Great horned owls (GHOWs; Bubo virginianus) are widely distributed in the Americas and are one of the most common Strigiformes species presented to wildlife rehabilitation centers in North America.1–3 Traumatic injuries such as wounds and fractures from collisions are the major reported causes for the admission of birds of prey to veterinarians in rehabilitation centers.3–8 Therefore, there is a need to study analgesic drugs for use in acute pain management when these animals are admitted. Improvement in pain management will provide a faster return to normal function following trauma or surgery and overall improvement of animal welfare.
Hydromorphone is a semisynthetic full μ-opioid agonist that has been used in human medicine for postoperative and cancer-related pain.9,10 In humans, hydromorphone is estimated to be approximately 7.5 to 8.5 times more potent than morphine for chronic pain and between 5 and 7 times more potent than morphine for acute pain.11 Hydromorphone has been extensively studied in dogs and cats12–16 and other species of mammals such as horses17 and guinea pigs18 and, together with other μ-opioid agonists, is the recommended first choice for treating moderate to severe pain.19 In mammals, hydromorphone is metabolized to form the metabolite hydromorphone-3-glucuronide (H3G).20 Hydromorphone pharmacokinetics have been previously investigated in orange-winged Amazon parrots (Amazona amazonica),21 cockatiels (Nymphicus hollandicus),22 and American kestrels (Falco sparverius),23 however, to our knowledge, the metabolite H3G has not been evaluated in any avian species to date.
Recent studies24–28 evaluating the efficacy of opioid drugs using the American kestrels as a model for raptorial species support the clinical use of opioids in pain management. However, there are limited studies evaluating these drugs in Strigiformes and most dosage recommendations for opioids come from extrapolation of data from other avian species. The pharmacokinetics of butorphanol tartrate have been investigated in GHOWs, but the efficacy of this drug as an analgesic has not been evaluated in this species.29 A recent pharmacodynamic study30 evaluating the thermal antinociceptive effects of hydromorphone in GHOWs at 0.3- and 0.6-mg/kg doses resulted in significantly higher mean thermal foot withdrawal thresholds at 0.5, 1.5, and 3 hours and at 0.5 and 1.5 hours, respectively, when compared to the control group. Other drugs used for pain management, like meloxicam and gabapentin, have also been studied in GHOWs, providing additional information on their pharmacokinetic profiles.31,32
The objective of this study was to measure plasma concentrations and pharmacokinetic parameters of hydromorphone and to determine whether it forms the metabolite H3G following the administration of hydromorphone (0.6 mg/kg) IM and IV for each route of administration. It was hypothesized that a single dose of hydromorphone administered IM would result in high bioavailability, a pharmacokinetic profile comparable with other avian species studied, and would be metabolized to H3G.
Materials and Methods
Animals
Six adult healthy great horned owls, 3 females and 3 males of ages ranging between 5 and 28 years old with mean ± SD body weights of 1.160 ± 0.240 kg, were utilized in this study. The same 6 birds also participated in a previous pharmacodynamic study30 8 months prior to the evaluation of the thermal antinociceptive effects of hydromorphone. These birds are deemed nonreleasable due to previous orthopedic injuries or visual deficits and are permanent residents of the California Raptor Center, School of Veterinary Medicine, University of California-Davis. Other than their initial injuries, the GHOWs were deemed healthy to participate in the study based on history, physical examination, and a complete blood count performed during their annual examinations. None of the birds received other medications or underwent any anesthetic procedures for at least 1 month prior to the start of the study. Prior to the day of sample collection, 3 of the owls were housed together in a large display exhibit and 3 were housed individually in 8 X 10 ft enclosures. The birds were offered frozen-thawed day-old chicks (4 to 5 chicks per owl) and water ad libitum in water bowls daily. During the days of sample collection, each bird was kept in individual carriers to minimize capture handling and stress. The study was approved by the Institutional Animal Care and Use Committee at the University of California-Davis (protocol No. 22094).
Study design
For this study, 2 experiments were performed. In preparation for each experiment, food was offered in the early evening but removed within 2 hours if not consumed to ensure a 12-hour fasting period. For the IM experiment, the owls were manually restrained with leather gloves, and a dose of 0.6 mg/kg of a 10-mg/mL solution of hydromorphone hydrochloride (Pfizer Hospira Inc) was administered in the right pectoral muscles using a 0.5-mL insulin syringe with a 28-gauge needle (BD Micro-Fine, BD Lo-Dosep; Becton, Dickinson and Co). Following a 6-week washout period, the owls were manually restrained, and the same medication was administered at the same dose into the left jugular vein with the same size needle and syringe. All injections IM and IV were performed by the same researcher (DSMG).
Blood samples were collected at 5 minutes and 0.5, 1.5, 2, 3, 6, 9, and 12 hours postadministration. For both experiments and for all time points, 0.4 mL of whole blood was collected from the right jugular vein or medial metatarsal veins using a 0.5-mL insulin syringe with a 28-gauge needle (BD Micro-Fine, BD Lo-Dose), with a total volume collected of approximately 3.2 mL/bird, which was < 10% of blood volume and < 1% of the bird’s body weight. Needles were removed from syringes, and blood samples were transferred to lithium-heparin tubes (BD microtainer), placed in a cooler with ice packs, and centrifuged (3,500 X g for 8 minutes) within 30 minutes of collection. Plasma was extracted using a disposable transfer pipette, placed into labeled 2-mL cryovials, and stored at –80 °C for 139 days for the IM samples and 97 days for the IV samples until analysis; all samples were analyzed at the same time. At the end of both experiments, each bird received supportive care, which included the administration of 50 mL/kg, SC lactated Ringer’s solution (Lactated Ringer’s Injection, USP; Baxter) to compensate for the fasting and intravascular volume loss and 0.5 mg/kg, SC meloxicam (OstiLox; Vetone) for additional pain management after multiple phlebotomies. The owls were visually monitored at each time point for adverse effects such as abnormal mentation, tremors, and gastrointestinal abnormalities (vomiting, diarrhea).
Determination of plasma hydromorphone concentrations
Five minutes prior to the IM injection, a blood sample of 1 mL was collected from each bird to harvest blank plasma for preparation of the calibration curve and quality control samples for liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis; the collected blank plasma was used for both the IM and IV experiments. Plasma calibrators were prepared by dilution of the hydromorphone and hydromorphone-glucuronide working standard solutions (Cerilliant) with drug-free owl plasma to concentrations ranging from 1 to 500 ng/mL. Calibration curves and negative control samples were prepared fresh for each quantitative assay. Quality control samples (owl plasma fortified with analyte at 3 concentrations within the standard curve) were included with each sample set as an additional check of accuracy.
Prior to analysis, 50 µL of plasma was diluted with 150 µL of acetonitrile (ACN):1 M acetic acid (9:1, vol:vol) containing 0.01 ng/μL of the d3-HYD internal standard (Cerilliant) to precipitate proteins. The samples were vortexed for 1.5 minutes, refrigerated for 20 minutes, vortexed for an additional 1 minute, and centrifuged at 4,300 rpm/3,830 X g for 10 minutes at 4 °C, and 10 µL was injected into the LC-MS/MS system. The concentrations of hydromorphone and H3G were measured in plasma by LC-MS/MS using positive electrospray ionization [ESI(+)]. Quantitative analysis was performed on a TSQ Altis triple quadrupole mass spectrometer coupled with a Vanquish liquid chromatography system (Thermo Scientific). The spray voltage was 3,500 V, the vaporizer temperature was 350 °C, and the sheath and auxiliary gas were 50 and 10, respectively (arbitrary units). Product masses and collision energies of each analyte were optimized by infusing the standards into the TSQ Altis. Chromatography employed a Zorbax Eclipse-XDB-Phenyl 3 X 100-mm, 3-μm column (Agilent Technologies, Inc) and a linear gradient of ACN in water with a constant 0.2% formic acid at a flow rate of 0.45 mL/min. The initial ACN concentration was held at 0% for 0.2 minutes, ramped to 75% over 3.8 minutes, and ramped to 95% over 0.3 minutes before reequilibrating for 4 minutes at initial conditions.
Detection and quantification were conducted using selective reaction monitoring of the initial precursor ion for hydromorphone (mass-to-charge ratio [m/z], 286.1), HG3 (m/z, 462.2), and the internal standard d3-HYD (m/z, 289.1). The response for the product ions for hydromorphone (m/z, 128.1, 157.1, and 185.1), HG3 (m/z, 185.1, 227.0, and 286.1), and the internal standard d3-HYD (m/z, 185.1) was plotted and peaks at the proper retention time integrated using Quanbrowser software (Thermo Scientific). Quanbrowser software was used to generate calibration curves and quantitate analytes in all samples by linear regression analysis. A weighting factor of 1/X was used for all calibration curves.
The concentration-response relationships (relationship between calibrators and the LC-MS/MS instrument response) for hydromorphone and H3G were linear and gave correlation coefficients of 0.99 or better. The precision and accuracy of the assay were determined by assaying quality control samples in replicates (n = 6; Table 1). Accuracy was reported as percent nominal concentration and precision as percent relative standard deviation. The technique was optimized to provide a limit of quantitation of 1 ng/mL and a limit of detection of approximately 0.5 ng/mL for both hydromorphone and hydromorphone-glucuronide.
Accuracy and precision values for LC-MS/MS analysis of hydromorphone and hydromorphone-3-glucuronide concentrations in great horned owl (Bubo virginianus) plasma.
| Concentration (ng/mL) | Accuracy (% nominal concentration) | Precision (% relative SD) | |
|---|---|---|---|
| Hydromorphone | |||
| 7.5 | 113 | 4.0 | |
| 35.0 | 98.0 | 5.0 | |
| 160 | 99.0 | 3.0 | |
| Hydromorphone-3-glucuronide | |||
| 7.5 | 97.0 | 2.0 | |
| 35.0 | 96.0 | 4.0 | |
| 160 | 108 | 3.0 |
Pharmacokinetic analysis
The peak concentration (Cmax) and time to peak plasma concentration (tmax) were determined by visual inspection of the concentration-time data. Noncompartmental analysis was used for the determination of pharmacokinetic parameters using a commercially available computer software program (Phoenix Winnonlin v8.3). The area under curve (AUC) from time 0 to infinity (AUC0–∞) was determined using the linear up-log down trapezoidal rule. Mean absorption time (MAT) following IM administration was calculated as the difference between IM mean residence time (MRTIM) and IV mean residence time (MRTIV).
Results
No adverse effects were noted for the duration of the study. All birds were bright and responsive at each collection time point. The birds were quiet after handling but did not have other outward signs of stress or sedation. There was no change in urofeces as subjectively assessed. All GHOWs were alert and able to return to their enclosures 12 hours postdrug administration. The following morning after sample collection, all birds were reported to be bright and alert and had eaten their diets.
Plasma concentration versus time curves for IV and IM hydromorphone and H3G are presented (Figures 1 and 2). The calculated pharmacokinetic parameters are summarized (Tables 2 and 3) for the IV and IM parameters, respectively. At 6 and 9 hours postadministration IV, there were 1 and 4 birds, respectively, with plasma concentration below the limit of quantification. At 12 hours postadministration, all birds, except for 2 in the IM administration experiment, had hydromorphone concentrations below the limit of quantitation. Hydromorphone-3-glucuronide was measured in all individuals at the 5-minute time point from the IV route but was only detected in 4 birds via the IM route at the same time point. Hydromorphone-3-glucoronide was measured in the remaining time points in all birds except for 1 bird in the IV experiment at the 9- and 12-hour time points.
Mean ± SD hydromorphone (black circles) and hydromorphone-3-glucuronide (white circles) plasma concentrations over time in 6 great horned owls after a single dose of hydromorphone hydrochloride (0.6 mg/kg, IV). Time 0 was the time of drug injection. Values below the limit of quantitation or nondetectable are not represented in the figure.
Citation: American Journal of Veterinary Research 84, 5; 10.2460/ajvr.22.12.0218
Mean ± SD hydromorphone hydrochloride (black squares) and hydromorphone-3-glucuronide (white squares) plasma concentrations over time in 6 great horned owls after a single dose of hydromorphone hydrochloride (0.6 mg/kg, IM).
Citation: American Journal of Veterinary Research 84, 5; 10.2460/ajvr.22.12.0218
Mean ± SD values of noncompartmental pharmacokinetic parameters of hydromorphone hydrochloride and hydromorphone-3-glucuronide following administration of a single dose of hydromorphone hydrochloride (0.6 mg/kg) IV to 6 great-horned owls (Bubo virginianus).
| Parameter | Hydromorphone | Hydromorphone-3-glucuronide |
|---|---|---|
| Cl (mL/min/kg) | 62.11 ± 14.65 | – |
| Vdss (L/kg) | 4.29 ± 0.50 | – |
| Cmax (ng/mL) | – | 26.9 ± 10.9 |
| t1/2λ (h) | 1.35 ± 0.59 | 4.04 ± 2.21 |
| λz (1/h) | 0.59 ± 0.25 | 0.20 ± 0.08 |
| AUC0–∞ (h·ng/mL) | 168.45 ± 38.10 | 102.8 ± 42.4 |
| AUCextrap (%) | 2.17 ± 1.16 | 11.4 ± 5.75 |
| AUClast (h·ng/mL) | 165 ± 38.35 | 92.1 ± 42.2 |
| AUMC0–∞ (h·h·ng/mL) | 206.37 ± 78.25 | 498.7 ± 220.4 |
| AUMClast (h·h·ng/mL) | 178.08 ± 70.87 | 300 ± 150.6 |
| MRT–∞ (h) | 1.19 ± 0.22 | – |
λz = Slope of terminal portion of plasma concentration curve. AUC0–∞ = Area under the curve from time 0 to infinity. AUCextrap = Percentage of the area under the curve that was extrapolated to infinity. AUClast = Area under the curve to the last measurable concentration. AUMC0–∞ = Area under the moment curve from 0 to infinity. AUMClast = Area under the moment curve to the last measurable concentration. Cl = Total systemic clearance. Cmax = Maximal plasma concentration. MRT–∞ = Mean residence time to infinity. t1/2λ = Terminal half-life. Vdss = volume of distribution at steady state.
Mean ± SD values of noncompartmental pharmacokinetic parameters of hydromorphone hydrochloride and hydromorphone-3-glucuronide following administration of a single dose of hydromorphone hydrochloride (0.6 mg/kg) IM to 6 great horned owls (Bubo virginianus).
| Parameter | Hydromorphone | Hydromorphone-3-glucuronide |
|---|---|---|
| Cmax (ng/mL) | 225.46 ± 0.21 | 41.14 ± 12.00 |
| tmax (h) | 0.22 ± 0.22 | 1.5 ± 0.54 |
| t1/2λ (h) | 1.62 ± 0.36 | 4.65 ± 3.37 |
| λz (1/h) | 0.44 ± 0.08 | 0.19 ± 0.067 |
| AUC0–∞ (h·ng/mL) | 278.55 ± 44.10 | 242.7 ± 78.10 |
| AUCextrap (%) | 1.01 ± 0.21 | 198.4 ± 14.80 |
| AUClast (h·ng/mL) | 275.8 ± 44.0 | 198.4 ± 46.6 |
| AUMC0–∞ (h·h·ng/mL) | 434.6 ± 97.4 | 1,847.4 ± 1,895.4 |
| AUMClast (h·h·ng/mL) | 399.2 ± 87.8 | 781.3 ± 197.1 |
| MRT–∞ (h) | 1.56 ± 0.24 | – |
| MAT (h) | 0.37 ± 0.29 | – |
| F (%) | 170.8 ± 37.6 | – |
F = Bioavailability. MAT = Mean absorption time. tmax = Time to peak plasma concentration.
See Table 2 for the remainder of the key.
Discussion
The objective of the present study was to establish the pharmacokinetic parameters of hydromorphone and whether it formed its metabolite H3G following administration of 0.6 mg/kg, IM and IV. This dose was elected based on the pharmacodynamic research of hydromorphone in GHOWs, where the highest effective dose evaluated of 0.6 mg/kg was associated with increased thermal foot withdrawal times when compared with the control group.30 Hydromorphone IM was rapidly absorbed, had high bioavailability, and had a short half-life (t1/2). The dose evaluated in the present study was well tolerated and not associated with significant adverse effects. However, as interventions such as venipuncture and restraint can confound the accurate assessment of agitation and sedation scores, a dedicated study focusing solely on such scores for hydromorphone may prove valuable but was not the main focus of this study.
The bioavailability of IM hydromorphone was > 100% for all birds. Although values exceeding 100% are not common, they are possible and are attributable to either experimental or mechanistic factors.33 Experimental factors do not appear to explain the > 100% IM bioavailability. The same formulation was used for both routes of administration; for IV administration, the drug was directly injected as opposed to using a catheter or tubing where the drug might be expected to adhere to tubing; the same group of birds received the drug via both routes of administration; and there were no apparent analytical assay concerns. It was also considered that different storage duration leading to drug degradation could be an explanation for IM bioavailability > 100%. However, the IM experiment was performed first and the IM samples were stored longer than the IV samples. If IM samples were frozen longer and drug degradation had occurred, it would be expected that the bioavailability would be lower, not higher. Additionally, a study34 identified that hydromorphone is stable when frozen at −20 °C for over 3 years. Even though H3G was not measured in that study, it could be speculated that if it was not stable, the concentration of hydromorphone would increase as the metabolite degrades. One common mechanistic factor that can also be ruled out in the current study is nonlinear elimination kinetics, which can result in erroneous pharmacokinetic calculations. As the terminal half-life was comparable between routes of administration in the current study, this does not appear to be a factor. The > 100% bioavailability is likely attributable to a more complex mechanistic factor and requires further investigation.
In mammals, hydromorphone has been associated with a short t1/2 and rapid clearance (Cl). In horses, the mean t1/2 at the highest dose evaluated (0.08 mg/kg, IV) was 41.2 minutes with a mean Cl of 92.7 mL/min/kg.17 In dogs, following a dose of 0.1 mg/kg, IV, the t1/2 was 34 minutes and Cl 106.8 mL/min/kg35; however, in cats, the same dose resulted in a mean terminal t1/2 of 97.8 minutes, and clearance was significantly slower at 27.6 mL/kg/min, which was associated with their lower capacity to glucorinidate.14,36 In the current study, a higher dose of hydromorphone than those previously studied in mammals resulted in a longer t1/2 of 1.62 ± 0.36 hours and a Cl of 62.1 ± 14.7 mL/min/kg. These differences in pharmacokinetic parameters likely represent pharmacokinetic differences between mammals and avian species and/or dose-dependent pharmacokinetics. This is of particular interest given that other opioid medications have been classified as poor candidates for allometric scaling,37 and it is a good example to demonstrate that dedicated avian species-specific pharmacokinetic studies are needed for each drug studied.
There are more similarities and comparable parameters with other avian species, including the shorter elimination half-life, rapid plasma clearance, and large volume of distribution.21–23 For example, in American kestrels (Falco sparverius) at the same dose of 0.6 mg/kg IV, the t1/2 was 1.25 h, Cl was 62.32 mL/min/kg, and volume of distribution at steady state (Vdss) was 4.26 L/kg, which were very similar values to those described in the current study. In orange-winged Amazon parrots (Amazona amazonica) following administration of almost twice the dose administered to the GHOWs (1 mg/kg hydromorphone, IM), similar values for Vdss 4.24 L/kg, Cl 64.2 mL/min/kg, and t1/2 1.74 h were obtained.21 The bioavailability in orange-winged Amazon parrots was high at 97.6 ± 61.1%21; however, the results of bioavailability (F) > 100% as seen on the GHOWs were not observed in other species.
A dose of 0.6 mg/kg of hydromorphone in GHOWs yielded plasma concentrations > 1 ng/mL for 9 hours after IM and 6 hours after IV administration. Thermal antinociceptive effects were detected at 1.5 hours after administration of that dose of hydromorphone in the pharmacodynamic counterpart30 of this study, which corresponded with a mean hydromorphone plasma concentration of 57.7 ± 8.43 ng/mL at that time point. These concentrations would not represent the minimum effective concentrations in GHOWs considering that in the pharmacodynamic counterpart of the current study, a dose of 0.3 mg/kg IM resulted in thermal antinociceptive effects at 3 hours, which was associated with much lower plasma concentrations. In fact, in the counterpart pharmacodynamic study,30 it was speculated that the shorter duration of action of the 0.6 mg/kg when compared to the 0.3 mg/kg might have been due to a type II error. In a similar set of studies27 in American kestrels, mean plasma hydromorphone concentration > 1 ng/mL would have been associated with thermal antinocioception. In orange-winged Amazon parrots, also using a similar methodology to the studies being discussed, the lowest mean plasma concentrations that would have been associated with thermal antinociception was 17.4 ng/mL.21 In humans, minimum effective plasma concentrations associated with analgesia were found to be 4 ng/mL.38 In a different study,39 plasma concentrations of 20 ng/mL in humans produced only 50% of maximum pain relief.
In mammals, hydromorphone is metabolized primarily by conjugation with glucuronic acid to form the metabolite H3G in a similar manner to morphine that forms morphine-3-glucuronide (M3G), the main metabolite of morphine.20 In the study reported here, H3G was detectable in all birds at 30 minutes following drug administration by both the IV and IM routes and had measurable plasma concentrations up to 12 hours postadministration, which was the last time point measured. The quantifiable concentrations of this metabolite suggest that hydromorphone in GHOWs may have a similar metabolic profile as mammals. In veterinary medicine, this metabolite has been documented in horses, with a linear concentration-response relationship between hydromorphone and H3G,17,40 similar to what was found in the present study. In horses, at a dose of 0.04 mg/kg IV, H3G was detected up to 18 hours after administration; in the present study, all owls had quantifiable plasma concentrations with a mean of 1.96 ng/mL at the 12-hour time point. For horses, the mean Cmax of H3G was 52.6 ± 7.8 ng/mL and the t1/2 was 4.55 hours.17 In GHOWs at a much higher dose of 0.6 mg/kg, the mean Cmax was 26.9 ± 10.9 ng/mL. The mean t1/2 of H3G is longer than that of hydromorphone in this species; this should be taken into account when considering multiple and frequent dosing, as dose-dependent neuroexcitatory effects have been described in rats.41
To the authors’ knowledge, this is the first study to document the presence of the H3G metabolite in an avian species. It is unknown if H3G has analgesic effects; however, the structural analog M3G has been associated with neuroexcitant signs in rats without analgesic effects. In contrast, the morphine metabolite morphine-6-glucuronide has been associated with analgesic effects,20,42 but an analgesic analog metabolite has not been identified for hydromorphone. A study41 investigating the neuroexcitant effects of H3G in rats revealed that it can induce dose-dependent-excitatory behaviors such as myoclonus, changes in body postures, and tonic-clonic seizures. In that study,41 the authors also found H3G had more potent neuroexcitatory effects than the M3G analog. In the present study, there were no obvious excitatory behaviors observed in the GHOWs. In the pharmacodynamic study,41 0.3 and 0.6 mg/kg of hydromorphone were associated with mild to moderate sedation and in 2 owls were associated with mild tremoring after administration of the 0.6 mg/kg at the 30 min time point evaluation. Increased agitation scores were observed in orange-winged Amazon parrots after administration of higher doses of 1 to 2 mg/kg of hydromorphone. Speculation could be made that increased agitation could be explained by the higher concentrations of the H3G metabolite, but further studies are necessary to confirm this.43
We acknowledge some limitations of the present study, including the small sample size, which reduces the power of the study and increases the margin of error. A larger sample would have allowed comparison between different variables such as sex and age group. However, maintaining a large, uniform population of wild species is challenging and not feasible in most settings. A cross-over study design would have been better suited for a study of this nature and, in this case, would have elucidated if the increased bioavailability was associated with an experimental factor. Agitation and sedation scores were not monitored given that repeated interventions such as phlebotomy could interfere with the interpretation, and further studies evaluating the agitation and sedation scores for GHOWS are required.
As initially hypothesized, the pharmacokinetic parameters for hydromorphone following IV or IM administration obtained from GHOWs were determined and found to be comparable to other species of birds. Hydromorphone IM rapidly attained plasma concentrations, had high bioavailability and a short half-life (t1/2), and was metabolized to H3G. This study documents the presence of H3G in an avian species. This metabolite has been associated with neuroexcitatory effects in rats; therefore, this must be considered in different individuals and with more frequent dosing. Further studies, including investigation of multiple doses and different routes of administration such as oral or intranasal, are needed to explore all potential benefits, safety, and adverse effects of hydromorphone in GHOWs.
Acknowledgments
The authors thank Julie Cotton, William Thein, and all the volunteers at the California Raptor Center for providing support, resources, and space to conduct this study. The authors also thank Jenna Gerds, Lexi Durant, Quinn Neil, Hikaru (Ray) Shiraishi, and Marissa Monopoli for their assistance in owl handling, care, and sample processing.
This study was supported by the American Association of Zoo Veterinarians Wild Animal Health Fund.
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