Quantitative and Qualitative Behavioral Measurements to Assess Pain in Axolotls (Ambystoma mexicanum)

Abstract

Effective pain relief in animals relies on the ability to discern pain and assess its severity. However, few objective measures exist to assess the presence and severity of pain in axolotls, and few resources are available regarding drugs and appropriate doses to provide pain relief in this species. This study evaluated behavioral tools for cageside pain assessment and validated a reproducible and reliable quantitative method to evaluate analgesic efficacy in axolotls. Animals were divided into control and treatment groups (n = 6 per group); treatment groups received buprenorphine through injection (50 mg/kg every 24 h for 48 h intracelomically) or butorphanol immersion (0.50 or 0.75 mg/L every 24 h for 48 h). Qualitative behavioral tests, adapted from other amphibian studies, included tapping on the home tank, directing water jets or physically touching specific anatomic points on the animal, and placing a novel object in the home tank. Quantitative methods used to produce noxious stimuli were the acetic acid test and von Frey aesthesiometers. Animals that were treated with analgesics did not demonstrate a significant difference compared with controls during behavioral assessment at 1, 6, 12, 25, 30, and 48 h after analgesia administration. The acetic acid test revealed a reproducible, concentration-dependent pain response. However, a significant difference in the AAT response was not observed between control and treated groups with the tested analgesics and doses.

Although several differences have been demonstrated in pain responses of frogs, newts, and salamanders, A- and C-type nociceptive fibers are present in most vertebrate animals, including amphibians.^4,10,23 Although axolotls (Ambystoma mexicanum, also known as Mexican salamanders) are classified in a different family and order from newts and frogs, respectively, pain receptors are likely conserved within the class. However, nociceptor fiber distribution and number may vary.²³ Spinothalamic projections (by means of a brainstem–thalamus tract) conveying cutaneous sensory information to the thalamus are not well understood in amphibians but are known to exist.^15,26 In addition, thalamocortical projections that convey sensory signals to the telencephalon are poorly organized and only contain scant numbers of fibers in amphibians.²⁶ The neuroanatomy of frogs²⁹ suggests the transmission of noxious stimuli (nociception) and the processing of the sensory information (pain) is poorly represented in amphibians and that most CNS pathways are related to spinal (that is, does not ascend to the brain) and long-loop (that is, ascends to the brain) reflexes to the brain stem and thalamus.¹³ Nevertheless, the presence of sensory projections to the brains of amphibians suggests these animals perceive pain and that appropriate analgesics should be used whenever performing experiments that can cause pain.

Axolotls are capable of robust, life-long epimorphic regeneration and can recover from virtually any nonfatal injury.²⁰ Their versatility as a research model to study developmental biology and tissue regeneration has led to their active use in laboratory research since the 1860s.⁷ Over the years, much has been published about the general anatomy, biology, and behavior of axolotls; recently the 32-Gb axolotl genome has been sequenced and assembled,¹⁶ thus increasing the utility of the animal for developmental, evolutionary, and regenerative studies. However, despite the use of this species for more than 150 y in experimental research, sometimes involving invasive surgical procedures such as limb amputation, guidance is sparse regarding the use of analgesics in axolotls.

Federally funded animal research must adhere to the Public Health Service Policy on Humane Care and Use of Laboratory Animals, which states that “procedures that may cause more than momentary or slight pain or distress to animals will be performed with appropriate sedation, analgesia, or anesthesia” unless the withholding of sedation, analgesia, or anesthesia is justified for scientific reasons in writing by the investigator.¹² Given the nature of regenerative medicine research, animals frequently undergo surgical injury for assessment of the healing process. Providing appropriate pain relief in axolotls is important for maintaining animal welfare and adhering to federal regulations.

The 2 primary classes of analgesics available to provide pain relief in research animals are opioids and NSAID. The decision to use one class of drugs over another is dependent on desired clinical outcomes, degree of expected pain, and effect of the drugs on scientific outcomes. In the field of regenerative medicine, the early stages of tissue repair and healing rely on a robust immune response.¹⁹ Consequently, NSAID usually are avoided, whereas opioids are typically first-line drugs of choice when studying tissue regeneration. Still, no choice of drug is without complications. For example, morphine and fentanyl suppress the immune system, whereas buprenorphine has a more favorable immunomodulatory and neuroendocrine profile.^8,14,21 In addition, as a class of analgesics, opioids have well-known side effects in mammals, including but not limited to immunologic effects, hormonal changes, increased sedation or sleep disturbances, induction of hyperalgesia, reduced psychomotor performance, respiratory depression, gastrointestinal and genitourinary dysfunction, pruritus, urticaria, and cardiac effects due to increased histamine release.^2,6 Therefore, when studying analgesic efficacy and effects of analgesics in laboratory animal research, especially in uncommon model animals, it is important to differentiate behavior induced by the drugs compared with the animals’ response to acute or chronic noxious stimuli. In the current study, we observed and recorded baseline behavior in all subjects to establish each animal's normal status. We then used this baseline as an index of the resulting behavior after each animal received each analgesic compound. This methodology was instituted to ascertain whether observed changes were purely associated with the administration of the analgesics.

The few studies that are available concerning antinociception in amphibians have used frogs (Xenopus laevis and Rana pipiens) and newts (Notophthalmus viridescens). Studies in frogs (X. laevis and R. pipiens) have described mechanical, thermal, and chemical methods to assess pain and analgesic efficacy.³⁰ In addition, increased latency to respond to noxious stimuli after the administration of morphine suggests that, as in mammals, frogs have an endogenous opioid-responsive system.²⁴ Various quantitative testing methods have been developed to measure pain in laboratory animals. Specific tools to measure different modalities of nociception in laboratory animals include electronic and manual aesthesiometers to measure mechanical sensitivity and hot-plate or radiant heat devices to measure thermal sensitivity. Devices relying on thermal stimuli to induce pain are difficult to adapt to aquatic species. Chemical nociceptive sensitivity can easily be assessed by using the acetic acid test (AAT),¹⁸ a testing method that is easily performed by using common wet-lab supplies. Described in 1983 for Northern leopard frogs (R. pipiens), the AAT was developed because the investigator hypothesized that previous pain studies in amphibians used inappropriate modalities to induce pain in frogs.¹⁸ The AAT is performed by diluting glacial acetic acid to make a 15-M stock, after which serial dilutions are made to create equally spaced (logarithmic) test solutions. Starting from the lowest concentration, testing is performed by placing a small volume of the acetic acid solution onto the animal's skin and continues with increasing concentrations until the animal responds—a point noted as the nociceptive threshold. In light of the published use of AAT and aesthesiometers to assess pain in amphibians,^25,27,28,30 these methods were modified for use in the axolotls in the current study.

Literature searches revealed scant published scientific studies investigating general signs of pain or distress in axolotls or recommending analgesic doses. Given evidence from prior studies in amphibians on pain relief after the administration of opioids and the further characterization of cutaneous nociceptors, amphibians sense noxious and nonnoxious thermal, mechanical, and chemical stimuli through Aδ and Aβ nociceptors and polymodal C-fiber nociceptors, the same signaling pathways as those found in mammals. Given that endogenous opioid receptors are present in amphibians, the current study aimed to evaluate 2 opioid-based drugs, previously studied in the newt,¹³ to provide pain relief in axolotls. Recent studies comparing newt and axolotl forelimb regeneration have shown that proposed cellular mechanisms underlying the repair of skeletal muscle after limb amputation follow different pathways in these 2 amphibians.²² The considerable diversity in achieving epimorphic regeneration, even in these 2 related urodeles, suggests that sufficient species-specific differences exist to suggest that simply translating protocols from newts to axolotls is ill-advised.⁷

Prior to any testing of analgesics, we tested the repeatability and reproducibility of the 2 quantitative pain assessments in axolotls. The 2 assessments provide different insights into stimulus processing—noxious stimuli as used in AAT are detected through cutaneous chemical nociceptors and conduct information along unmyelinated C fibers and thinly myelinated Aδ fibers, whereas nonnoxious stimuli activate cutaneous mechanoreceptors that rapidly conduct along Aβ-like fibers.³⁰ From our review of literature and pilot assessments, we hypothesized that buprenorphine and butorphanol will induce nociceptive changes in axolotls, as measured by using an adapted AAT or von Frey fiber (vF) qualitative measurement system, but will not produce behavioral changes at published doses used in other amphibian species.

Materials and Methods

Animals.

Male, adult, breeding and nonbreeding wild-type axolotls (weight, exceeding 60 g) were purchased from the Ambystoma Genetic Stock Center (University of Kentucky; Lexington, KY). The animals were individually housed in open-top polypropylene, static, mouse cages (19.0 × 10.5 × 6.1 in.). For environmental enrichment purposes, a single commercial rodent tunnel or 2-in. schedule 40 PVC pipe (approximate length, 6 to 8 in.) was placed in each cage, and enough 50% Holtfreter solution^1,11 (5 to 6 L) was added to cover the rodent tunnel to allow the axolotls to swim freely into and hide in the tunnel. Tap water was treated (NovAqua Plus and AmQuel Plus, Kordon, Hayward, CA); 50% Holtfreter solution was made by mixing the appropriate amounts of salts (1.75 g NaCl, 0.050 g CaCl₂, 0.025 g KCl, and 0.100 g NaHCO₃ per 1 L of treated tap water) in a clean container (for example, commercial trash can washed by using custom cage-wash settings to ensure removal of all detergents) with treated tap water, allowing the solution to age at least 24 h to allow conditioning of the tap water and chlorine to outgas (container was kept partially uncovered for approximately 24 h to allow chlorine outgassing). Before its use, the quality of each batch of 50% Holtfreter solution was checked (EasyStrips 6-in-1 Aquarium Test Strips and Ammonia Test Strips, Tetra, Blacksburg, VA). The animals were housed in a windowless room to prevent exposure to natural light, which facilitates algae growth; instead a 12:12-h artificial light:dark cycle (using standard fluorescent lighting) was used at an ambient temperature of 15 to 19 °C (59 to 66 °F).

Animals were free-choice fed a diet of sinking Soft-Moist Salmon Diet (Rangen, Buhl, ID) on Mondays, Wednesdays, and Fridays, allowing as long as 2 h for the animals to feed, given that adult axolotls exhibit low levels of activity when left undisturbed. Immediately after feeding, animals were transferred to clean plastic rodent cages; tunnels were rinsed and replaced weekly. All axolotls were checked daily for overall health, with inspection of the condition of gills and dorsal fin for signs of stress and for the presence of feces to ensure appropriate gastrointestinal function. All animals were acclimated to laboratory conditions for at least 5 d before any experimental procedures were initiated. All animal use was approved by the IACUC of Wayne State University, which is an AALAC-accredited institution.

Study design.

A sample size of 6 animals per group (control compared with treatment groups) was determined by assuming α = 0.05 and β = 0.05 and estimating a large effect size (σ = 0.25 for laboratory-bred animals and assuming clear behavioral differences with and without analgesia).⁵ In some cases, animals were used in multiple experiments; when animals were used in multiple experiments, a minimal 1-wk washout between treatments was permitted to allow the animals’ behavior to stabilize before proceeding. Our original set of experiments used 18 animals: 6 animals each in the control, buprenorphine, and butorphanol groups. After this first round of experiments, some animals showed signs of anorexia and constipation in the buprenorphine group and were euthanized because they met criterion/criteria in our IACUC protocol for animals that were clinically sick. We then ran additional experiments of 12 animals: 6 animals in control and high-dose butorphanol. Any animals that were still healthy in the first group were used in this second round of experiments.

Approximately 24 h prior to administration of the control or study compounds, each group of 6 axolotls was tested for baseline quantitative pain assessments and behavioral parameters. The next day, the study began, using the same personnel (2 for behavioral testing and one for quantitative testing) as for baseline assessment.

The 2 quantitative pain techniques—commercially available von Frey fibers and a modified AAT—were evaluated in naïve axolotls to determine which method produced more consistent responses. Each technique (vF or AAT) was evaluated by using a different group of 6 animals. The animals’ response to noxious mechanical and chemical stimulation was measured by using a vF aesthesiometer or various concentrations of acetic acid, respectively, to establish a baseline response. The quantitative method that produced the most repeatable and reproducible results was used exclusively in subsequent experiments.

To keep the operators blind to treatment, the study supervisor uniformly labeled 2 syringes for each animal, with each syringe marked with animal identification only. The study supervisor then aspirated equal volumes for control (amphibian saline as 0.63% NaCl), study drug for injection, or study drug for immersion bath. Although each animal was assigned to one treatment only, each animal received an injection of drug or control and mixing of drug or control into the home tank. This practice prevented bias from the operators looking for trends in animal behavior.

Quantitative pain assessment.

Quantitative pain assessments used in other laboratory animals were adapted for use in axolotls. Specific details regarding mechanical and chemical testing are found in subsequent sections. The application of noxious stimuli, both for quantitative assessment validation and analgesic efficacy experiments as measured by qualitative assessments, was performed as outlined in the schedule of experiments (Figure 1)

Figure 1.

Study schedule for each dose, with the study repeated twice (experiments A and B).

Mechanical stimulation using vF filaments.

For these tests, axolotls were kept in their home polypropylene static mouse cage. By using manual vF aesthesiometers (Touch Test Sensory Evaluator, Stoelting, Wood Dale, IL), increasing force was applied at the site of evaluation (Figure 2 A, lateral to dorsal fin, in line with hindlimb) until nociceptive behavior (for example, movement of limbs, contraction of dorsal trunk muscles [musculus dorsalis truncis], or full escape) was observed, at which point the applied force was recorded. Care was taken such that the flexible fiber was placed perpendicular to the site of evaluation before the force was applied. In addition, the manual aesthesiometers were brought toward the animal from a caudal direction to prevent it from visually detecting the approaching stimulus.

Figure 2.

Experimental setup for quantitative pain assessments. (A) von Frey fibers brought to hindlimbs from caudal direction to prevent animal from seeing assessment tool. (B) Animal resting completely submerged in angled housing before acetic acid testing. (C) Housing placed flat to expose dorsal surface of axolotl for acetic acid testing.

vF probes are supplied in force increments that are not linear but are standard for commercially available manual vF aesthesiometers. To make appropriate comparisons about the repeatability of vF aesthesiometers, the evaluators were mapped to an ordinal scale that included 0. The smallest gauge filament (0.008 g of force) was defined as the scale's zero point, because it bent when it touched the water surface simply due to surface tension. Commercially available vF sets are sold with 15 standard evaluators, such that the ordinal scale spanned the range from 0 to 14. This range was used in the statistical analysis of data.

Chemical stimulation using AAT.

For these tests, each axolotl was placed in a small polypropylene static mouse cage (11.5 × 7.5 × 5.0 in.) with enough 50% Holtfreter solution to cover half of its body, leaving the dorsal surface above the waterline. Between tests, the polypropylene mouse cage was tilted to one side by using a block or wedge to increase the water level that the animal experienced and allowing it to be fully submerged, thereby reducing the stress of being only partially submerged during AAT (Figure 2 B).

AAT was performed according to previously published reports in frogs.³⁰ Glacial acetic acid was serially diluted to produce 15 dilutions evenly spaced along a logarithmic scale (0.03 to 15.00 M). Testing started with placing a negative control, a single drop (20 μL) of 50% Holtfreter solution, to ensure that the animal did not respond simply due to the mechanical stimulation of the droplet. Testing proceeded by placing a single drop (20 μL) of the weakest concentration of acetic acid lateral to dorsal fin, in line with hindlimb (Figure 2 C). The animal was observed for a repeatable behavioral response (that is, wiping, turning, escape behavior). When a response was not observed within 5 s, the area was rinsed by using 50% Holtfreter solution. The next acetic acid concentration was then placed on the opposite side of the body. Testing proceeded in this manner, alternating sides to prevent over sensitization to a single area, until the nociceptive threshold—the lowest concentration to produce a behavioral response— was reached. Once a response was observed, a subsequent test was performed with the next highest concentration on the alternate side. If the alternate side also produced a positive response, the previous dose was recorded as the threshold. When no response occurred on this alternate side, testing continued as described. This approach ensured that the lowest concentration was captured appropriately, given potential differences between the left and right sides of the animal.

To make appropriate comparisons regarding the repeatability of AAT, the logarithmic concentrations were mapped to an ordinal scale that included 0. The weakest concentration of diluted acetic acid (0.03 M) was defined as the scale's zero point, because its use did not produce any response during pilot testing, controlling for mechanical stimuli with a negative control of 50% Holtfreter solution. With 16 concentrations of acetic acid mixed prior to the study, the ordinal scale for AAT ranged from 0 to 15.

Qualitative behavioral assessments.

The following tests were investigated to determine the behavioral experiences provoked by real or perceived tissue damage due to noninvasive stimuli. The schedule for qualitative assessments is found in Figure 1.

Feeding.

At least 1 wk of food consumption was measured to establish a baseline intake. Feeding behavior was measured by using a highly palatable treat: black worms. A few (3 to 5) black worms were placed in clear sight in front of the animal. Animals were given a maximum of 5 min to feed. A positive response occurred when the animal ate the worms, rather than simply snapping in the general direction of the worms.

Cageside assessments.

Operators blinded to treatment assessed the animals at cageside at least twice daily at predetermined time points. A single blinded operator consistently performed the cageside assessment methods to ensure repeatability of testing; the other operator provided an independent observation of behavior. Assessment methods included observing body posture and physical responses after gently tapping on cage, squirting 3 to 5 mL of 50% Holtfreter solution from a syringe at the base of the head with an inline jet aimed from a caudal-to-rostral direction, squirting a transverse jet of 50% Holtfreter solution aimed at the forelimb toward the dorsal fin, and squirting a transverse jet of 50% Holtfreter solution aimed at the hindlimb toward the dorsal fin; gently touching the animal at the midbody and tail with a gloved finger; and response to the placement of a novel object near the animal. The animals were scored by using a Likert-style system (0, no response; 1, minor response; 2, nominal response; and 3, major response) developed from observations during pilot studies. The scoring rubric is found in Figure 3

Figure 3.

Response key. Likert scores for behavioral testing based on pilot studies and observations.

Drugs.

The effects of different doses of butorphanol and buprenorphine on quantitative pain assessments and qualitative behavioral parameters to noxious stimuli were evaluated before and after the use of analgesia. The schedule of assessments is outlined in Figure 1. Six animals each were assigned to control (amphibian saline, 0.63% NaCl) and analgesic groups (buprenorphine and butorphanol). Buprenorphine (10 mg/mL; Penro Specialty Compounding, Colchester, VT) was administered as a single dose (50 mg/kg) through intracoelomic injection every 24 h for 48 h. The intracoelomic injection site was at the midpoint between the dorsal fin and right rear appendage. The location is caudoventral to the kidneys and cranial to the bladder, in the caudal aspect of the coelom; this site was away from major organs such as the liver, spleen and stomach. The syringe was aspirated prior to injection to confirm that the needle was not in the gastrointestinal tract, bladder, or a blood vessel. Pharmaceutical butorphanol (10 mg/mL; MWI Veterinary Supply, Boise, ID) was studied at 2 concentrations (0.50 and 0.75 mg of butorphanol per liter of tank water) and was directly administered into the 50% Holtfreter solution of the animal's cage every 24 h for 48 h. Prior studies (data not shown) using consecutive HPLC measurements confirmed that the 50% Holtfreter solution did not degrade or adversely interact with butorphanol over time.

Statistics.

All scoring methodologies were developed from observations during pilot studies. For qualitative behavioral tests, animals were scored by using a Likert-style system (that is, 0 to 3) for all measures except for feeding, which was simply dichotomous (yes or no). These observations are ordinal data, except for feeding, which yielded categorical data. For the quantitative tests, the logarithmic acetic acid concentrations were mapped to an interval scale that includes 0 (that is, 0 to 15). Although the recorded measurements are considered interval-type data, the results were not presumed to be normally distributed, and a nonparametric statistical approach was used. When the efficacy of a control or treatment effect was evaluated, the change (δ or Δ) in quantitative or qualitative response was referenced to the baseline measurement. Therefore, the statistic that is being evaluated is change in response. Findings from qualitative testing, including change in feeding, were included to assess whether analgesics changed behavior; results are not indicative of the efficacy of analgesia.

With the qualitative measurements recorded as ordinal data and recoding of a nonnormal (logarithmic) scale to an interval scale, the significance of treatment effects for qualitative and quantitative data at each time point was assessed through Kruskal–Wallis one-way ANOVA for unrelated samples. If the omnibus test showed significance, posthoc comparisons were run by using Mann–Whitney tests for qualitative and quantitative data on a succinct set of tests (compared with control treatment) to prevent inflation of type I error rates. At most, 2 post hoc Mann–Whitney tests were performed (control compared with treatment 1 and control compared with Treatment 2); therefore the critical level of significance in posthoc testing was corrected from α_critical = 0.05 to α_critical = 0.05 / 2 = 0.025. For feeding behavior, we had the independent nominal variable of treatment group (for example, control compared with various treatments) and dependent nominal variable of change in feeding behavior from baseline (for example, positive change, negative change, or no change). To investigate associations between 2 nominal (and categorical) variables, the Goodman and Kruskal λ test was used. All analyses were performed by using SPSS statistics software (versions 23 through 25, IBM North America, New York, NY).

Results

Comparison of quantitative methods.

Testing showed that the difference between the minimal and maximal scores was smaller for the AAT (range, 0 to 2; Figure 4, left) than for vF assessments (range, 2 to 8; Figure 4, left). From these simple descriptive statistics and the difficulty in consistently applying the manual vF evaluators on axolotls (for example, inability to keep probe perpendicular to axolotl body due to mucus coating and body curvature), AAT was used for all subsequent quantitative tests. The reproducibility of AAT was investigated in another set of 6 axolotls. Although the difference between the minimal and maximal scores for the AAT in this set of animals was larger (range, 1 to 3; Figure 4 right) than for the first series of tests, the range is still less than the variation of the vF evaluators.

Figure 4.

Repeatability of responses to serial assessments using von Frey (vF) aesthesiometers and acetic acid testing (AAT; right). Reproducibility of assessments using AAT in different animals (left). ●, vF trial 1; ▪, vF trial 2; ▲, vF trial 3; +, AAT trial 1; ×, AAT trial 2; and –, AAT trial 3.

Drug response.

Plots of quantitative and qualitative results (except feeding behavior) are shown in Figures 5 through 7. In the initial set of qualitative behavioral tests (except feeding behavior) assessing 50 mg/kg buprenorphine and 0.5 mg/L butorphanol (Figure 5), some behavioral tests at each time point showed significant (P < 0.025 and P < 0.005, respectively) differences when behavior was compared between treated and control animals. However, no consistent trend in overall behavioral responses according to analgesia usage over time was apparent for either study drug. The qualitative results for feeding behavior comparing 50 mg/kg buprenorphine and 0.5 mg/L butorphanol with controls were not statistically significant when the Goodman and Kruskal λ was used (no graph is produced in this test). In quantitative testing (Figure 7 A), no significant difference emerged between 50 mg/kg buprenorphine and 0.5 mg/L tank water of butorphanol compared with control. This first round of testing is designated experiment A in subsequent discussions.

Figure 5.

Results of qualitative assessments in experiment A. +, P < 0.025; ×, P < 0.005; ●, control; ▪, 0.5 mg/L butorphanol; and ▲, buprenorphine. (A) 1 h. (B) 6 h. (C) 25 h. (D) 30 h.

Figure 7.

Results of quantitative testing in (A) experiment A and (B) experiment B. No values were significantly different compared with controls. Legend: ● Control; ▪ Butorphanol (0.5 mg/L); □ Butorphanol (0.75 mg/L).; ▲ Buprenorphine.

During experiment A, adverse effects occurred in 2 of the 6 animals given 50 mg/kg buprenorphine. Fecal output ceased after the completion of the first round of experiments, meeting criterion for euthanasia. Necropsy of the 2 affected animals demonstrated dark discoloration of the gastrointestinal tissue (Figure 8 A through C) near the site of injection, with marked colonic distension and fecal impaction. Given the clear clinical observation that the fecal output was reduced, we decided to eliminate buprenorphine from further use in this study, and we performed another round of tests (experiment B) to compare a higher dose of butorphanol (0.75 mg/L) with the control.

Figure 8.

Adverse effects after buprenorphine injection. (A) Dissected and (B) in situ gross necropsy findings. Mobile phone camera (Huawei 6P; resolution, 96 × 96 dpi; 1× digital zoom). (C) Injection site schematic. (D) Affected and (E) unaffected gastrointestinal tract. TUNEL staining. (F) Affected and (G) unaffected gastrointestinal tract. Hematoxylin and eosin staining. Magnification, 40×.

After a minimal washout period of 1 wk, we repeated the qualitative and quantitative tests in experiment B. Qualitative behavioral tests (except feeding behavior) comparing 0.75 mg/L butorphanol with controls (Figure 6) revealed significant differences (P < 0.05, P < 0.025) between treated and control animals at 1, 6, and 25 h after the administration of butorphanol. However, no consistent trend in overall behavioral responses was clear. The qualitative results for feeding behavior during experiment B were not significantly different between treated and control animals according to the Goodman and Kruskal λ (no graph is produced in this test). In quantitative testing (Figure 7 B), high-dose butorphanol did not differ from control.

Figure 6.

Results of qualitative assessments in experiment B. *, P < 0.05; +, P < 0.025; ●, control; □, butorphanol. (A) 1 h. (B) 6 h. (C) 25 h. (D) 30 h.

Discussion

Previous studies in frogs (R. pipiens and X. laevis) have described mechanical (manual vF aesthesiometers), thermal (light energy), and chemical (acetic acid test [AAT]) methods to assess pain and analgesic efficacy in a quantitative manner;³⁰ however, all of these methods require modifications to produce repeatable results in axolotls. In the current study, we evaluated vF and AAT quantitative assessments to establish the repeatability and reproducibility of these 2 tests and to determine which is a better tool to use to quantify pain in axolotls.

In comparing AAT and vF as quantitative tools to assess pain in amphibians, our present findings are in direct contrast to the reproducibility and repeatability of these same methods when used in frogs. One study³⁰ found that vF had less interindividual variation than AAT when assessing pain thresholds in R. pipiens. This measurement variation may be due to differences between species or experimental methods. Concerning interspecies differences, the cutaneous mechanoreceptors of the lateral line system may be distributed differently between animals, or sensory-discriminative pathways may differ between frogs and axolotls. In addition, the experimental setups differ between the previous frog study³⁰ and the experiments we presented here. At least 2 d before testing, the frogs in the previous study³⁰ were transferred from their home environment to individual plastic cages containing 2 cm of water covering the cage bottom (reduced to 0.5 cm on the day of testing), clearly leaving the frog's hindlimb exposed for probing and testing. In contrast, to minimize extraneous nonnoxious stimuli and environmental cues that might prime a response, such as restraining the animals or allowing them to see the test devices, the aesthesiometers probed the animals or acetic acid was placed on them while the axolotls were in a quiescent and restful state in their home or test cage, submerged in 6 to 8 cm of water with their dorsal surface briefly exposed only for testing. In addition, to prevent a coached response, probes or pipettes were brought toward the animal from the caudal body aspect, preventing the animal from seeing the device's approach. Despite these precautions, unlike results in frogs,³⁰ vF testing showed more measurement variation than AAT.

Multiple factors contribute to the difficulty in translating vF testing to axolotls. Although care was taken such that the flexible fiber was placed perpendicular to the site of evaluation before the force is applied, the shape of the animal (curved body lateral to dorsal fin) and its natural mucous coating sometime make it difficult to consistently apply the evaluator perpendicular to the animal's skin. Conversely, using an adjustable-volume pipettor, a small-caliber pipette tip, and a very small volume of acetic acid allows for the more precise application of this noxious stimulus. Furthermore, after a simple practice (that is, cage tilting) was developed to keep the animal predominantly submerged when exposing the dorsal surface for only the few seconds needed to apply the acetic acid solution, we found that AAT is easier than vF to perform. Although noxious stimuli, such as those for AAT, are transmitted along unmyelinated C fibers and thinly myelinated Aδ fibers, whereas nonnoxious stimuli, such as mechanical sensation, activate rapidly conducting Aβ-like fibers,³⁰ thus perhaps accounting for variation between species, the human factors necessary to perform the tests and experimental setups must not be ignored when assessing a measurement tool's usefulness. The results of the 2 different quantitative approaches to measure pain in axolotls showed that an adapted AAT is more repeatable and reproducible in causing a noxious stimulus in this model animal compared with the nonnoxious stimuli using the manual vF fibers.

After adopting AAT for use in axolotls, we next explored the efficacy of available analgesics for this model animal. Because axolotls at our institution are used in studies of regenerative medicine, we chose opioids (buprenorphine and butorphanol) because they have been shown to be less immunomodulating than NSAID in other species^8,14,21 and because the early inflammatory response is important in mediating a robust healing response.¹⁹ The desire to allow inflammation to run its course in these injury models precludes the use of NSAID. Furthermore, the known immunosuppressive effects of morphine and fentanyl prohibited their inclusion into the study design. Therefore, we tested mixed partial opioid-receptor agonist–antagonist antinociceptives to determine whether they provide antinociception in these animals.

The behavioral parameters previously reported in newts¹³ were used in our study to evaluate the behavioral effects of analgesia administration in axolotls. In the previous study,¹³ the newts demonstrated significant differences in behavior (for example, feeding, movement after tapping on cage, body posture) between groups undergoing bilateral forelimb amputation without analgesic compared with the same surgical procedure after the administration of 2 different opioid analgesics (buprenorphine by intracoelomic injection and butorphanol by immersion). That study¹³ did have a surgical control, which received anesthesia alone, but an analgesic only group was not evaluated. In our study, we evaluated whether the analgesics alone had an effect on behavioral parameters in the absence of a surgical procedure. Our findings demonstrated that the analgesics alone—at the doses evaluated—do not have a statistically significant effect on behavioral parameters in axolotls. The author of the previous study¹² concluded that analgesics, as described, provided pain relief, in light of the results of qualitative behavioral testing in newts. However, the author¹³ further posited that quantitative testing could further validate the behavioral results. Therefore, quantitative data can extend the usefulness of subjective behavioral measurements.

Although studies caution the application of doses across unrelated species in the same class (for example, Amphibia),⁷ without guidance on doses and route of application, we used buprenorphine (intracoelomic injection) and butorphanol (immersion) in the same doses and routes as in eastern red-spotted newts.¹³ At the same doses in axolotls (buprenorphine: 50 mg/kg by intracoelomic injection; butorphanol: 0.50 mg/L directly into home cage water) compared with control animals, we found no significant difference in behavior or response to AAT between the control and treated groups. Given the study design, we repeated the qualitative and quantitative experiments at a 50% higher dose of butorphanol (0.75 mg/L directly into home cage water) but again found no significant difference in behavior or response to AAT between control and treated animals. Of note, many of the statistically significant differences between treated and control animals showed that the treated animals had a higher mean rank in response. This result is interpreted as treated animals being more sensitive to that behavior test than the control animals. For example, the inline jet showed a consistent significant difference to butorphanol compared with controls, but with higher mean ranks at all time points, thus suggesting that animals were more sensitive to the inline jet during assessments when treated with butorphanol. Unlike the previous study,¹³ the results presented here do not include an underlying model of pain. Although a dose–response to increasing concentrations of acetic acid was noted, there was no significant difference in the quantitative assessments comparing analgesic-treated animals with control animals. This result suggests that the choice of drug, drug dose, or route of delivery used in these experiments is suboptimal for axolotl analgesia when measured quantitatively.

Buprenorphine is commercially available as 0.3-mg/mL (Buprenex, Reckitt Benckiser, Slough, United Kingdom) and 1.8-mg/mL (Simbadol, Zoetis, Florham Park, NJ) formulations. Using these formulations and an average axolotl mass of 75 g, the volume of buprenorphine to achieve a dose of 50 mg/kg would be 12.5 mL for Buprenex or 2.1 mL for Simbadol—volumes that are quite large for such small animals. To reduce the required volume of injection into the animals under study, the buprenorphine that we used in the reported experiments was compounded (Penro Specialty Compounding) as a 10-mg/mL formulation. In addition, we used commercially available butorphanol (Torbugesic, Zoetis), because it was directly administered into the animals’ cage water. We hypothesize that the adverse events noted when using buprenorphine can be due to reduced gastrointestinal motility, a known side effect in μ-opioid pharmacology¹⁷ in other species, or the concentration of the buprenorphine dose (10 mg/mL) and its compounding formulation given through the intracoelomic route may be irritating to the gastrointestinal system. Although TUNEL staining is strong near the rectum of the 2 animals that were necropsied, suggesting DNA damage, hematoxylin and eosin staining does not indicate any pathology in this area (Figures 8 D through G). In addition, the TUNEL staining results could be due to increased gastrointestinal epithelial turnover. This finding could be caused by the primary effect of opioids causing reduced gastrointestinal motility or a local reaction to the intracoelomic injection of this buprenorphine formulation and concentration. In light of the adverse clinical outcome previously noted, buprenorphine was removed from further use.

Our hypothesis that buprenorphine and butorphanol will induce nociceptive changes but not behavioral changes at published doses used in other amphibian species is partially confirmed. From the observed behavioral responses and quantifiable measurements, the failure to find any significant differences, combined with noted adverse clinical outcomes, underscores the pitfalls in translating drugs and doses across species within the same class.⁷ The work in our current study provides guidelines for future studies of pain measurement and control in axolotls. The AAT is a well-established tool to quantitatively assess pain in laboratory animals, and it can effectively be adapted for use in axolotls. Future studies should explore additional doses of butorphanol through immersion routes and the efficacy of other opioids and nonNSAID-based analgesics as additional options for pain control in axolotls. In addition, if used in surgically invasive experiments like those commonly found in regenerative medicine studies, the effects of these drugs on the healing response in axolotls should be established. For example, the cardioprotective aspects of μ-opioid agonist signaling^3,9 are part of cardiac regeneration studies at our institution. Given that axolotls are a well-known model for epimorphic regeneration throughout its entire lifespan,²⁰ the lack of information on pain control relegates this species as an untapped resource for understanding mechanisms regulating tissue regeneration.