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In: Zoological Restraint and Anesthesia, D. Heard (Ed.)
Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA.
Capture and Immobilization of Free-living Jaguars (Panthera onca) (2-Dec-2004)
S. L. Deem
Department of Animal Health, Smithsonian National Zoological Park, Washington DC, USA.
Introduction
Jaguars (Panthera onca) (Fig. 1) are the largest felid species in the New World and the only member of the genus
Panthera, the roaring cats, that occurs in the Americas. They are the third largest cat species, being outsized only by
lions (P. leo) and tigers (P. trigris). Although not the largest felid, jaguars have the strongest jaw in relation to head size
of any of the cats, a fact that should be remembered whenever planning to capture and immobilize these animals. The
body weight of jaguars is 90 - 120 kg for males and 60 - 90 kg for females, with a large variation in body size. Jaguars
live in a wide variety of tropical habitats, ranging from montane forest and wet savannah to tropical rain forest and
deciduous tropical forest. The largest documented jaguars occur in wet savannahs while jaguars that live in more forested
regions tend to be smaller in size [1].
Figure 1. Male jaguar (Panthera onca). (Photo by Deem SL). - To view this image in full size go to
the IVIS website at www.ivis.org . -
Historically the range of jaguars was the southern USA through Central and South America as far south as southern
Argentina. Their current range is limited to a broad belt from central Mexico through Central America to Northern
Argentina [2,3] (Fig. 2). The jaguar is known to be rare or extinct in many parts of its former range and it is
approximated that 10,000 are left with several subspecies being rare.
Figure 2. Distribution of jaguar (Panthera onca). (With permission from: Sanderson EW, Redford
KH, Chetkiewicz C-LB, et al. Planning to save a species: the jaguar as a model. Con Biol 2002;
16:58-72.). - To view this image in full size go to the IVIS website at www.ivis.org . -
Jaguars have been cultural icons for many indigenous American people, including the Mayans and Incas, and have
fascinated humankind for millennia. In the more recent past, this fascination has been evident in the many scientific
studies that have been conducted, primarily focusing on the natural history of jaguars [4-7].
Studies on the home range of jaguars have found large variation, with animals in the Pantanal of Brazil having ranges
twice as large as those in a forested area of Belize. In the Pantanal, study males had a range of 50 - 76 km2 and females
25 - 38 km2 [7]. In the Belize study, males had a range of 28 - 40 km2 and females approximately 10 km2 [6].
Studies on the food habits of jaguars have been extensive with findings showing the variability related to the prey
available and habitat type [5,6,8-10]. Large prey items include tapir, peccary, deer, and in some cases livestock.
However, jaguars are opportunistic feeders and will consume capybara, sloths, armadillos, fish, reptiles, birds, insects,
and, in coastal regions, sea turtles.
The biggest conservation threats for jaguars are due to habitat fragmentation and hunting of "problem cats" or cats for the
commercial sell of their pelts [3]. Although the specific health threats to free-ranging jaguars are largely unknown at this
time, they are probably similar to those cited for the health concerns of wildlife in general and include anthropogenetic
influences, often associated with increased contact that wildlife have with livestock, domestic carnivores and humans, as
well as habitat fragmentation and contamination of their habitats [11]. Additionally, jaguar-human conflicts which
include hunting of jaguar directly (often due to real or perceived high levels of predation on livestock), as well as hunting
of their prey, is believed to be the number one "health" threat.
Many infectious and non-infectious diseases have been documented in captive jaguars. Non-infectious problems include
a high incidence of neoplasia [12-15] which may be associated with husbandry in captivity and/or longevity. Many
infectious agents have been documented to cause morbidity and/or mortality including protozoan [16], bacterial [17] and
viral pathogens (i.e., canine distemper, feline infectious peritonitis) [18,19]. Additionally, there is serologic evidence of
infection with canine distemper and feline immunodeficiency virus, [18,20,21]. It is also assumed that jaguars are
susceptible to the common respiratory disease agents (i.e., Chlamydia sp., herpesvirus-1, and calicivirus) in domestic and
non-domestic cats. Few studies have been conducted on the health status of jaguars in the wild with the majority of data
on parasite infection and infestation [1,22,23]. However, it can be expected that the loss of habitat, and associated
increased contact of jaguars with humans and their domestic animals, may lead to an increase in the incidence of
infectious and parasitic diseases in free-ranging jaguars.
Reference ranges for physiological data values are available for captive jaguars. The hematologic values of 18 jaguars
held at the Zoological Park in London are presented in one paper [24]. The International Species Information System
(ISIS) provides tables with hematology and blood chemistry values based on sex and age [25]. The combined values for
both sexes and all ages is presented here in Table 1 [25].
Physiological reference ranges for captive jaguars (Panthera onca) submitted to the International
Species Information System from 39 member institutions [25].
Parameter Mean Standard Deviation Sample size * Animals **
WBC (103/ul) 12.01 4.099 191 98
RBC (106/ul) 7.26 1.36 161 84
Hemoglobin (g/dl) 11.8 2.3 166 86
Hematocrit (%) 34.8 5.7 199 102
MCV (fl) 48.8 9.3 159 82
MCH (pg/cell) 16.6 3.9 154 79
MCHC (g/dl) 33.7 3.3 165 86
Platelet Count (103/ul) 273 107 37 30
NRBC/100 WBC 1 1 11 11
Reticulocytes (%) 0.0 0.0 5 5
Segmented Neutrophils (103/ul) 8.56 3.92 179 90
Lymphocytes (103/ul) 2.15 2.09 182 94
Monocytes (103/ul) 0.35 0.39 142 84
Eosinophils (103/ul) 0.297 0.307 135 77
Basophils (103/ul) 0.051 0.1 41 26
Neutrophilic Bands (103/ul) 0.813 1.657 77 46
Calcium (mg/dl) 9.8 0.8 148 80
Phosphorus (mg/dl) 5.0 1.1 131 74
Sodium (mEq/L) 151 4132 75
Potassium (mEq/L) 4.0 0.4 132 75
Chloride (mEq/L) 121 5123 70
Bicarbonate (mEq/L) 170.0 316.7 4 3
CO2 (mEq/L) 16.0 2.9 60 36
Osmolarity (mOsmol/L) 303 730 19
Many field researchers are currently immobilizing free-ranging jaguars for studies to answer the many questions that
remain unknown about their natural history, biology, genetics and health status. Unfortunately, many of these researchers
Physiological reference ranges for captive jaguars (Panthera onca) submitted to the International
Species Information System from 39 member institutions [25].
Parameter Mean Standard Deviation Sample size * Animals **
Iron (ug/dl) 84 20 15 7
Magnesium (mg/dl) 2.66 0.54 5 5
BUN (mg/dl) 24 9155 88
Creatinine (mg/dl) 2.0 0.7 152 84
Uric Acid (mg/dl) 0.3 0.3 59 34
Total Bilirubin (mg/dl) 0.2 0.1 133 78
Direct Bilirubin (mg/dl) 0.0 0.1 45 26
Indirect Bilirubin (mg/dl) 0.1 0.1 45 26
Glucose (mg/dl) 137 55 154 86
Cholesterol (mg/dl) 246 60 140 78
Triglyceride (mg/dl) 32 19 75 40
CPK (IU/L)*** 317 279 62 44
LDH (IU/L)*** 163 162 90 55
ALP (IU/L)*** 33 33 147 80
AAT (IU/L)*** 55 25 119 69
AST (IU/L)*** 35 16 150 84
GGT (IU/L)*** 3 3 56 33
Amylase (U/L) 1816 901 40 25
Lipase (U/L) 14 12 12 8
Total Protein (g/dl) 7.3 0.6 142 78
Globulin (g/dl) 3.9 0.8 113 62
Albumin (g/dl) 3.4 0.4 114 62
Total Triiodothyronine (ng/ml) 154.0 0.0 1 1
Total Thyroxine (ug/dl) 2.5 2.0 8 7
Body Temperature (ºF) 100.9 2.0 107 67
Weight: 1.8 - 2.2 years old (Kg) 53.49 13.85 9 9
Weight: 9.5 - 10.5 years old (Kg) 65.76 11.07 24 17
Weight: 19.0 - 21.0 years old (Kg) 56.39 13.38 12 7
Sample size* - Number of tests run per parameter.
Animals** - Number of animals sampled per parameter.
*** CPK - Creatinine phosphokinase; LDH - Lactate dehydrogenase; ALP - Alkaline Phosphatase; AAT - Alanine aminotransferase;
AST - As
p
artate aminotransferase; GGT - Gamma
g
lutam
y
ltransferase.
have little or no training in veterinary skills, such as anesthesia and troubleshooting during an anesthetic emergency.
Although veterinarians should always be available and part of the jaguar handling team in the field, this often is not the
case. This chapter provides a brief review of the literature on the capture and anesthesia of free-ranging jaguars,
recommendations for the safe immobilization of free-ranging jaguars, and gives information on troubleshooting for
anesthetic emergencies in the field. This chapter has been written for veterinarians and field biologists with previous
training in basic anesthesia principles.
Immobilization Procedure
General Principles -
Any person who immobilizes a wild jaguar must remember that she/he is solely responsible for the health of that animal
from the time the drug is administered (or from the time the animal is captured or treed) until the animal has fully
recovered from the anesthetic agent(s). It is imperative that anyone engaged in the immobilization of free-ranging jaguars
know how to handle the anesthetized cat, monitor physiologic parameters, and respond to medical emergencies should
they arise. Although many anesthetic agents are relatively safe in felid species, anesthetic emergencies can and do occur
even under the best of circumstances.
Unlike the hospital setting where anesthesia is more controlled, there are unique problems related to immobilization of
free-ranging wildlife in general, and large cats in particular. The capture method may itself result in injuries. Jaguars are
aggressive cats and often when trapped will bite on cage material. Free-ranging jaguars have succumbed to tooth root
abscesses following fracture of canine teeth from the capture procedure [1]. If chased by dogs and darted while in a tree,
the fall itself may cause injury. For this reason, it is best to not dart a jaguar above 5 meter high in a tree to avoid
traumatic falls. Jaguars are often highly stressed during capture. The capture team must strive to minimize stress due to
the effects that stress may have on physiologic parameters that may compromise the animal once anesthetized. Lastly, it
must always be remembered that capture in the wild of a potentially dangerous animal, such as a jaguar, has inherent
risks for the capture team.
Capture Methods -
There are a variety of capture and immobilization methods for free-ranging felids [26]. Wilson et al. [27] provide an
excellent overview of capture methods for medium to large sized mammals. Methods that have been used to capture free-
ranging jaguars include treeing the animal using dogs, padded foot-hold traps, snares (i.e., Aldrich snares) and cage or
box traps [6,7,28]. The later two methods may or may not include bait (i.e., live goat or pig) to lure the animal to the trap.
Once the jaguar is treed or trapped, it can then be darted. The capture method employed for each jaguar capture should be
based on the immobilization team’s previous experience, methods that have been successful in the region (if studies
exist), habitat, and current weather conditions. In every capture and immobilization procedure the top priority is for a
safe anesthetic event for both the jaguar and the people involved with the procedure.
Pre-anesthetic Management -
Once a jaguar has been captured, it is important to perform the anesthesia as quickly as possible. When in a cage, the
possibility of damaging canine teeth is high and may increase with prolonged time in the cage. A technique to minimize
stress includes tranquilizer tablets [29,30] which are commonly used with padded foot-hold traps, but may be of value
with the other capture methods. As is true for field immobilization in general, you should not take a lot of time once you
begin your initial approach to dart the captured animal. An approximation of the body weight for the calculation of drug
must be done to minimize a drug dose error. It is imperative that you have all your immobilization equipment ready prior
to approaching the cat as reviewed in Osofsky and Hirsch [31].
Anesthetic Administration -
Anesthetic agents should only be administered to free-ranging wild jaguars using remote drug delivery systems (RDDS).
There are a variety of RDDS available for the practitioner, nicely reviewed by Bush [32] and Nielsen [33]. A blowpipe,
or possibly a pole syringe, may be used for immobilizing jaguars in a cage, foot-hold trap or snare. In all other field
situations, it is best to use a rifle or pistol (i.e., Telinject tm, Cap-Chur tm, Dan-Inject tm). It is beyond the scope of this
chapter to cover the principles of all these products. The practitioner must be familiar with the instrument he/she chooses
for use in the field. Darting animals is always associated with some degree of risks. Serious damage to the animal (and
human participants) can and does occur if inappropriate instruments are used and/or if instruments are used
inappropriately.
Dart and needle selection is also important in preparing for a jaguar immobilization. Darts that are too heavy and needles
that are too long/thick can cause serious damage on impact. Damage is also possible if the charge of the dart or the
charge of the rifle/pistol is too high. Needles available include collared, plain and barbed. A collared needle is often
employed during immobilization procedures because it remains in the animal and ensures total drug injection.
Unfortunately, if the jaguar is not adequately immobilized and cannot be restrained, the dart will remain in the animal
and may cause problems.
The author recommends the use of 1.5 x 30 mm (18 gauge x 1 - 1/4 inch) collared needles for immobilizing free-ranging
adult jaguars. However, if the jaguar is treed or trapped prior to darting, a non-barbed (plain) needle can be used. Non-
barbed (plain) needles cause less trauma to the tissues but often do not remain in the animal as long as collared needles
and thus may not inject all the drug prior to falling from the animal. Again, the practitioner must be comfortable and
familiar with the equipment he/she takes to the field. When darting a jaguar, it is safest to aim for the proximal region of
a rear limb (Fig. 3). Some researchers recommend darting the triceps region of the front arm. If the anesthesiologist
elects to use the front limb, it must be remembered that the thoracic region and head are in very close proximity to the
intended site.
Figure 3. Dart placement sites for darting free-ranging jaguars (Panthera onca). (From: Deem and
Karesh [47]). - To view this image in full size go to the IVIS website at www.ivis.org . -
Serious harm can be inflicted on the cat if the dart hits one of these regions. It is for this reason that the author
recommends the hind leg unless the anesthesiologist is darting the jaguar at close range (i.e., in a box trap) (a shorter
needle may be more appropriate when using the tricep region.) When aiming for the rear limb, darts should be placed in
the caudal most aspect of the muscle mass to avoid the femoral bone and the sciatic nerve. Needles and darts must be
disinfected prior to use on the next animal to prevent the spread of diseases. Although disinfection is often the only
available means of equipment care in the field, it is best to sterilize needles between animals.
Anesthesia
Literature review - The anesthetic protocols that have been used on free-ranging jaguars and are published in the
literature are listed in Table 2. The author has included the sample size for those studies that published it. If you choose
to use one of these protocols, you should refer to the cited paper for detailed information.
Table 2. Literature review of anesthesia dosages reported in the literature for the chemical
immobilization of free-ranging jaguars (Panthera onca)
Drug Dosage (mg/kg) Sample Size Reference
Ketamine 10 - 12 n/a* [1]
Ketamine 7 - 40 9[34]
Ketamine 22 7[6]
Ketamine
Diazepam
11.8
0.25 2[1]
Ketamine
Xylazine
Atropine
3
0.6
0.05
1[1]
Ketamine
Xylazine
Diazepam
7
0.5
10
1[1]
Ketamine
Xylazine
10.6 - 11.5
1.3 - 1.4 2[35]
Ketamine
Xylazine
11
18[36]
Ketamine
Xylazine
6.6
0.66 n/a [37]
Ketamine
Medetomidine
Atipamezole
1.46 - 3.48
0.36 - 0.087
0.122 - 0.163
2[38]
Telazol 6.6 - 16.4 11 [39]
Telazol 10 n/a [40]
Telazol 3.9 11 [1]
Telazol 3.5 - 9.1 6[34]
* n/a - Not available
There are no published reports on the use of inhalant anesthesia used to immobilize free-ranging jaguars. Although
portable anesthetic machines are available and have been employed during the maintenance of anesthesia for various
free-ranging species, they are not practical and seldom available for field work. If inhalant anesthesia is an option for
your particular project, you must first be familiar with inhalant anesthetics and available equipment for use in cat species
[41].
Recommended Protocol - Currently, the author recommends the following anesthetic regimen for use by field personnel
with little experience in immobilizing free-ranging jaguars. This regimen should provide an adequate plane of anesthesia
for short-term work on the jaguar (i.e., radio-collar application, morphometric measurements, biomaterial collections)
while requiring a minimal level of technical skill in anesthesiology.
Telazol (6 - 10 mg/kg ) IM as the dose for immobilization in free-ranging jaguars. The darter has the option to include
150 mg ketamine in the initial dart based on work by Hoogesteyn and Cavalcanti (unpublished data). Supplemental
ketamine at a dose of 1 - 1.5 mg/kg, IV or 1 - 2 mg/kg IM, as needed to maintain an adequate level of anesthesia. (No
supplemental ketamine should be delivered for at least 10 minutes after the initial dart containing telazol.) Atropine at a
single dose of 0.04 mg/kg either SC or IM may also be administered if the cat has excessive salivation.
There have been anesthesia related problems with telazol use in large cat species, in particular tigers [42]. The author
knows of no similar problems with the use of telazol in jaguars, but again, the practitioner should be prepared to deal
with unexpected reactions.
One possible exception to this recommendation is the use of ketamine (10 - 20 mg/kg) alone when immobilizing any
jaguar that has first been treed using dogs. This exception is based on anecdotal information that some jaguars have
fallen from trees when immobilized with telazol or a ketamine / xylazine combination (H. Quigley, personal
communication).
Antagonists
zFlumazenil is the antagonist for zolazepam (the benzodiazepine component of telazol) and can be administered,
once all procedures are completed, at an IM dose of 1.0 mg of flumazenil for each 40 mg of telazol used.
Flumazenil should not be administered for a minimum of 30 minutes after the initial dose of telazol was delivered
to ensure the tiletamine component of telazol is nearly completely metabolized. There should also be at least 30
minutes between the administion of any supplemental ketamine administration and flumazenil.
zYohimbine is the antagonist for xylazine and should be administered at 0.125 mg/kg IM and should only be
delivered once the procedure is completed and at least 30 minutes after the last dose of the cyclohexane
(ketamine) was given.
zAtipamezole is the antagonist for medetomidine and can be administered once all procedures are completed, at a
dose of 4 - 5 x the medetomidine dose. For example, if 40 ug/kg of medetomidine was used for immobilization,
reversal with atipamezole should be at a dose of 160 - 200 u/kg. This should be delivered IM. Atipamezole
should not be administered for a minimum of 30 minutes after the last dose of cyclohexane (ketamine) was given.
Supplemental Drugs
There will be occasions when the initial anesthetic agent(s) does not provide adequate immobilization or when the effect
of the anesthetic agent(s) begins to wane (i.e., increased animal movements, increased respiration and heart rate) prior to
all procedures (i.e., radiocollar application, sample collection) being completed. In these cases, it may be necessary to
administer supplemental drugs for adequate anesthesia to allow safe handling. The following should be kept in mind if
one is faced with either of these situations.
zKetamine at a dose of 1 - 1.5 mg/kg IV or 1 - 2 mg/kg IM, as needed to maintain an adequate level of anesthesia
should be a safe dose in MOST adult jaguars.
zDiazepam (valium) at the dose of 5 - 10 mg/jaguar should be administered slowly IV to any jaguar with extreme
muscle rigidity, muscle tremors, and/or seizures. Diazepam can be administered again IV after 3 minutes if there
is no response to the initial injection. If the jaguar still does not respond following the second injection, another
cause of the seizure activity should be considered. If a vein cannot be located (i.e., moving animal), diazepam can
be injected IM. Caution should be exercised in administering a second dose of diazepam following an IM
injection due to a potentially slower rate of metabolism with IM injections.
zNEVER use telazol as the supplemental drug. If telazol is the initial immobilizing agent and it has not provided
adequate anesthesia or if its anesthetic effects have worn off, it is best to supplement with ketamine either IV or
IM. The dose of ketamine will depend on the plane of anesthesia prior to supplementation. 25 - 50 mg IV or 50 -
100 IM mg of ketamine total per jaguar should be a safe dose in MOST adult jaguars.
zNEVER use xylazine, medetomidine and midazolam as the supplemental drug. They should only be administered
in combination with another drug (i.e., ketamine) for induction of anesthesia. It is best to supplement with
ketamine either IV or IM. The dose of ketamine to deliver will depend on the plane of anesthesia prior to
supplementation. 25 - 50 mg IV or 50 - 100 mg IM of ketamine should be a safe dose in MOST adult jaguars.
If you are not sure of how much of the original drug(s) was successfully administered (i.e., poor dart placement, dart
bounced in and out quickly), you should wait at least 15 minutes following the initial dart prior to administering any
additional agents.
Anticholinergics
Some authors recommend the addition of atropine to the anesthetic protocol for the anticholinergic property of
decreasing salivary secretions. However, atropine can be associated with negative side effects, most commonly on the
heart and gastrointestinal tract. In field situations it may be more appropriate to administer atropine only to those cats that
are displaying excessive salivation during the immobilization procedure. A single dose should be administered:
Atropine - 0.04 mg/kg SC or IM.
Animal Handling and Monitoring
Standard equipment for handling and monitoring the anesthetized jaguar should include those listed in Table 3. The
author has listed just the bare minimal essentials that should be available whenever a jaguar is immobilized in the field.
All handling equipment (i.e., towels, non-disposable gloves, veterinary supplies) should be disinfected prior to use on
another animal to prevent the spread of diseases.
Immediately after the animal is darted and an initial assessment of the respiratory rate (RR; 8 - 24 breathes/minute) and
heart rate (HR; 70 - 140 beats/minute) are deemed within normal limits, the dart should be collected (avoid handling the
needle) and put in a safe place. It is best to have one person immediately take the physiologic parameters while a second
person is in charge of the dart. The dart site on the animal should not be touched to avoid contact with drug residues and
blood. People who will have contact with the immobilized animal should wear latex gloves during the immobilization
procedure to avoid the transmission of infectious diseases between the animal and him/herself, as well as to minimize
contact with drug residues at the injection site. The animal should be placed in a position that allows it to breathe easily
(Fig. 4).
Figure 4. Jaguar (Panthera onca) in lateral recumbency during anesthesia. (Photo by Deem SL). - To
view this image in full size go to the IVIS website at www.ivis.org . -
Preferably, the jaguar should be placed in lateral recumbency. The head and neck should be placed in a position that
allows air to flow through the mouth and trachea. The mouth should be kept lower than the back of the throat and neck so
saliva flows out of the mouth and not into the trachea.
Once the animal is anesthesized and placed in the proper position, the eyes must be protected. A triple antibiotic eye
ointment (i.e., Trioptic-P tm) should be applied in both eyes to prevent them from drying due to the lack of the normal
blink response which is often the case when using ketamine and telazol anesthetics. A towel (nonabrasive material
preferably) should then be placed over the eyes to protect them from the sun and dirt, as well as to minimize stressful
stimulus to the animal. Cotton balls may be placed in the outer ear canal to minimize auditory stimulus. However, should
one choose to use these, one must remember to remove them at the completion of the immobilization procedure. It is
Table 3. Standard equipment for handling and monitoring the anesthetized
jaguar
Monitoring Equipment
- Stethoscope
- Thermometer
- Pulse oximeter
Emergency Equipment
- Laryngoscope
- Endotracheal tubes
- Ambu bag or Oxygen tank
- Anesthetic reversal agents (see above)
- Emergency drugs (see below)
- Portable ice packs
- Dental repair kit
- Surgical pack
- Bandage material
important to reduce the risk of wound infection by screwworm (Cochliomyia hominivorax). Topical betadine and a fly-
strike ointment can be applied to the dart site, and to any abrasions that occur during the procedure, to protect against
screwworm.
During all jaguar immobilizations, the physiological parameters (i.e., respiratory rate, heart rate, and temperature) MUST
be monitored. If these values fall outside the normal range, the immobilization team should be alerted to a potential
impending emergency and be ready to respond in the appropriate manner. The normal physiologic parameters for an
immobilized free-ranging jaguar are the following:
Temperature (T) 37 - 39.5ºC (98.6 - 103.1ºF)
Respiratory Rate (RR) 8 - 24 breathes/minute
Heart Rate (HR) 70 - 140 beats/minute
Both respiratory rate and heart rate should be monitored every 5 minutes and the temperature should be taken
every 10 minutes.
Monitoring these parameters can best be done by use of a thermometer, visual observation of chest wall expansion, and
either palpation of the femoral pulse or use of a stethoscope. A rectal thermometer should be placed in the anus (digital
thermometers are the best and easiest to use in the field) and the temperature monitored at 10 minute intervals during
anesthesia. Respiration can be monitored by watching the thorax move when the animal breathes. The easiest way to
determine the respiratory rate per minute, is to count the thoracic movements during 15 seconds and then multiply this
number by four. If one does not have a stethoscope in the field, then light digital pressure over the femoral artery will
provide a measure of the heart rate. Alternatively, a stethoscope can be used to auscultate the heart directly over the
lateral aspect of the cranial thorax.
The recognition of what are normal jaguar responses to anesthetic agents is also imperative. Jaguars immobilized with
telazol and ketamine usually will have increased salivation, open eyelids, whole body muscle rigidity (including jaw
tone), and intact reflexes (i.e., corneal, pedal). Jaguars should maintain swallowing and coughing reflexes with these
agents, but should not have muscle tremors and seizure-like activity.
In addition to the drugs necessary for safe and effective anesthesia (which includes a triple antibiotic eye ointment), a
number of supportive medications are valuable for field work. Ivermectin (200 mcg/kg SC) should be administered to
prevent screwworm infestation. Fluid therapy with Lactated Ringer’s solution (10 - 20 ml/kg IV or SC) for rehydration
should be provided especially if the jaguar was trapped for an extended period and/or was highly stressed and
hyperthermic. A long-acting antibiotic such as penicillin G benzathine (40,000 IU/kg IM) should be administered,
especially for jaguars that have sustained significant trauma from the dart or a fractured tooth, had vomited during the
procedure, or had active lesions at the time of immobilization. Both topical fly-strike and triple antibiotic ointments
should be placed on the dart site as well as any active skin lesions.
Troubleshooting Common Anesthetic Emergencies in the Field
Any person who immobilizes a wild jaguar must remember that he/she is solely responsible for the health of that animal
from the time the drug is administered (or from the time the animal is captured or treed) until the animal has fully
recovered from the anesthetic agent(s). It is imperative that anyone engaged in the immobilization of free-ranging jaguars
know how to handle the anesthetized cat, monitor physiologic parameters, and respond to medical emergencies should
they arise. Although many anesthetic agents are relatively safe in felid species, anesthetic emergencies can and DO occur
even under the best circumstances.
The author will not present a comprehensive review for all aspects of anesthesia-related veterinary emergencies.
However, the author will provide highlights on the most common emergencies that may arise during the immobilization
of free-ranging jaguars. The author strongly recommends that researchers performing jaguar immobilization do further
reading on this subject [33,43-46].
The most common anesthesia emergencies in free-ranging jaguars are respiratory depression and arrest, cardiac arrest,
seizures, hyperthermia, and wounds including canine tooth fractures. Additional problems include vomiting and
aspiration, shock, capture myopathy, and dehydration as reviewed by Deem and Karesh, [47]. In Table 4, lists the drugs
commonly used in emergency situations in the field.
Figure 5. Collecting blood from the medial saphenous in an anesthetized jaguar (Panthera onca).
(Photo by Deem SL). - To view this image in full size go to the IVIS website at www.ivis.org . -
Drugs may be delivered either intramuscularly or intraveneously. If the intravenous route is chosen, the anesthesiologist
must be familiar with the drug he/she is administering and be sure to deliver at an appropriate rate. Vessels for delivering
drugs IV (and that are suitable for the collection of blood) include the jugular, cephalic, medial and lateral saphenous,
femoral, and lateral tail veins (Fig. 5).
The appropriate vessel to use will be determined on a case by case basis. For example, the jugular vein may be difficult
to approach for the administration of diazepam to a seizuring jaguar. The needle size will depend on the vessel used, but
generally the ideal needle size should be 1 - 1-1/2 inch and 18 - 22 gauge.
I. Respiratory Depression and Arrest - Results in tissue hypoxia caused by inadequate oxygenation of blood
hemoglobin and is probably the number one anesthetic emergency encountered in the field.
Diagnosis of respiratory depression/arrest is based on:
zthe jaguar taking few or no breathes (i.e., less than 4; no chest expansion) per minute;
zblue/gray mucous membrane (mm; gums);
zoxygen saturation is < 80% on pulse oximetry (if available).
During field immobilization there are a number of causes for respiratory depression/arrest including 1) drug-induced
depression of the respiratory center; 2) airway obstruction due to malpositioning, excessive salivation or regurgitation,
laryngeal edema; 3) pressure on the diaphragm from gastrointestinal contents; 4) excessive build up of carbon dioxide
which alters normal respiration; and 5) inapparent underlying disease process.
Treatment of respiratory depression / arrest should include the following:
1. DO NOT PANIC (this is true for all anesthetic emergencies!).
2. Do not administer any additional immobilization drugs.
3. Be sure the head and neck are in good positions (extended with no objects compressing them) so air can move
through the mouth and trachea. Be sure there is no vomit or foreign objects blocking the trachea.
4. Intubate immediately if an endotracheal tube (ETT) is available. Administer oxygen through the ETT using an
ambu bag, your own breath, or an oxygen tank (if available).
5. If no ETT or supplemental source of oxygen is available, use intermittent pressure on the chest to attempt to
move air through the lungs. The jaguar should already be in lateral recumbency. Push down firmly on the chest at
regular intervals (i.e., press for 1 second, wait for 1 second, press for 1 second and so on). Alternatively, you may
attempt mouth-to-mouth or mouth-to-nose resuscitation. Exhale into the jaguar’s mouth or nose for a count of 2
sec and then inhale away from the cat’s mouth/nose for a count of 2 sec.
6. Administer 1 - 2 mg/kg doxapram IV (or IM in the tongue muscle if one cannot quickly find a vein).
Note - Doxapram can cause arousal, especially in a cat immobilized with telazol, and caution for human safety
Table 4. Quick reference of emergency drugs for troubleshooting anesthetic emergencies in
the anesthetized free-ranging jaguar (Panthera onca) with amounts in milliliters to give to an
80 kg jaguar.
Drug Concentration Dose Amount
Atropine 2.25 mg/ml 0.04 mg/kg 1.4 ml
Diazepam 5 mg/ml 0.1 mg/kg 1.6 ml
Dexamethasone 4 mg/ml 2 mg/kg 40 ml
Doxapram 20 mg/ml 1.5 mg/kg 1.5 ml
Epinephrine 1:1000 0.02 mg/ml 1.6 ml
Lactated Ringers Solution n/a 20 ml/kg 1600 ml
Sodium bicarbonate 1 mEq/ml 1 mEq/kg 80 ml
must be considered if one elects to use this drug as a respiratory stimulant. Some veterinary anesthesiologist no
longer recommend the use of this drug. If respiratory arrest is not corrected with steps 1 - 5 above, the author
recommends the use of doxapram as a last attempt for resuscitation. If a person must inject the drug into the
tongue, he/she should be very careful not to traumatize the oral cavity.
7. Administer appropriate anesthestic antagonist if available (i.e., flumazenil, yohimbine, atipamezole). However,
do this cautiously as the antagonist will only reverse the drug it antagonizes and the jaguar may be semi-
anesthetized and difficult to handle after the antagonist is administered.
II. Cardiac Arrest - Is usually preceded by respiratory arrest and is defined as the loss of effective cardiac function
resulting in cessation of circulation. This is the most serious anesthetic emergency encountered during field
immobilization.
Diagnosis of cardiac arrest is based on:
zWeak or absent pulse or heart sounds;
zBlue/gray mucous membranes (gums);
zPoor capillary refill time measured by applying digital pressure to the mucous membrane until the mm turns pale
and then releasing the pressure and monitoring the seconds it takes until the mm color returns to normal (this
value should be < 2 sec);
zDilated pupils;
zCold extremities;
zLoss of consciousness (hard to evaluate if the animal is anesthetized).
The most common causes of cardiac arrest during field immobilization are 1) drug-induced; 2) respiratory failure leading
to hypoxia; 3) acid-base or electrolyte imbalance; and 4) underlying disease process.
Treatment of cardiac arrest should include the following:
1. Do not administer any additional immobilization drugs.
2. Be sure the animal can breathe prior to starting cardiac massage (see above).
3. Begin external cardiac massage. The jaguar should already be in lateral recumbency.
Apply firm pressure downward over the heart. Compression of the heart should be for a count of 1 and release for
a count of 1 with 60 - 100 cycles/minute. If an assistant is available he/she should palpate the femoral pulse to
ensure adequate pressure, to circulate blood, is being applied during cardiac compressions.
4. Administer 0.02 mg/kg of 1:1000 (1.0 mg/ml) epinephrine IV or intracardially and continue with external cardiac
massage. This dose is approximately 1.6 mg (1.6 ml) per 80 kg adult jaguar.
5. Administer 20 ml/kg cool Lactacted Ringer’s Solution as an IV bolus (i.e., a single rapid infusion).
6. If no response, repeat 4 above at 5 minute intervals indefinitely.
III. Seizures - Are defined as disturbances of cerebral function characterized by a violent, involuntary contraction or
series of contractions of the voluntary muscles.
Diagnosis is made based on clinical signs that include the following:
zUncontrolled muscle and/or whole body spasms;
zRigid extension of the limbs.
Causes include 1) drug-induced (i.e., ketamine and tiletamine); 2) trauma; and 3) hypoglycemia.
Treatment includes the following:
1. Administer 10 mg (total) diazepam IV slowly over 10 - 15 seconds.
2. Repeat step above if no improvements within 3 minutes.
3. Monitor body temperature to determine if secondary hyperthermia results from the seizure activity.
IV. Hyperthermia - Is defined as an increase in body temperature to a point where oxygen demand exceeds supply due
to increased metabolism.
Diagnosis of hyperthermia: easily determined by rectal thermometer.
Temperatures > 41ºC (105.8ºF) are true emergencies.
Causes of hyperthermia in field immobilization include 1) internal heat production due to excessive physical exertion; 2)
external heat absorption; 3) drug-induced compromise of thermoregulation; and 4) inability to use behavioral
thermoregulation.
Treatment of hyperthermia includes the following:
1. Do not administer any additional anesthetic agents.
2. Make sure the jaguar is in the shade.
3. Use portable "cold" packs that can be placed in the groin, axillae (armpit) and belly of the jaguar.
4. Cool the jaguar by applying water over the body and/or alcohol to the extremities (legs and feet).
5. Administer cold water enema if tubing is available.
6. Administer 20 ml/kg cool Lactacted Ringer’s Solution as an IV bolus (i.e., rapid fluid infusion).
7. Take the temperature every 5 - 10 minutes to determine if the temperature is decreasing. Continue to wet the
animal if the temperature remains high.
8. Administer antagonist IV (IM if a vein is not readily identified). However, do this cautiously as the antagonist
will only reverse the drug it antagonizes and the jaguar may be semi-anesthetized and difficult to handle after the
antagonist is administered.
9. If it is believed that the hyperthermia is due to muscle rigidity and a light plane of anesthesia, diazepam at a dose
of 5 - 10 mg/jaguar TOTAL can be administered slowly IV to decrease muscular activity.
Note - Hypothermia (< 35ºC = < 95ºF) - decreased body temperature to point of cellular death - is much less likely under
most field conditions in which jaguar will be immobilized. However, this may occur (i.e., high altitude regions) and
should be treated by warming the animal.
V. Wounds - Are often associated with the dart site as well as by trap or chase injuries. (Be especially cognizant of any
oral lesions and/or broken teeth).
Diagnosis: based on clinical signs. The severity of the wound will dictate the treatment modality chosen:
zPhysical examination to evaluate for traumatic lacerations and lesions;
zOral examination to evaluate for oral lesions and broken teeth.
Treatment should always include:
1. Clean the wound with a povidone-iodine or 2% chlorhexidine solution. If neither of these is available, use soapy
water.
2. If necrotic tissue is present and the field personnel are familiar with veterinary surgical techniques, debride the
dead tissue and repeat step.
3. Only suture those wounds that you KNOW are fresh (i.e., caused by the dart) and that require sutures to minimize
further tissue damage. Again, only field personnel who are familiar with veterinary surgical techniques should
attempt to suture any wounds.
4. Apply topical antibiotic and fly-strike ointment to wound site.
5. Administer long-acting antibiotic IM (i.e., Penicillin G benzathine 40,000 IU/kg IM).
6. Administer ivermectin 200 ug/kg SC (to prevent screw worm infestation at site of broken skin).
Treatment of broken teeth
It is imperative that a fractured tooth (most commonly a canine is broken during jaguar captures and immobilizations) be
repaired to minimize pain and infection associated with the tooth. A calcium hydroxide product (i.e., Dycal ®) can be
used to cap the tooth pulp. Instructions for application come with tooth repair kits.
Post-anesthetic Recovery
The recovery period is just as important for proper handling and monitoring as the induction and maintenance periods. It
is not uncommon for anesthetic related morbidity and mortality to occur during this period; in fact, most anesthetic
complications occur during induction and recovery. Although there are reversal drugs for the zolazepam component of
telazol (flumazenil), xylazine (yohimbine), and medetomidine (atipamezole), jaguar recoveries can not be completely
reversed with one specific antidote as is available for narcotic immobilization agents (i.e., carfentanil, etorphine)
commonly used in hoofstock. For this reason, it is important to ensure that the jaguar does not cause injury to itself or to
people involved in the immobilization during the recovery period.
During recovery, the jaguar should be positioned so that it can breathe easily and will not harm itself on objects near it.
The animal should be placed in lateral recumbency with the head and neck extended. Abrasive material should not be
under the head due to possible head movements that could lead to corneal abrasions. People in the area remain quiet and
should NOT stimulate the jaguar. It should recover at its own pace as it metabolizes the anesthetic agent(s). Stimulation
will not result in a faster recovery, but it may cause the jaguar to injure itself.
If the jaguar was originally captured in a box trap, it may be beneficial to let the animal recover in the cage where it is
dark and quiet. However, it must be remembered that when the cat is awake enough for release, the danger to field
personnel may be significant when opening the cage. While in the cage and recovering, the animal may also be
aggressive and cause harm to itself. Thus, if one is to use a box trap for recovery, it requires judgement to be sure the
jaguar is awake enough prior to release, but does not cause harm to itself while still in the cage. Alternatively, when no
trap is available (i.e., treed by dogs or darted from a blind), the animal can be placed in a quiet, padded (i.e., with leaf
litter), and protected (i.e., not near ledges, hard structures) area to recover on its own. Risks are involved with both
recovery methods.
References
1. Hoogesteijn R, Mondolfi E. The Jaguar. Caracas: Armitano Publishers 1992.
2. Sanderson EW, Redford KH, Chetkiewicz C-LB, et al. Planning to save a species: the jaguar as a model. Con Biol
2002; 16:58-72.
3. Swank WG, Teer, JG. Status of the jaguar - 1987. Oryx 1987; 23:14-21.
4. Crawshaw PG. Comparative ecology of ocelot (Felis pardalis) and jaguar (Panthera onca) in a protected subtropical
forest in Brazil and Argentina. Doctoral thesis, University of Florida, 1995.
5. Emmons LH. A field study of ocelots (Felis pardalis) in Peru. Revue d’ Ecologie (Terre Vie) 1997; 43:133-157.
6. Rabinowitz AR, Nottingham BG. Ecology and behaviour of the jaguar (Panthera onca) in Belize, Central America. J
Zool Lond 1986; 210:149-159.
7. Schaller GB, Crawshaw PG. Movement patterns of jaguar. Biotropica 1980; 12:161-168.
8. Jorgenson JP, Redford KH. Humans and big cats as predators in the Neotropics. Symp Zool Soc Lond 1993; 65:367-
390.
9. Taber AB, Novaro AJ, Neris N, et al. The food habits of sympatric jaguar and puma in the Paraguayan Chaco.
Biotropica 1997; 29:204-213.
10. Terborgh J. The big things that run the world - a sequel to E.O. Wilson. Con Biol 1988; 2:402-403.
11. Deem SL, Karesh WB, Weisman W. Putting theory into practice: wildlife health in conservation. Con Biol 2001;
15:1224-1233.
12. Ladiges WC, Foster JW, Jones MH. Malignant hemangioendothelioma in a jaguar (Panthera onca). J Zoo An Med
1981; 12:36-37.
13. Bossart GD, Hubbell G. Ovarian papillary cystadenocarcinoma in a jaguar (Panther onca). J Zoo An Med 1983;
14:73-76.
14. Frazier KS, Hines ME, Ruiz C, et al. Immunohistochemical differentiation of multiple metastatic neoplasia in a
jaguar (Panther onca). J Zoo Wildl Med 1994; 25:286-293.
15. Kollias GV, Calderwood-Mays MB, Short BG. Diabetes mellitus and abdominal adenocarcinoma in a jaguar
receiving megestrol acetate. J Am Vet Med Assoc 1984; 11:1383-1386.
16. Cirillo F, Ayala M, Barbato G. Giardiasis and pancreatic dysfunction in a jaguar (Panthera onca): case report,
evaluation, and comparative studies with other felines. In: Proceedings of the Am Assoc Zoo Vet 1990; 69-73.
17. Abdulla PK, James PC, Sulochana S, et al. Anthrax in a jaguar (Panthera onca). J Zoo An Med 1982; 13:151.
18. Appel MJG, Yates RA, Foley GL, et al. Canine distemper epizootic in lions, tigers, and leopards in North America. J
Vet Diagn Invest 1994; 6: 277-288.
19. Fransen DR. Feline infectious peritonitis in an infant jaguar. In: Proceedings of the Am Assoc Zoo Vet 1973; 261-
264.
20. Barr MC, Calle PP, Roelke ME, et al. Feline immunodeficiency virus infection in nondomestic felids. J Zoo Wildl
Med 1989; 20:265-272.
21. Brown EW, Yuhki N, Packer C, et al. Prevalence of exposure to feline immunodeficiency virus in exotic felid
species. J Zoo Wildl Med 1993; 24:357-364.
22. Patton S, Rabinowitz A, Randolph S, et al. A coprological survey of parasites of wild neotropical felidae. J Parasit
1986; 72:517-520.
23. Seymour KL. Panthera onca. Mammalian Species 1989; 340:1-9.
24. Hawkey CM, Hart MG. Haematological reference values for adult pumas, lions, tigers, leopards, jaguars and
cheetahs. Res Vet Sci 1986; 41:268-269.
25. International Species Information System. Medical animal record keeping system. Apple Valley, Minnesota, 1999;
www.worldzoo.org.
26. de Wet T. Physical capture of carnivores. In: McKenzie AA, ed. The Capture and Care Manual. South Africa:
Wildlife Division Support Services CC and The South African Veterinary Foundation, 1993; 255-277.
27. Wilson DE, Cole FR, Nichols JD, et al. Measuring and monitoring biological diversity. Standard Methods for
Mammals. Washington DC: Smithsonian Institution Press, 1996; 409 p.
28. Crawshaw PG. Capture methods of large felids, with special reference to the jaguar (Panthera onca).
www.savethejaguar.com 2002
29. Balser DS. Tranquilizer tabs for capturing wild carnivores. J Wildl Manag 1965; 29: 438-442.
30. Sahr DP, Knowlton FF. Evaluation of tranquilizer trap devices (TTDs) for foothold traps used to capture gray
wolves. Wildl Soc Bull 2000; 28:597-605.
31. Osofsky SA, Hirsch KJ. Chemical restraint of endangered mammals for conservation purposes: a practical primer.
Oryx 2000; 34:27-33.
32. Bush M. Remote drug delivery systems. J Zoo Wildl Med 1992; 23: 159-180.
33. Nielsen L. Chemical Immobilization of Wild and Exotic Animals. Ames: Iowa State University Press, 1999; 342 p.
34. Crawshaw PG. Recommendations for study design on research projects on neotropical felids. In: Felinos de
Venezuela-Biología, Ecología y Conservacíon. Memorias del Simposio Organizado por Fudeci, 1992; 187-222.
35. Lopez de Buen L, Aranda Sanchez JM. Nota zoologica. Anestesia de mamiferos silvestres con la combinacion
ketamina-xilacina. Biotica 1986; 67-71.
36. Quigley H. Ecology and conservation of the jaguar in the Pantanal Region, mato Grosso do Sul, Brazil. PhD
Dissertation. University of Idaho, 1987.
37. Quigley K. Hornocker Wildlife Institute Immobilization and Biological Sampling Protocols. Idaho: Hornocker
Wildlife Institute Inc.
38. Hoogesteijn R, McBride R, Sunquist M, et al. Medetomidine and rubber-padded leg-hold traps in Venezuelan cat
studies. Cat News 1996; 25:22-23.
39. Morato RG, Moura CA, Crawshaw PG. Chemical restraint of free ranging jaguars (Panthera onca) with tiletamine-
zolazepam combination. In: Jaguars in the New Millenium Proceedings, 2001.
40. Morato RG. Reprodução em onca pintada (Panthera onca): avaliação do método de contenção e de obtenção de
sêmen, caracterização do ejaculado, biometria testicular, níveis séricos de testosterona e sazonalidade. Universidade de
São Paulo, São Paulo: Master Thesis 1997; 120 p.
41. Steffey EP. Inhalation anesthetics. In: Lumb and Jone’s Veterinary Anesthesia, 3rd ed. Thurmon JC, Tranquilli WJ,
Benson GJ eds. Baltimore: Williams and Wilkins, 1996; 297-329.
42. Armstrong D. Adverse reactions to telazol in tigers. Tiger Beat 1990; 3:11.
43. Evans AT. Anesthetic emergencies and accidents. In: Thurmon JC, Tranquilli WJ, Benson GJ, eds. Lumb and Jone's
Veterinary Anesthesia, 3rd ed. Philadelphia: Wilkins and Wilkins Co, 1996; 849-860.
44. Fowler ME. Medical problems during restraint. In: Restraint and Handling of Wild and Domestic Animals, 2nd ed.
Ames: Iowa State University Press, 1995; 78-99.
45. International Wildlife Veterinary Services, Inc. Wildlife Restraint Series. Salinas: International Wildlife Veterinary
Services Inc, 1991.
46. Kreeger TJ. Handbook of Wildlife Chemical Immobilization. Laramie: International Wildlife Veterinary Services
Inc, 1996.
47. Deem SL, Karesh WB. The Jaguar Health Program Manual. www.savethejaguar.com/fieldvet_health_manual.pdf,
2001; 1-47.
All rights reserved. This document is available on-line at www.ivis.org. Document No. B0183.1204.