Thiafentanil:3-5mg or Etorphine: 3-5mg + Thiafentanil:0.03mg/kg +
Cow (110- Etorphine:3-4mg +
Azaperone:80mg or Medetomidine: 5-6mg or Xylazine:0.1mg/kg
Detomidine:10 mg or Xylazine: 5-10mg
Bull: Naltrexone 55-90mg or Diprenorphine:12-17mg;
Cowl: Naltrexone 45-75mg or Diprenorphine:10-13mg Diprenorphine 2.5x Etorphine or Naltrexone 30xThiafentanil IV or
+ Atipamezole 5x Medetomidine/Detomidine or Naltrexone 15x Etorphine 100xCarfentanil IM or IV
Table 29 – Different anesthetic protocols for the Plains zebra.
Stallion Etorphine: 4-7mg + Azaperone:40-60mg
(175- or Medetomidine: 5mg or Detomidine:5-10 Etorphine:0.0085-0.01mg/kg +
275Kg) mg or Xylazine: 40-60mg Acepromazine:0.035-0.04mg/kg / Detomidine: 0.1mg/kg +
Mare Etorphine: 4-6mg + Azaperone:40-60mg Tiletamine+Zolazepam:5.5-8.3mg/kg + Butorphanol:0.13mg/kg
(175- or Medetomidine: 5mg or Detomidine:5-10 Detomidine:0.06-0.08mg/kg
275Kg) mg or Xylazine: 40-60mg
Naltrexone 60-100mg or Diprenorphine Naltrexone: 0.1mg/kg + Atipamezole
Reversal Diprenorphine 0.045mg/kg IV (for Etorphine)
10-18mg (Stallion) or 10-15mg (Mare) 2xDetomidine IV
Table 30 – Different anesthetic protocols for the Bushbuck.
Xylazine:5mg or Detomidine:3-5mg No data No data
/Etorphine:3mg+Azaperone:40mg
Naltrexone 45-60mg or Diprenorphine 6-
Reversal No data No data
Table 31 – Different anesthetic protocols for the African lion.
TZ:0.5mg/kg + Medetomidine: 0.03mg/kg /
TZ:3-5mg/kg / Ketamine:5mg/kg+
Xylazine:0.5mg/kg / Ketamine:5mg/kg +
Lion (120-
Medetomidine:0.05mg/kg / TZ:1.6-4.2mg/kg / Ketamine:3-10mg/kg+
Butorphanol:0.2-0.3mg/kg + Xylazine:0.3-1mg/kg / Ketamine:2-6mg/kg +
TZ: 0.6-1mg/kg + Medetomidine:0.015-
Medetomidine:0.05mg/kg + Medetomidine:0.03-0.07mg/kg /
0.025mg/kg / TZ: 4-6mg/kg/ Ketamine:7-
Midazolam:0.15mg/kg Butorphanol:0.003-0.04mg/kg +
10mg/kg+ Xylazine:1-4mg/kg /
TZ:0.5mg/kg + Medetomidine: 0.03- Medetomidine:0.1-0.4mg/kg + Midazolam:0.1-
Ketamine:1.9-5.7mg/kg +
0.05mg/kg / TZ l:3-5mg/kg / 0.3mg/kg / Butorphanol:0.003-0.04mg/kg +
Medetomidine:0.02-0.08mg/kga
Lioness Ketamine:5mg/kg+ Xylazine:0.5mg/kg / Medetomidine:0.1-0.4mg/kg + Midazolam:0.1-
(120- Ketamine:5-7mg/kg + Medetomidine:0.03- 0.3mg/kg + Ketamine:1-2mg/kg
250kg) 0.05mg/kg / Butorphanol:0.2-0.3mg/kg +
Medetomidine:0.05mg/kg +
Midazolam:0.15mg/kg
Atipamezole 5xMedetomidine /
Atipamezole0.1-0.29mg/kg (if a) / Yohimbine Atipamezole / Naltrexone / Yohimbine/
Reversal Yohimbine:1mL/50Kg IV (if Xylazine)
0.1mg/Kg IV if Xylazine Flumazenil if need it (doses without data)
/Naltrexone:2xButorphanol
Table 32 – Different anesthetic protocols for the Cheetah.
Male (35- TZ:1.6-4.2mg/kg / Ketamine:3-10mg/kg+
TZ:0.5mg/kg + Medetomidine: 0.03-
72Kg) Xylazine:0.3-1mg/kg / Ketamine:2-6mg/kg +
0.05mg/kg / TZ:3-5mg/kg / Ketamine:3- TZ: 1.6-7.8mg/kg /
Medetomidine:0.03-0.07mg/kg /
5mg/kg+ Xylazine:0.5mg/kg / Ketamine:3- Medetomidine:0.035mg/kg+
Butorphanol:0.003-0.04mg/kg +
5mg/kg + Medetomidine:0.03mg/kg / Butorphanol:0.02mg/kg +
Medetomidine:0.1-0.4mg/kg +
Butorphanol:0.2mg/kg + Midazolam:0.15mg/kga / Saffan®:5mL IV
Female Midazolam:0.1-0.3mg/kg /
Medetomidine:0.15mg/kg + (alphaxolone 9mg/mL + alphadolone:3mg/mL)
(35-72Kg) Butorphanol:0.003-0.04mg/kg +
Midazolam:0.03mg/kg / Saffan®:60-120mg / Ketamine:1.57-2.5mg/kg + Medetomidine:
(alphaxolone 9mg/mL + 0.031-0.07mg/kgb
Midazolam:0.1-0.3mg/kg + Ketamine:1-
alphadolone:3mg/mL)
2mg/kg
Atipamezole 5xMedetomidine / Atipamezole:0.18mg/Kg +
Atipamezole / Naltrexone / Yohimbine/
Reversal Yohimbine:1mL/50Kg IV (if Xylazine) / Flumazenil:0.006mg/Kg + Naltrexone:
Flumazenil if needed (doses without data)
Naltrexone 2xButorphanol 0.25mg/Kg / bAtipamezole: 0.15-0.3mg/kg
Table 33 – Different anesthetic protocols for the Leopard.
TZ:1.6-4.2mg/kg / Ketamine:3-10mg/kg+
Xylazine:0.3-1mg/kg / Ketamine:2-6mg/kg +
TZ:1-3mg/kg + Medetomidine: 0.05- Butorphanol:0.003-0.04mg/kg +
Male (23- 0.08mg/kg / TZ:5-10mg/kg / Ketamine:8- Medetomidine:0.1-0.4mg/kg +
Ketamine:5-10mg/kg + Xylazine:1-4mg/kg
91Kg) 10mg/kg+ Xylazine:1mg/kg / Ketamine:5- Midazolam:0.1-0.3mg/kg /
10mg/kg + Medetomidine:0.05-0.08mg/kg Butorphanol:0.003-0.04mg/kg +
Atipamezole 5xMedetomidine / Atipamezole / Naltrexone / Yohimbine/
Yohimbine:1mL/50Kg IV Flumazenil if needed (doses without data)
Table 34 – Different anesthetic protocols for the African wild dog.
Male (19- TZ:0.5mg/kg + Medetomidine: 0.03- Ketamine:3-5mg/kg + Medetomidine:0.005-
35Kg) 0.05mg/kg or Xylazine 0.5mg/kg / TZ:3- 0.1mg/kg a/ Medetomidine:0.045mg/kg +
Medetomidine: 0.04-0.06mg/kg +
5mg/kg / Ketamine:5mg/kg+ Butorphanol:0.24mg/kg + Midazolam:0.3mg/kg
Butorphanol: 0.18-0.3mg/kg + Midazolam:
Medetomidine:0.1mg/kg / Fentanyl:2.5mg / Xylazine: 0.7-1.1mg/kg + Fentanyl: 0.1mg/kg
Female 0.18-0.4mg/kg / Ketamine:1.5-5mg/kg +
+ Xylazine:0,5mg/kg / + bolus o 10mg of Xylazine and 0.5mg of
(19-35Kg) Medetomidine:0.04-0.1mg/kg / TZ:1-4 mg/kg
Butorphanol:0.15mg/kg + Medetomidine: Fentanyl / Ketamine: 1.6mg/kg + Xylazine:
0.05mg/kg 2.2mg/kg / TZ:1-4mg/kg
Atipamezole:0.1mg/Kg if a, 3mg/Kg if b (plus
Atipamezole 5xMedetomidine; Naltrexone Naltrexone:10mg + Flumazenil:0.2mg);
Reversal 2xButorphanol; Yohimbine (no dose Yohimbine:0.125mg/Kg + No data
available) if Xylazine Naloxone:0.04mg/Kg if c; Yohimbine:0.2mg/Kg
if d
Injection (Medicine)
Documents Similar To A Short Review of the Chemical Immobilization Principles in Some Common African Wildlife Species
saais
Cedy L. Cielo
taichi7
JP Fernandez
BOOMTIME
Gina Castillo
bwester2222
edward_terisno
Harish Chand
Salman Shalwani
Ljubica Prusac
Arup Nag
Healthcare Representative
JANGAN LUPA DISAMBUNG MARWAH !!!!!!!!!!
Marwa Hayon
competition invite 1
Amit Project
Rakesh Meel
National Safety Month: Keeping aging loved ones safe at home
50yearsafter
sunmars
OSHA3491QuickCardPictogram.pdf
dimasfebrianto
Health Policy . Reading, Discussion and Questions Assignments Due on February 10 and 24. 2013 (1)
Immanuel Ronald Lewis
use a helmet
Nature's Pathways Aug 2011 Issue - Northeast WI Edition
Nature's Pathways
More From Andrés González Rodríguez
Aliment a Cio Naves
Andrés González Rodríguez
NOM-056-ZOO-1995 Pruebas Laboratorios Aprobados 22 de Febrero de 1999 DOF
12.Terapeutica rapaces
Guía de Los Derechos y Obligaciones de Los Inspeccionados
Theeffectsofcaptiveexperienceonreintroduction (1)
Kevin Puertas
Enfermedades de Las Aves
Ester Bonnie Rojas Silva
inter oto
Mirleys Merlano
Drugs and Alcohol Modified
MonicaDzhaleva
drug_addiction_SI_v16_prefaceinToC-final.pdf
NachoConde
Metabolic Report 2017
Alice Nguyen
27_HEALING_SECRETS_0309.pdf
2012 -j Popul Ther Clin Pharmacol- Neonatal Abstinence Syndrome
Pablo Vercelli
PTSD and Veterans
Tom Slotcavage
MINI-CASE-3-CKD
YetTamparong
Dinitrophenol Notes
antonio carlos
GHS SDS E Sikafloor Curehard -24 2015
huo lenghak
Klipper Montaño Ontaneda
ch 21.ppt
Mylyn Mas
ayusuryani
HPV Vaccine Support Group Concerned at Side-effects but Irish Government Ignores the Parents
Rita Cahill
Examination of Orthopedic and Athletic Injuries Starkey PDF
xi-xu
chain of infection
Fatin Muhammed
Chap 11. Promoting Regular PA
David Montoya Dávila
kniga-nemechek.pdf
Complement fixation test
GIST (Gujarat Institute of Science & Technology)
MN_2018-01-23
moorabool
FlyingWalrus
Sensory Motor Integration
AdelinaPanaet
Cardiac Re Synchronization Therapy
sockembopem
Electronic Torture - RADAR Neurological Reprogramming Experiments
stop-organized-crime
oral pathology piks
zeedental
Barriers to ICTC and ART Linkages Among Men Who Have Sex With Men in Karnataka Single Pages
Priya Pillai
Aŋdrés Melo
Electromyographic Analysis of Core Trunk, Hip, And Thigh Muscles During 9 Rehabilitation Exercises
jopepinho5796
Vademecum Nuclear Medicine - Leslie - Landes Biosc
Slobodan Sreckovic
Catatan Obat 1
Andrew Soerijadi