Using Waterless Alcohol-based Antiseptic for Skin Preparation and Active Thermal Support in Laboratory Rats

Rodents are frequently used for models that require surgical procedures. At our institution, laboratory rats are increasingly preferred for investigations of neurologic disorders, cardiovascular interventions, and assessment and treatment of addictive and depressive behaviors. For these types of studies, surgical preparations of the head and neck areas are necessary for catheterization and instrumentation. Based upon our former work in laboratory mice, we sought to improve rat surgery outcomes and confirm the efficacy of a waterless alcohol-based (WAB) antiseptic for skin disinfection prior to incision. In addition, we wanted to investigate whether active warming efforts improved perioperative body temperatures for rats to aid in return to consciousness. Prior to cranial surgical incision and placement in stereotactic equipment, rats were assessed after skin preparation with WAB and after thermal interventions, including prewarming cages for 30 min before anesthesia and delivery of warmed fluid (NaCl) supplementation. Core temperatures were recorded and aerobic culture swabs collected from surgical sites at multiple time points. As previously shown in mice, bacterial counts in rats were effectively diminished by WAB agents. Assessment of intraoperative body temperature trajectories did not identify appreciable differences between control rats and rats that were exposed to prewarming or warmed fluid supplementation or both. However, heavier male rats recovered more rapidly from isoflurane anesthesia than did lighter male and female rats. Although these thermal support measures did not significantly improve anesthetic recovery times in rats, animals warmed for 30 min trended toward a faster return to righting reflex after exposure to isoflurane. These findings confirm that WAB antiseptic is an acceptable option for skin preparation in rats and suggest that continued evaluation of thermal interventions remains of interest for improved outcomes in rat surgery.

High standards of animal welfare and quality of research outcomes are best promoted through concentrated training on surgical methods, extensive presurgical planning, and compliant execution of IACUC-approved surgery protocols.¹ The Guide for the Care and Use of Laboratory Animals and the American College of Laboratory Animal Medicine similarly advocate for aseptic techniques in survival surgeries performed on laboratory animal species.^1,26Typically, skin preparation for veterinary patients consists of hair removal and cleaning of the surgery site by multiple applications of relevant chemical agents and liquid rinses to minimize the chance for postoperative infections.^{2,3,17,25,29,33} Comparative studies in human, veterinary, and laboratory animal medicine have found several types and application protocols of skin cleaning agents to be equivalent in antimicrobial efficacy.^{10,12,16,22-25,29,41,46-48} Given that modern animal programs typically only permit use of laboratory rodents of defined health and pathogen status, repetitive application of agents to achieve skin antisepsis in rodents may be unnecessary and has been shown to contribute to hypothermia in laboratory mice during procedures.^6,14,21In human and veterinary medicine, waterless alcohol-based (WAB) antiseptics are commonly used for hand-scrubbing by the surgery team, but are not specifically used for preparation of the skin site prior to surgery.^{4,7,11,20,35,36,48-50} In the public realm, WAB solutions like ‘hand sanitizer’ gels, which contain ethanol or isopropyl alcohol or both, evaporate spontaneously without the need for rinsing. In addition, hand sanitizer gels promptly reduce the number of detectable aerobic microorganisms although they do not provide the sustained antimicrobial activity that is required for presurgical skin preparation of medical patients.^4,13 WAB antiseptics, which contain a combination of ethanol and chlorhexidine, are as effective as typical aqueous scrub agents in reducing bacteria on the skin of laboratory mice, yet require only 2 applications (no rinse needed) and mitigate heat loss during surgical skin preparation.¹³ The current study extended our previous work in mice by evaluating WAB antiseptic for surgical skin preparation in rats.^6,13,44Rats have long been a valuable model for behavioral neuroscience, cardiovascular, and addiction treatment investigations. Often, surgical preparations of the head and neck areas are necessary for catheterization and instrumentation. Building on literature indicating that prewarming of patients prior to anesthesia could promote thermoregulation and recovery,^5,8,28,42 we compared prewarming practices - the provision of warmed cage environments before anesthesia and in the early stages of the anesthetic procedure and provision of warmed intraperitoneal fluids - in study animals. To our knowledge, this is the first evaluation of WAB agents in rat surgery. The current report describes the first use of WAB antiseptic for cranial skin preparation, in combination with presurgical thermal interventions, in rats.     

Evaluation of In-cage Filter Paper as a Replacement for Sentinel Mice in the Detection of Murine Pathogens

Recent studies have evaluated alternatives to the use of live animals in colony health monitoring. Currently, an alternative method that is suitable for all rack types and that has been verified to detect the infectious agents most commonly excluded from mouse colonies is unavailable. We compared the use of filter paper placed on the inside floor of mouse cages to the traditional use of sentinel mice in the detection of several prevalent murine pathogens including mouse hepatitis virus (MHV), murine norovirus (MNV), minute virus of mice (MVM), mouse parvovirus (MPV), Theiler murine encephalomyelitis virus (TMEV), Helicobacter spp., Syphacia obvelata, and Aspiculuris tetraptera. Experimental groups comprised 7 cages containing either 2 pieces of filter paper on the cage floor or 2 ICR sentinel mice. Soiled bedding from pet-store mice was transferred to the experimental cages weekly for 8 wk. At 1 and 2 mo after bedding transfer, the filter papers were evaluated by PCR and sentinel mice were tested by serology and fecal PCR. Filter papers detected all pathogens as effectively (MHV, MNV, MPV, MVM, TMEV S. obvelata, and A. tetraptera) or more effectively (Helicobacter spp.) than sentinel mice at both time points. Filter papers more readily detected pathogens with a high copy number per RT-PCR analysis than a low copy number. Helicobacter spp. were not detected by sentinel mice at either time point. These results indicate that the use of filter paper placed on the interior floor of empty mouse cages and exposed to soiled bedding is efficient in detecting bacteria, endoparasites, and most of the common mouse viruses included in an animal health monitoring program.

Health monitoring programs are essential for animal facilities to maintain healthy animals, reduce zoonotic disease risk to personnel, and produce reliable research results.^25,27,35 Several methods exist to perform health monitoring in rodent colonies, including direct sampling of colony animals, sampling of sentinel animals, and environmental sampling. Direct sampling of colony animals eliminates the need to rely on the transmission of pathogens and the cost of purchasing additional sentinel animals. However, this technique uses valuable experimental animals and can lead to false-negative results, depending on the organism tested, prevalence of infection within the colony, and strain and age of the colony animals.^3,22,37 The use of contact sentinels has a higher probability of pathogen transmission compared with soiled-bedding sentinel animals or exhaust air testing, but using contact sentinels is impractical to implement in large colonies and may contribute to the spread of pathogens.^9,16,25,29 Soiled-bedding sentinels are useful for monitoring large colonies and can be used with any type of rack design but are limited to the detection of pathogens primarily transmitted fecoorally.^{2,9,15,29,32,33} The effectiveness of soiled-bedding sentinels in detecting pathogens can vary depending on the infection prevalence, frequency and amount of soiled bedding transferred, and strain and age of mice used as sentinels.^{3,4,9,15,29,46} The use of microisolation caging reduces the spread of pathogens between colony animals, resulting in a low infection prevalence, which can make detection of pathogens by sentinel animals more challenging. The number of contact or soiled-bedding sentinel animals required to detect an infection at low prevalence can become costly and may not be feasible for many institutions.^{7,9,12,28,42,44,45} In addition, sentinel testing relies on the infection of mice, purchase of mice, and euthanasia of live animals. If serologic detection is used, seroconversion can take 1 wk to several months, depending on the pathogen being tested, and its use is limited mainly to the detection of viruses and a few bacterial species.^17,25,27Compared with the use of live sentinel animals, environmental sampling using PCR of particles collected from prefilters at the exhaust air duct in ventilated cage systems can provide earlier and more reliable detection of ectoparasites, respiratory infectious agents, and Helicobacter spp.^{19-21,32,33,38,50} Although environmental sampling can eliminate the need to purchase and euthanize live sentinel animals, these techniques are not compatible with all ventilated rack designs and configurations.^2,11 In the last few years, 2 studies have evaluated environmental sampling techniques using PCR of filter media from the lids of cages maintained on IVC racks where exhaust air is filtered at the cage level.^11,14 One group used this cage-level method for reliable detection of pinworms and fur mites in cages housing 4 mice each.¹⁴ Another study found that the same technique was reliable for the detection of murine norovirus, Helicobacter spp., Pasteurella pneumotropica, Entamoeba muris, and Spironucleus muris in cages that did not contain mice.¹¹ The testing of filter paper from the lids of cages not occupied by mice allows for reduction in the animals used and is compatible with most cage types but requires shaking cages twice weekly, which may be time consuming. To our knowledge, no environmental sampling technique that can be performed at the cage level without sentinel animals or shaking of cages has been evaluated for efficiency regarding the most commonly excluded pathogens in laboratory mouse colonies.^18,27,29,31Our current study evaluated a new method of health monitoring that could replace the use of live sentinel animals and that can be implemented at the cage level, therefore making it compatible with any caging and rack system. We placed pieces of 2 types of filter paper on the interior floor of empty mouse cages, transferred soiled bedding, and compared the pathogen detection rate of these filters with traditional sentinel animals. The pieces of filter paper were mixed with soiled bedding weekly by using a new clean spoon. To confirm the use of our technique for health monitoring, we evaluated this technique in the presence of several pathogens commonly screened for in health monitoring of laboratory mouse colonies. We hypothesized that both types of filter papers, when placed on the interior floor of cages, would be as efficient as sentinel mice at detecting common mouse pathogens after soiled-bedding transfer.     

Effects of Oxygen Supplementation on Injectable and Inhalant Anesthesia in C57BL/6 Mice

Oxygen supplementation is rarely considered when anesthetizing laboratory mice, despite reports that mice become profoundly hypoxic under anesthesia. Little is known about the effects of hypoxia on anesthetic performance. This article focuses on the effects of oxygen supplementation on physiologic parameters and depth of anesthesia in male and female C57BL/6 mice. Anesthesia was performed via common injectable anesthetic protocols and with isoflurane. Mice anesthetized with injectable anesthesia received one of 3 drug protocols. Low-dose ketamine/xylazine (100/8 mg/kg) was chosen to provide immobilization of mice, suitable for imaging procedures. Medium-dose ketamine/xylazine/acepromazine (100/10/1 mg/kg) was chosen as a dose that has been recommended for surgical procedures. High-dose ketamine/xylazine/acepromazine (150/12/3 mg/kg) was chosen after pilot studies to provide a long duration of a deep plane of anesthesia. We also tested the effects of oxygen supplementation on the minimum alveolar concentration (MAC) of isoflurane in mice. Mice breathed supplemental 100% oxygen, room air, or medical air with 21% oxygen. Anesthetized mice that did not receive supplemental oxygen all became hypoxic, while hypoxia was prevented in mice that received oxygen. Oxygen supplementation did not affect the MAC of isoflurane. At the high injectable dose, all mice not receiving oxygen supplementation died while all mice receiving oxygen supplementation survived. At low and medium doses, supplemental oxygen reduced the duration of the surgical plane of anesthesia (low dose with oxygen: 22 ± 14 min; low dose without supplementation: 29 ± 18 min; medium dose with oxygen: 43 ± 18 min; medium dose without supplementation: 61 ± 27 min). These results suggest that mice anesthetized with injectable and inhalant anesthesia without supplemental oxygen are routinely hypoxic. This hypoxia prolongs the duration of anesthesia with injectable drug protocols and affects survival at high doses of injectable anesthetics. Because of variable responses to injectable anesthetics in mice, oxygen supplementation is recommended for all anesthetized mice.

Anesthesia is frequently required for mice used in biomedical research, but anecdotal communications suggest that mice receive significantly less anesthetic monitoring and supportive care than do other research species. Monitoring of anesthetized mice is often minimal due to lack of specialized monitoring equipment, and the fact that many rodent surgeries are performed by a single person who acts as both surgeon and anesthetist. Supportive care during anesthesia is limited by a lack of supporting experimental evidence. The lack of monitoring and supportive care may increase the mortality rate in anesthetized mice.Previous studies have shown that mice anesthetized with both inhalant and injectable anesthetics without supplemental oxygen become profoundly hypoxic.^{1,6,8,9,19,26,39,41} While mice in these studies appear to recover normally from anesthesia, little is known about the effects of hypoxia on physiologic parameters, anesthetic depth, and perioperative mortality. Respiratory complications, including hypoxia and hypoventilation, are second only to cardiovascular complications as a cause of perioperative mortality in veterinary species, and in humans, hypoxemia accounts for over 50% of deaths under anesthesia.⁴ To mitigate the risk of hypoxia under anesthesia, oxygen supplementation is commonly provided to anesthetized humans and animals, but is rarely provided to mice in research settings.^6,19All anesthetics affect respiratory function; ketamine and isoflurane are particularly known to cause respiratory depression in mice and rats by impairing the normal physiologic responses to hypoxemia and hypercapnia.^{9,12,20,23,28} The peripheral chemoreceptors, primarily in the carotid body, normally sense dropping arterial partial pressure of oxygen (PaO₂) while central chemoreceptors located in the medulla sense changes in pH and rising partial pressure of carbon dioxide (PaCO₂).^22,23,29,40 Both sets of chemoreceptors compensate by initiating increases in respiratory rate and tidal volume.^{23,28,31,34,40} Injectable and inhalant anesthetic agents depress the function of these chemoreceptors, preventing the increases in respiration that compensate for hypoxia and hypoventilation.^22,29Pulse oximetry is commonly used to monitor peripheral oxygen saturation and detect the presence of hypoxia. Pulse oximeters use the difference in light absorption of oxygenated hemoglobin and deoxygenated hemoglobin in arterial blood to provide an estimate of arterial oxygen content, abbreviated as SpO₂.¹⁷ An SpO₂ of less than 90% to 95% generally corresponds to a PaO₂ of less than 60 to 80 mm Hg, which is considered hypoxic in most species of mammals.^7,17 Because of the small size of mice, species-specific pulse oximetry equipment is necessary to obtain this measurement. Therefore, measurement of SpO₂ in anesthetized mice is not routinely performed, meaning that hypoxia under anesthesia generally goes unrecognized, and is likely more common than is appreciated by our field.The purpose of this study was to confirm that mice become hypoxic after receiving a ketamine/xylazine based anesthetic admixture or isoflurane, which are commonly used anesthetics in mice and to investigate the effects of oxygen supplementation on anesthetic depth, physiologic values, and anesthetic requirements in these mice.^9,35 We hypothesized that mice not receiving supplemental oxygen would be hypoxic, as indicated by lower SpO₂ while anesthetized, and that supplemental oxygen would correct this hypoxia. We also hypothesized that oxygen supplementation would increase the doses of injectable and inhalant anesthesia necessary to maintain mice at a surgical plane of anesthesia.     

Wellbeing of Mice Euthanized with Carbon Dioxide in Their Home Cage as Compared with an Induction Chamber

The AVMA Guidelines on Euthanasia state that, to decrease potential distress of animals, the home cage should be used for the euthanasia of mice. The current study evaluated this recommendation by comparing behavioral and physiologic changes in ICR and SJL mice that were euthanized by using a 30% volume per minute displacement rate of 100% CO₂ in either their home cage or an induction chamber. Blood samples were collected to assess blood glucose, serum corticosterone, and serum noradrenaline as markers of physiologic wellbeing. Behavioral assessment was performed (with emphasis on behaviors including rearing, jumping, sniffing at the gas inlet, and grooming) from the introduction of gas to the estimated time to loss of consciousness (i.e., the time period when the animal would be expected to experience pain or distress). Despite significant differences between mouse strains, no significant differences were detected in the physiologic or behavioral parameters assessed when comparing the home cage with the induction chamber. This finding suggests that— from the perspective of a mouse—either the home cage or an induction chamber can be used for induction of anesthesia with CO₂ during the euthanasia procedure.

General recommendations for the process of euthanizing rodents used in research include the maintenance of stable groups, reduction of transport prior to the euthanasia process, and selection of euthanasia methods that minimize pain and distress.^7,16 An additional recommendation is that the wellbeing of rodents is improved when the rodent is euthanized within its home cage.^7,16 This recommendation is based on a large body of literature that demonstrates transient increases in blood pressure, heart rate, and corticosterone after a cage change.^{8,17,23,24,26} Few studies have evaluated the effect of using an induction chamber or home cage during anesthesia or euthanasia with anesthetic overdose.^11,22This question is critical because of an apparent professional dichotomy in best practices for the improvement of animal wellbeing in euthanasia and anesthesia. Although the best practice recommendation for euthanasia is to use the home cage during induction with an inhaled anesthetic (generally CO₂ or halogenated anesthetics),^7,16 common inhalant anesthesia recommendations support the use of an induction chamber. Both CO₂ (especially when combined with O₂)⁹ and halogenated anesthetics are anesthetics, which can create a surgical level of anesthesia and, if used as an overdose, euthanasia,²⁸ therefore the wellbeing of the animal during induction is the same in both situations. In other words, because the sequence of experiences for anesthesia and euthanasia are initially similar, there will be pain or distress associated with the induction of anesthesia for any anesthetic used (such as CO₂¹⁰ or isoflurane⁴ or any injectable combinations of anesthetic agents),¹³ regardless of the intended outcome of anesthesia or euthanasia. If the home cage does improve animal wellbeing during euthanasia, then it follows that the home cage will also improve animal wellbeing during the induction of anesthesia. However, in both cases, practical reasons may underlie the use of an induction chamber for anesthesia. For example, there may be a need to anesthetize only select members of a cage cohort, or it may be beneficial to use a smaller induction chamber that can be sealed to minimize personnel exposure to the gases in use.To determine whether induction of anesthesia in the home cage promotes animal wellbeing during the euthanasia process as compared with the use of an induction chamber, I compared behavioral and physiologic changes of individually housed ICR and SJL mice that were anesthetized prior to euthanasia using a 30% volume displacement per minute rate of 100% CO₂ in either their home cage or an induction chamber. The physiologic responses assessed included rapid markers of activation of the HPA axis (e.g., noradrenaline) and more delayed markers of activation of this axis (e.g., corticosterone and blood glucose).^4,27 The behavioral parameters assessed included the frequency in which each mouse engaged in behaviors suggestive of anxiety (for example, jumping, digging) or exploration (for example, rearing, sniffing).^4,27 I hypothesized that mice anesthetized in their home cage would demonstrate fewer indices of distress than those anesthetized in the induction chamber.     

Evaluation of Treatment Options for Ulcerative Dermatitis in the P Rat

Rotational outbred adult rats, phenotypically selected to prefer drinking alcohol (“P” rats) frequently present with self-inflicted wounds and ulcerative dermatitis, similar to that seen in C57BL/6 mice. Historically, veterinary interventions used to address this clinical condition have included triple antibiotic ointment (TABO), Columbia wound powder (CPW), nail trims, or plastic tubes that allow affected animals to hide. More recent studies have suggested that nail trims are the most successful intervention in mice, but this has not been evaluated previously in rats. In this study, we evaluated nail trims in rats and also tested whether placing a pumice stone in the cage would reduce the need for nail trims to reduce self-inflicted wounds. Our hypothesis was that interacting with the pumice stone would dull/trim the rats’ nails without causing stress or illness and allow the wounds time to heal. We used 66 P rats that were assigned to 1 of 6 treatment groups (pumice stone, TABO, CWP, huts, nail trims, and an untreated control group) of 11 rats each. Rats were transferred to this study from a colony of experimentally naïve animals that had evidence of dermatitis. The wounds were photographed and measured for 12 wk at 2 wk intervals. At the end of the study, representative skin samples from the site of the wound were collected for histopathologic evaluation of inflammation. Our data showed no significant differences in the inflammation scores. The rats treated with nail trims healed significantly more often than did all of the other treatment groups. This suggests that nail trims are the most effective intervention for treating self-inflicted wounds in P rats.

Ulcerative dermatitis (UD) is a common disease of mice, with a prevalence rate ranging from 4% to 21%, especially in strains with a C57BL/6 background.^1,2,18 Clinical signs of UD in mice include moderate to severe pruritus-induced self-trauma that progresses from a small, superficial excoriation to deep ulcerations, tissue granulation, and crusts.^2,18 UD can progress to degloved skin sections in severe cases. The exact etiology of this UD is unknown, although multiple factors have been suggested, including sex, genetics (strain), age, diet, season and primary follicular dystrophy.^17,18,34,40 Some studies have indicated that the clinical etiology is behavioral in origin, with the lesions resulting from self-mutilating behavior in response to pruritis and skin inflammation.^6,13The alcohol-preferring (P) and alcohol-nonpreferring (NP) lines of rats were developed by phenotypic selection from a Wistar foundation stock.^4,21 They were developed to study ethanol drinking behaviors and the consequences of addiction. Indiana University has one of 2 primary breeding colonies of these rats in the world. Clinically, P rats in our breeding colonies, in which rats are kept for 8 to 12 mo, and are not handled on a daily basis, present with UD, including pruritis-induced self-trauma, with a prevalence of approximately 10%. This prevalence is not inconsistent with the prevalence of dermatitis reported in mice.^2,18 C57BL/6J mice are frequently used in addiction studies, suggesting a possible correlation between addiction and self-mutilation or dermatitis.^12,27,36 This potential relationship suggests that these clinical observations are not unexpected, given links between addiction and compulsive behaviors such as trichotillomania and skin picking.^11,29,30Because the etiology of UD is unknown, the identification of appropriate preventive measures is challenging. One suggested preventative measure is the addition of environmental enrichment to the animal''s cage, such as tubes or shacks to hide in or chew up, provision of additional nesting material, or food enrichment.^3,24,32 Several topical and injectable therapies have been used to treat UD, including vitamin E,^16,20,22 antioxidants,^8,15 and ibuprofen⁹ in drinking water. These therapies rely on antiinflammatory mediated analgesia, alleviating pruritus, antioxidative protection, immune suspension, antisepsis, and local anesthesia. The reported efficacy of these therapies varies widely and these treatments are usually ineffective.^26,34,40Nail trimming is another treatment for UD that has shown some degree of success in mice.^1,2 Nail trims may interfere with the maintenance and progression of UD lesions as a mechanical intervention that interrupts the itch-scratch cycle, even though this intervention may not address the underlying pathophysiologic factors contributing to the self-injurious behavior.³⁸ Studies in the literature have suggested that nail trims can be effective in as many as 90% of UD cases in mice.^1,2,26,37,40 However, this intervention has not previously been investigated in rats.This study aimed to identify a treatment or intervention that was reliably successful for amelioration of dermatitis in the P rat. Although nail trims appear to be the most successful intervention in mice, the larger size of rats makes restraint for nail trims more challenging and potentially stressful for both the rat and research personnel, particularly if the animal is not accustomed to regular restraint. We hypothesized that pumice stones placed in the cage would result in dulling of the nails, providing a similar outcome to that achieved with a nail trim while minimizing any potential stress associated with restraint. We speculated that walking over or manipulating the pumice stone would dull the rats’ nails, providing an alternative to nail trims. Based on current suggested treatments in mice, we compared the nail trim and pumice stone interventions with other commonly used interventions such as enrichment (Shepherd Shacks, Shepherd Specialty Papers, Watertown, TN) and topical medication used in mice at our facility (Columbia Antiseptic Wound Powder (SC Sturtevant, Bronxville, NY) and topical antibiotic ointment containing bacitracin zinc (USP 400 units), neomycin sulfate (USP 3.5 mg), and polymixin B sulfate (USP 5000 units) (Phoenix Global Supply Group, White Plains, NY). These treatments and interventions were compared with an untreated control group to assess their effectiveness. We hypothesized that the pumice stone would provide a refinement over nail trims by removing the need to restrain the rat, and that nail trims and the pumice stone would result in better outcomes, such as improved healing, when compared with the other treatments and interventions.     

Pharmacokinetics of Buprenorphine and Sustained-release Buprenorphine in Common Marmosets (Callithrix jacchus)

Buprenorphine is an essential component of analgesic protocols in common marmosets (Callithrix jacchus). The use of buprenorphine HCl (BUP) and sustained-release buprenorphine (BSR) formulations has become commonplace in this species, but the pharmacokinetics have not been evaluated. Healthy adult (age, 2.4 to 6.8 y; 6 female and 6 male) common marmosets were enrolled in this study to determine the pharmacokinetic parameters, plasma concentration–time curves, and any apparent adverse effects of these compounds. Equal numbers of each sex were randomly assigned to receive BUP (0.02 mg/kg IM) or BSR (0.2 mg/kg SC), resulting in peak plasma concentrations (mean ± 1 SD) of 15.2 ± 8.1 and 2.8 ± 1.2 ng/mL, terminal phase t_1/2 of 2.2 ± 1.0 and 32.6 ± 9.6 h, and AUC_0-last of 16.1 ± 3.7 and 98.6 ± 42.7 ng×h/mL. The plasma concentrations of buprenorphine exceeded the proposed minimal therapeutic threshold (0.1 ng/mL) at 5 and 15 min after BUP and BSR administration, showing that both compounds are rapid-acting, and remained above that threshold through the final time points of 8 and 72 h. Extrapolation of the terminal elimination phase of the mean concentration–time curves was used to develop the clinical dosing frequencies of 6 to 8 h for BUP and 3.0 to 3.5 d for BSR. Some adverse effects were observed after the administration of BUP to common marmosets in this study, thus mandating judicious use in clinical practice. BSR provided a safe, long-acting option for analgesia and therefore can be used to refine analgesic protocols in this species.

Renewed interest in common marmosets (Callithrix jacchus) as models for biomedical research is being driven by the fields of gene-editing technology, neuroscience, and infectious disease.^{7,19,23,28,32,34} Research study aims as well as clinical interventions necessitate appropriate pain management protocols for these animals. Current recommendations for analgesia in common marmosets are extrapolated from other species or are based on anecdotal evidence. Dosage, duration of action, and potential adverse effects of analgesics used in this species require evaluation to refine guidelines for their use in clinical practice.Buprenorphine is the most commonly used opioid analgesic in many NHP species, including common marmosets.^6,24,33 It is a partial µ-opioid receptor agonist used for the treatment of moderate to severe pain, when included as part of a multimodal pain management approach. Buprenorphine''s widespread use in laboratory animal medicine is attributed to a relatively long duration of action and favorable safety profile when compared with other available opioid agents. A considerable amount of data on the efficacy and recommended dosage of buprenorphine is available for various laboratory species, including mice, rats, rabbits, cats, dogs, and pigs.^{2,10,14,17,20,35,41}Formulations of sustained-release buprenorphine (BSR) have been developed and have become commercially available, providing a longer acting option than the standard buprenorphine HCl (BUP) formulation. Longer-acting compounds are preferable, because they have the potential to improve animal welfare by reducing handling and the number of injections per animal, reducing adverse effects associated with peak plasma buprenorphine concentrations, and avoiding repeated trough plasma buprenorphine concentrations, which may result in inadequate analgesia. The uses of both formulations of buprenorphine in several common laboratory animal species, including mice, rats, guinea pigs, dogs, cats, and macaques, have been described.^{5,9,10,16,30,39}The pharmacokinetics of BUP and BSR have already been described in both cynomolgus and rhesus macaques, the most commonly used NHP in biomedical research. The widely accepted dosage range for BUP in macaques is 0.01 to 0.03 mg/kg administered either intramuscularly or intravenously. Dosing recommendations provided in one study, using a plasma threshold of 0.1 ng/mL, suggest that for macaques, BUP at 0.01 mg/kg IM should be given every 6 to 8 h and at 0.03 mg/kg IM should be given every 12 h.³⁰ The same study showed that a single subcutaneous dose of BSR at 0.2 mg/kg can be given every 5 d. A similar study demonstrated that BUP given to rhesus macaques at 0.03 mg/kg either intravenously or intramuscularly maintains plasma concentrations above 0.1 ng/mL for 24 and 12 h, respectively.¹⁸BUP dosing recommendations for New World NHP, such as common marmosets, tend to be lower than those of Old World species, such as macaques. These recommendations are a result of the more profound adverse effects (for example, respiratory depression, apnea, and death) seen in common marmosets when higher doses of BUP are used or when BUP is used in combination with anesthetic agents, such as alfaxalone or isoflurane.^1,4 Guidelines for BUP dosing in common marmosets range from 0.005 to 0.02 mg/kg IM.^4,21,24The purpose of the current study was to investigate the plasma concentration of buprenorphine over time after the administration of BUP and BSR in common marmosets. Observations were made to determine whether adverse effects occurred after the administration of BUP and BSR. We hypothesized that BUP dosed at 0.02 mg/kg IM would remain above a plasma threshold of 0.1 ng/mL for 6 to 8 h and that BSR dosed at 0.2 mg/kg SC would remain above the same threshold for 72 h.     

Effects of Analgesics on Tumor Growth in Mouse Models of Prostate Cancer Bone Metastasis

Murine models of tumor development often require invasive procedures for tumor implantation, potentially causing pain or distress. However, analgesics are often withheld during implantation because of concerns that they may adversely affect tumor development. Previous studies examining the effects of analgesics on the development and metastasis of various tumor lines show that the effect of analgesics depends on the tumor line and analgesic used. A blanket statement that analgesics affect the general growth of tumors is not adequate scientific justification for withholding pain relief, and pilot studies or references are recommended for each specific tumor cell line and treatment combination. In this study, we evaluated the effects of 2 commonly used analgesics on tumor growth in 2 models of prostate cancer (PCa) bone metastasis. We hypothesized that a one-time injection of analgesics at the time of intratibial injection of tumor cells would not significantly impact tumor growth. Either C57BL/6 or SCID mice were injected subcutaneously with an analgesic (carprofen [5 mg/kg], or buprenorphine [0.1 mg/kg]) or vehicle (0.1 mL of saline) at the time of intratibial injection with a PCa cell line (RM1 or PC3, n = 10 to 11 per group). Tumor growth (measured by determination of tumor burden and the extent of bone involvement) and welfare (measured by nociception, locomotion, and weight) were monitored for 2 to 4 wk. Neither carprofen or buprenorphine administration consistently affected tumor growth or indices of animal welfare as compared with the saline control for either cell line. This study adds to the growing body of literature demonstrating that analgesia can be compatible with scientific objectives, and that a decision to withhold analgesics must be scientifically justified and evaluated on a model-specific basis.

Many murine models of bone metastasis rely on intratibial injections of tumor cells.¹¹ These injections are assumed to be painful for mice because any procedure considered painful in humans should be assumed to be painful in animals, regardless of their age or species.²⁸ In humans, bone biopsy, a similar procedure to intratibial injection, is considered painful, and both local and systemic analgesics are recommended during or after the procedure.⁶Researchers have an ethical obligation to relieve experimental pain whenever possible. Pain can alter behavioral and physiologic responses, leading to immunosuppression, through glucocorticoid and catecholamine release.³³ Therefore, appropriate pain relief is both an ethical and scientific mandate; in many countries, the use of appropriate analgesics is legally required.^3,4 Investigators must therefore provide scientific justification if their experimental design requires the withholding of analgesics during painful procedures.^14,37Some protocols for intratibial injection in mice recommend administering a systemic analgesic before tumor cell injection,^8,24 although studies have shown that commonly used analgesics such as NSAIDs (nonsteroidal antiinflammatory drugs) and opioids can affect tumor progression.^16,26,36 NSAIDs reduce the chronic inflammation that has been linked to carcinogenesis,³⁹ while opioids can modulate immune response or cellular pathways that control cancer cell survival and migratory behavior.² Thus, researchers may claim the need to withhold analgesia during intratibial injection because it may interfere with establishing the tumor model. However, unrelieved pain should also be considered a significant research confounder. In one study, unrelieved surgical pain, independent of the tumor implantation procedure, increased tumor growth and metastasis, while administration of buprenorphine ameliorated these effects.²¹ In human cancer patients, further research evaluating the potential adverse and protective effects of analgesics is necessary before making a recommendation either for or against the administration of such drugs.^23,26,30,40 Review articles examining the effect of analgesics on the development and metastasis of various tumors show inconsistent results that are highly dependent on the tumor model and analgesic characteristics such as route of administration, drug type, or dose.^2,33 The variability of how analgesia affects the general growth of tumors renders general statements inadequate as scientific justification for withholding analgesics; pilot studies or model-specific references are recommended for each specific scenario.³³In addition to inappropriately extrapolating the effects of analgesics from the literature, researchers may be reluctant to administer analgesics because they have not seen data regarding the use of analgesics in their specific model provided in the literature. A review article assessing the documentation of anesthesia and analgesia in research animals found that the majority of articles inadequately described the use of anesthetics and analgesics, failed to include an explicit statement that analgesics were withheld, or did not include discussion of how pain management or untreated pain would affect results.9 This gap in literature may lead researchers to believe that analgesics cannot be used in their model or field of study.In the current study, we evaluated the effects of 2 commonly used analgesics (carprofen and buprenorphine) on tumor growth in 2 mouse models of prostate cancer (PCa) bone metastasis. We hypothesized that a single injection of an analgesic at the time of intratibial injection of PCa cells would not significantly alter tumor growth.     

West Nile Virus Seroprevalence in an Outdoor Nonhuman Primate Breeding Colony in South Florida

West Nile virus (WNV) was first detected in Florida in July 2001, with 404 human cases reported to the Centers for Disease Control and Prevention as of February 2020. The subtropical climate of Florida is ideal for the mosquitoes that transmit WNV. We investigated the WNV seroprevalence in 3 NHP species housed outdoors at The Mannheimer Foundation in South Florida. From January to December 2016, 520 3 to 30 y old NHP were sampled at our 2 closed sites in Homestead and LaBelle: 200 rhesus macaques (Macaca mulatta), 212 cynomolgus macaques (Macaca fascicularis), and 108 hamadryas baboons (Papio hamadryas hamadryas). The presence of WNV IgG antibodies in these animals was determined by serum neutralization assays, which found a total seroprevalence of 14%. Seroprevalence was significantly higher in the baboons (29%) than the rhesus (11%) and cynomolgus (9%) macaques. The probability of seropositivity significantly increased with age, but sex and site did not significantly affect seroprevalence. The frequency of WNV seropositivity detected in these outdoor-housed NHP suggests that screening for WNV and other vector-borne diseases may be necessary prior to experimental use, particularly for infectious disease studies in which viremia or viral antibodies could confound results, and especially for populations housed outdoors in warm, wet climates. As no seropositive subjects demonstrated clinical signs of WNV and WNV exposure did not appear to significantly impact colony health, routine testing is likely unnecessary for most NHP colonies. However, WNV infection should still be considered as a differential diagnosis for any NHP presenting with nonspecific neurologic signs. Mosquito abatement plans and vigilant sanitation practices to further decrease mosquito and avian interaction with research NHP should also be considered.

West Nile virus (WNV) is a zoonotic arbovirus belonging to the family Flaviviridae and genus Flavivirus.^15,75 It is a member of the Japanese encephalitis virus antigenic complex that includes St Louis, Murray Valley, and Japanese encephalitis viruses.^5,41,75 The WNV genome is comprised of a single molecule of linear, positive-sense, single-stranded RNA.⁴¹ Its structural proteins include nucleocapsid protein C, transmembrane glycoprotein M and spike protein E, with spike protein E being the primary target for antibody-mediated neutralization.^4,41 WNV is a neurotropic virus, and 2 human pathogenic strains exist: lineages 1 and 2.^41,67 Lineage 1 strains are the primary cause of viral encephalitis in humans and horses in the United States, while lineage 2 strains have caused human epidemics in Eastern and Southern Europe and continue to affect horses in Africa.^5,16,23,41The first documented case of WNV occurred in a Ugandan woman in 1937.^24,59 Viral strains from the Eastern Hemisphere reached New York in 1999, resulting in an outbreak that spread rapidly throughout the United States in subsequent years.^15,28 Presently, WNV is endemic in the contiguous United States, with over 51,000 reported human cases and over 2,300 reported human deaths as of January 2020.^10,12 Transmission most commonly occurs by a mosquito vector, with Culex spp. being the most common.^57,65 Wild birds are the primary reservoir and amplifier host, while mammals such as humans, NHP, and horses are considered dead-end hosts because the viral load in mammals is usually insufficient to propagate the virus.^57,70 As of 2016, WNV has been detected in over 300 species of birds and 66 species of mosquitoes in the United States.^11,13 WNV largely causes asymptomatic infection (approximately 80% of cases), but may manifest as febrile illness (approximately 20% of cases), which may progress to retinopathy, encephalitis, meningitis, and death in immunocompromised individuals (less than 1%).^17,43,49Two surveys in outdoor NHP colonies detected WNV seropositivity in 39% of 726 rhesus macaques (Macaca mulatta), 20% of 563 pig-tailed macaques (Macaca nemestrina), and 51% of 403 baboons (Papio spp.) at the Tulane National Primate Research Center (TNPRC; Covington, LA) in 2002,⁵¹ and 6.6% of 45 sooty mangabeys (Cercocebus atys) and 0% of 45 rhesus macaques at the Yerkes National Primate Research Center (YNPRC; Lawrenceville, GA) from 2004 to 2006.¹⁷ Natural infection of WNV was reported in one 25-y-old outdoor-housed barbary macaque (Macaca sylvanus) exhibiting acute neurologic signs,⁴⁹ while experimentally infected rhesus macaques, common marmosets (Callithrix jacchus), and olive baboons (Papio hamadryas anubis) did not develop neurologic symptoms.^70,72,74South Florida is a peninsula bordered by the Gulf of Mexico, Straits of Florida, and Atlantic Ocean³ with a year-round subtropical climate characterized by heavy rainfall, severe thunderstorms, elevated temperatures, and intense humidity.^33,35,46 These weather conditions are ideal for significant mosquito activity and thus, potential exposure to vector-borne diseases such as WNV.⁷³ Florida is also home to approximately 80 species of native and invasive mosquitoes.¹⁸ Mosquito surveillance from August 2016 to November 2018 in Miami-Dade County, FL found that 5 species accounted for 85% of the mosquitoes collected: 38% Culex nigripalpus, 23% Aedes taeniorhynchus, 13% Culex quinquefasciatus, 5% Aedes aegypti, and 4% Anopheles crucians.⁷³ All of these species are confirmed WNV vectors.Because of the marked differences in WNV seroprevalence reported among NHP species from different geographic locations, the typical absence of clinical signs, and the permissive climate and mosquito population in South Florida, the goal of this study was to determine the seroprevalence of WNV in the outdoor NHP colony at The Mannheimer Foundation. The objectives were to compare differences in: 1) 3 NHP species: rhesus macaques (Macaca mulatta), cynomolgus macaques (Macaca fascicularis), and hamadryas baboons (Papio hamadryas hamadryas); 2) age; 3) sex, and 4) the 2 Mannheimer Foundation sites, located in Homestead and LaBelle, FL. Based on previous studies,^17,51 we hypothesized that the total WNV seroprevalence would be at least 15%, be significantly higher in baboons compared to macaques, and increase with age, but not significantly differ by sex or site.     

Assessment of an Electronic Mechanical Sensory Threshold Testing Device (RatMet) in Wistar rats (Rattus norvegicus)

von Frey (vF) monofilaments are used to quantify mechanical hypersensitivity and nociception in rodents; however, this method of testing has been criticized due to inconsistencies in testing methods, filament properties, and nonlinearity. This study compared withdrawal thresholds measured by using vF monofilaments with those of a novel mechanical threshold testing device currently in development (RatMet) in a carrageenan inflammatory model in 9- to 11-wk-old male Wistar rats. Rats were randomly assigned to assessment of mechanical hypersensitivity after intraplantar carrageenan injection by using either vF monofilaments (n = 10) or the RatMet device equipped with 3 sizes of probe tips (0.9 mm [RM0.9], n = 15; 0.5 mm [RM0.5], n = 11; and 0.09 mm [RM0.09], n = 11). All RatMet probe sizes and vF monofilaments identified a reduction in withdrawal threshold after treatment. Systematic differences in threshold were identified between vF and both RM0.9 and RM0.5 groups; RM0.09 did not differ from vF. Withdrawal thresholds showed linear relationships with probe diameter, square root of probe diameter, and area of the RatMet probes. In contrast, exponential relationships were observed with the vF monofilaments. Furthermore, none of the RatMet probe results differed in accuracy when comparing a single test with the averages of 3 or 5 tests per time point. Overall, the RatMet device measurements have construct validity even when the number of testing replicates is low. These data indicate that the RatMet device produces data comparable with those from vF monofilaments, with the potential for a shortened testing period without a decrease in accuracy.

Correctly quantifying pain and nociception are core components of pain and analgesia research. Mechanical sensory threshold testing is a widely accepted and commonly used method to evaluate hypersensitivity. This testing method continues to play a role within the evolving armamentarium of pain and nociception assessment tools.¹⁸ Developing valid, reliable, and practical mechanical sensory threshold devices are crucial for nociception studies and analgesia development.¹⁵ vF monofilaments (correctly termed Semmes–Weinstein filaments) remain the ‘gold standard’ for quantifying mechanical nociception in animal pain models despite numerous limitations.^3,4,11von Frey monofilaments are used to quantify mechanical sensory thresholds by identifying the force required to stimulate a withdrawal response, most commonly measured on cutaneous surfaces such as the plantar surface of the hind paw or the tail of rodents.¹¹ These monofilaments are commonly used because they are convenient, simple to use, have good intrarater consistency, and are well-established in the literature as an acceptable mechanical hypersensitivity assay.^3,8Despite their popularity, vF monofilaments have drawbacks, including nonuniform surface area when applied,^3,4 augmented hypersensitivity of the animal due to a training effect or tissue damage,^5,6,11 estimated withdrawal thresholds, inconsistency in testing methodology,⁴ sensitivity to operator hand tremor, and interrater inconsistency.^2,3 Multiple attempts have been made to standardize the application protocol of vF monofilaments to mitigate these limitations.² The 50% withdrawal threshold technique⁵ is often cited. However, variations in this technique, including the time between applications, speed of filament application, range of filaments used, and pattern of application, are frequently applied and inconsistently reported in the literature.^2,3 A standardized protocol would address some shortcomings of the vF monofilaments, but the inherent properties of the standard monofilament kit pose important limitations. The buckling nature of vF monofilaments, combined with variation in tissue characteristics, is likely to affect the applied force, and uneven force distribution may confound the actual withdrawal threshold.^3,4,17In an attempt to mitigate some of the limitations of traditional mechanical threshold testing with vF monofilaments, electronic vF devices have been developed. These devices typically use a single, nonbending probe. The probe is applied to the testing site (for example, plantar surface of the hind paw), and the force is steadily increased until a withdrawal response occurs. As a result, electronic vF devices provide a continuous force measurement, shorter testing time, and a reduced number of probe applications.⁹Our study evaluated a novel electronic vF device (RatMet, Topcat Metrology). This device uses a single, slightly flexible probe (plastic material, polypropylene or nylon) that does not buckle, and generates a force–time graph that compares the actual force curve to the recommended force application rate. Importantly, the RatMet device obtains an accurate, measured withdrawal threshold, unlike the discrete data and calculated estimate of vF monofilaments.^3,4 These properties have the potential advantages of reducing the effect of hand tremor, presenting a constant probe area to the tissue surface, and providing real-time user feedback on probe application. A standardized, continuous-force application may result in greater consistency within and between experimenters, greater accuracy in measuring withdrawal thresholds, and less likelihood of developing of exaggerated hypersensitivity.^3,5,6The objectives of this study were 1) to establish whether the RatMet device has construct validity by comparing the withdrawal thresholds and fundamental probe characteristics from 3 sizes of RatMet probe tips with those of vF monofilaments in Wistar rats and 2) to determine if reducing the number of replicate applications of the RatMet device alters the withdrawal threshold data. The hypotheses were: 1) RatMet would detect changes in mechanical withdrawal thresholds after treatment of rats with carrageenan; 2) reducing the diameter of RatMet probe tips would lower the mechanical withdrawal thresholds; and 3) reducing the number of replicate applications of RatMet would not change the withdrawal thresholds.     

A Survey of Laboratory Animal Veterinarians Regarding Mouse Welfare in Biomedical Research

The quality of research animal welfare is undeniably linked to the quality of scientific results generated from the animals. Although mice are the most commonly used mammalian species in biomedical research, little information is available about what factors should be considered to promote future progress. To address this issue, the Animal Welfare Committee of the American Society of Laboratory Animal Practitioners (ASLAP) surveyed laboratory animal veterinarians to obtain their opinions about the welfare of mice and to consider the roles of 5 factors that significantly affect animal welfare in biomedical research: husbandry, clinical care, experimental use, regulatory oversight, and training. The survey revealed that 95% of veterinarians scored mouse welfare as acceptable to excellent, although areas for improvement remain. These areas include: 1) training of researchers performing experimental procedures; 2) the frequency of monitoring mice likely to experience pain and distress due to experimental manipulation; 3) inclusion of the institutional veterinary staff in the monitoring of mice likely to experience pain and distress; 4) continued improvement in the environmental enrichment provided to mice; 5) the ability of the IACUC to ensure that instances of noncompliance are fully addressed in order to prevent reoccurrence both within laboratories and among other research groups at the institution; and 6) reliance on non-veterinarians to perform examinations, diagnose disease, and prescribe the treatment of sick or injured mice.

In biomedical research, the quality of research animal welfare has an undeniable, internationally recognized link to the quality of the scientific results generated from their use.^9,35,45 Animal welfare is also imperative for the continued public acceptance of the use of animals in biomedical research. Animal welfare standards are mandated by the federal government^3,4,37,39 with the policies, guidelines, and assurance standards setting the minimal levels for appropriate animal care.^25,38 Laboratory professionals working with animals recognize the importance of the physical and mental wellbeing of the animals in their care.^12,16 While the provision of adequate animal welfare may sound very straightforward, decisions about acceptable standards of animal welfare can be challenging for an institution to enact consistently. This may be due to differences in interpretation of what constitutes adequate welfare, and the fact that many biomedical experiments require the scientifically justified, intentional compromise of certain aspects of the animal''s health or wellbeing to accomplish experimental goals. The situation is further complicated by standards for the care and regulatory oversight of mice and rats in research. The Animal Welfare Act (AWA) or AWA Regulations^3,4 provide no oversight for mice of the genus Mus and rats of the genus Rattus that are bred for use in biomedical research. Further, most institutions use a large number of rodents, making individualized medicine and record-keeping difficult.The factors that determine the care and welfare of mice can be grouped into 5 categories; husbandry, clinical care, experimental use, regulatory oversight, and training. Each of these categories must be considered because they are often interrelated, and failure to attend to any of these individual categories may compromise rodent welfare.     

Comparison of Direct and Indirect Methods of Measuring Arterial Blood Pressure in Healthy,Anesthetized African Green Monkeys (Chlorocebus aethiops)

Quantitative blood pressure measurement is a critical parameter for assessing cardiovascular health, monitoring physiologic status under anesthesia, and making clinical decisions. The placement of an arterial catheter is the most accurate way to measure blood pressure, but is invasive and perhaps uncomfortable for the patient, requires sedation or anesthesia, and is technically challenging for staff. Noninvasive devices and methods, including oscillometry, high-definition oscillometry, and Doppler flow can be used to measure blood pressure. However, the American College of Veterinary Internal Medicine guidelines state that blood pressure should be measured using devices that have been validated in the species of interest and under the circumstances in which the measurement is being made. The alternatives listed above have not been validated in this manner. The objective of this study was to compare indirect ultrasonic Doppler flow detection and oscillometric blood pressure measures with direct arterial measurement in 8 healthy, anesthetized African green monkeys using the methods of Bland-Altman to assess agreement. Our results indicated that neither Doppler flow nor oscillometry consistently estimates direct blood pressure measurements in anesthetized African green monkeys. In 6 female monkeys, which weighed less than the 2 male subjects, the indirect Doppler measurement more closely predicted direct mean arterial pressure, indicating Doppler flow may be useful for monitoring mean arterial pressure in small patients.

Quantitative blood pressure (BP) measurement is vital to managing anesthesia and to assessing patients and their response to therapy in veterinary and human medicine.¹⁷ Maintaining adequate BP is essential for tissue perfusion, and a mean arterial pressure (MAP) less than 60 mm Hg in most mammalian species can lead to hypoperfusion, resulting in renal failure and shock.¹⁵ Careful maintenance of BP under anesthesia is therefore vital, particularly for procedures that require long surgical time.²⁴ Measurement of direct arterial pressure through catheterization is the gold standard in both human and veterinary medicine.²⁸ However, this technique is technically challenging, uncomfortable for the patient, and unsuitable for many clinical and research situations.¹² Many alternative, indirect methodologies have been used in the clinical setting, but questions about their accuracy have been raised.^{2,8,12,17,24,32,36}Two common indirect methodologies currently used in the veterinary clinical setting are oscillometry and ultrasonic Doppler.¹⁵ Oscillometric devices estimate BP by measuring the amplitudes of arterial wall oscillations during compression.³⁵ Ultrasonic Doppler estimates arterial pressure by detecting changes in blood flow during external cuff compression of a peripheral artery.¹⁹ The accuracy of these techniques, relative to the gold standard, is important to verify. The American College of Veterinary Internal Medicine (ACVIM) guidelines state that BP should be measured in devices that have been validated in the species of interest under the circumstances in which the patient is being tested.¹ Indirect methodologies have been tested against the gold standard in cats, dogs, goats, sheep, baboons, marmosets, cynomolgus and rhesus macaques, and human patients.^{2,10,11,17,25,29,30,32-34,36,37} To date, the author could find only one study from 1985 that assessed indirect compared with direct BP methodologies in African green monkeys (AGM, Chlorocebus aethiops).²⁷ The study compared infrasonic and Doppler flow methodologies to direct arterial catheterization. Infrasonic devices use a transducer to detect low-frequency (20 to 30 Hz) sound waves associated with the arterial wall. This differs from classic oscillometry, in which the BP cuff senses pressure fluctuations caused by vessel wall oscillations in the presence of pulsatile blood flow.²³ In addition, the study was published prior to the publication of national standards for electronic or automated sphygmomanometers by the Association for the Advancement of Medical Instrumentation in 1987.³ Although the study found correlation between the methodologies, statistical analysis interpreted results with linear regression modeling.The current standard for evaluating agreement between 2 methods of the same medical measurement is the analysis proposed by Bland Altman.¹³ This method is superior to using a correlation coefficient as an indicator of agreement.^4,5 Each animal is compared with itself by calculating the difference between the animal''s BP measurements from 2 different measurement methods (direct and indirect BP), as well as calculating the average of those measurements. For example, if the direct BP for one animal indicated 120 mm Hg, and the indirect BP for the same animal displayed 100 mm Hg, the difference is 20 mm Hg. This difference is calculated for each animal and the overall mean difference is considered the bias of one measurement method, compared with the other. Graphs of each animal''s difference compared with the average of the 2 BP method measurements are also used to visually compare the 2 measurement methods.In this study, we compared indirect ultrasonic Doppler flow detection and classic oscillometric BP measures (hereafter referred to as simply as oscillometry) with direct measurement after catheterization of the left femoral artery with radiotelemetry in healthy, anesthetized AGMs. We evaluated the consistency and accuracy of each measurement to determine which method best predicts direct BP measurement and compared these values to ACVIM guidelines for validity of BP devices. Further, we used the methods of Bland-Altman to assess the level of agreement between the methodologies.AGMs are increasingly used in biomedical research to study behavior, AIDS, diabetes, genetics, infectious disease, neurobiology, and cell biology.¹⁶ Under most circumstances, AGMs require sedation or general anesthesia for routine clinical care, diagnostics, and research-related procedures.¹⁷ These anesthetic regimens include α2 agonists, opioids, dissociative agents, or inhalant gas anesthetics. These agents have been associated with moderate to severe hypotension in a variety of clinical settings, making BP monitoring a critical component of AGM care. However, validation of indirect as compared with direct BP measurement in this species is crucial to improving anesthetic decision-making and overall patient health. Per ACVIM guidelines, a BP system is validated if the following conditions are met: the mean difference of paired measurements for systolic and diastolic pressures treated separately is ±10 mm Hg or less with a standard deviation of 15 mm Hg or less; the correlation between paired measures for systolic and diastolic pressures treated separately is ≥ 0.9 across the range of measured values of BP; 50% of all measurements for systolic and diastolic pressures treated separately lie within 10 mm Hg of the reference method, and 80% of all measurements for systolic and diastolic pressures treated separately lie within 20 mm Hg of the reference method.⁶     

Effects of Cisapride,Buprenorphine, and Their Combination on Gastrointestinal Transit in New Zealand White Rabbits

Due to their effective analgesic properties, opioids are worthy of consideration for pain management in rabbits. However, this class of drugs causes undesirable effects including reduced gastrointestinal (GI) motility, reduced fecal output, and delays GI transit times and thus increases the risk of GI stasis. The risk of stasis discourages the use of opioids in rabbits, which could affect animal welfare. Gastroprokinetic agents such as cisapride are effective in promoting gastric emptying in many species, but whether this effect occurs in rabbits is unknown. This study assessed the efficacy of cisapride when administered as a single agent and in combination with buprenorphine in rabbits; efficacy was assessed by measuring GI transit times, fecal output, body weight, and food and water intake. Female New Zealand White rabbits (n = 10) were studied in a crossover, randomized design and received either vehicle and buprenorphine, cisapride and saline, cisapride and buprenorphine, or vehicle and saline (control) every 8 h for 2 d. Rabbits were anesthetized and administered radio-opaque, barium-filled spheres via orogastric tube. Feces was assessed via radiography for detection of the barium-spheres to determine GI transit time. GI transit time was significantly longer in buprenorphine groups than in control groups, regardless of the use of cisapride. Fecal output and food and water intake were lower for buprenorphine groups than control groups. Cisapride did not significantly alter GI transit, fecal output, or food and water intake. In addition, treatment group did not significantly affect body weight. In conclusion, buprenorphine treatment (0.03 mg/kg TID) prolonged GI transit time and reduced fecal output and food and water consumption in rabbits. Coadministration of buprenorphine and cisapride (0.5 mg/kg) did not ameliorate these effects, and the administration of cisapride at this dose did not appear to affect GI motility in female rabbits.

In the United States, more than 2 million rabbits are client-owned, many of which may undergo moderate to markedly painful procedures such as surgery that require alleviation of pain and distress.¹ Furthermore, according to the USDA''s 2018 annual animal usage report, approximately 133,000 rabbits were used for research, many of which were used in potentially painful procedures requiring analgesia.³ The opioid analgesic most commonly used in rabbits is a partial μ-opioid receptor agonist, buprenorphine.⁸ Buprenorphine prolonged gastrointestinal (GI) transit times and reduced fecal output and food and water intake in male rabbits.^7,18 These established side effects of opioids leave rabbits at risk of developing a potentially life-threatening condition known as GI stasis. GI stasis is characterized by decreased muscular contractions of the stomach and intestines, leading to blockage and the proliferation of harmful bacteria in the rabbit''s intestinal tract. Initial clinical signs include reduced appetite, small or no fecal pellets, and abdominal discomfort.⁴Although opioid-induced GI stasis is reported in other species, rabbits are more prone to GI stasis due to their unique digestive anatomy, and they can have severe clinical signs.^13,24 The rabbit GI system combines a simple stomach with hindgut cecal fermentation. The cecal digestive process of rabbits differs from that of other hindgut-fermenting animals in that, in rabbits, indigestible fiber is eliminated as rapidly as possible through constant digestion and peristalsis. Other fine particles and solutes are retained in the cecum as substrates for microbial fermentation. This rapid separation of fiber and energy concentrates is useful in view of rabbits’ small size and high metabolic rate.²⁴A rabbit can become prone to developing GI stasis when GI motility, appetite, and thirst are decreased due to stress, inappropriate diet, GI blockage, or other causes of hypomotility, such as the administration of opioids. Stasis can lead to proliferation of harmful GI bacteria and the development of potentially fatal intestinal blockages.^24,30 Current standard-of-care treatment for GI stasis in rabbits includes restoring appetite through forced feedings, correcting dehydration, and administering agents that promote GI motility.^16,24,30 However, the recommended dosages of gastroprokinetic agents such as cisapride in rabbits are extrapolated from other species. Little scientific evidence exists regarding whether these prokinetic drugs are effective in rabbits at published doses.Cisapride is a serotonin 5-hydroxytryptamine 4 receptor agonist with some 5-hydroxytryptamine 3 antagonist activity, which results in increased GI motility. Cisapride has broad prokinetic activity, increasing the motility of the colon, esophagus, stomach, and small intestine in humans, dogs, cats, and mice.^15,23,27,29 Cisapride and the antacid ranitidine have synergistic effects on increasing GI contractility in postmortem isolated rabbit intestine.^13,23 In addition, cisapride improved contractility in ex-vivo newborn and adult rabbit ileums.¹⁴ In mice, cisapride reversed morphine-induced delayed GI transit time.²⁷ Another option for treating GI opioid-induced side effects is a peripheral opioid receptor antagonist, such as methylnaltrexone. However, methylnaltrexone was found to be ineffective at reducing negative GI side effects associated with buprenorphine in rabbits.¹⁸ Given the broad prokinetic effects of cisapride and the different mechanism of action of cisapride as compared with peripheral opioid antagonists, veterinary clinicians may reasonably administer cisapride to ameliorate GI stasis in rabbits. Multiple veterinary formularies include the use of cisapride as a prokinetic agent in rabbits.^6,10,23The effects of buprenorphine on GI motility were assessed previously in male rabbits.¹⁸ One aim of the current study was to characterize the GI motility effects of buprenorphine in healthy, female New Zealand White (NZW) rabbits. Furthermore, given the lack of information regarding gastroprokinetic agents in rabbits and the potential positive analgesic use of opioids during invasive procedures, the second aim of our study was to evaluate the efficacy of cisapride in healthy, female NZW rabbits and its potential to ameliorate the negative GI side effects of buprenorphine. Demonstrating cisapride''s effectiveness in ameliorating buprenorphine-induced GI side effects could encourage veterinarians and researchers to use this drug to treat opioid-induced stasis in rabbits, thus directly benefitting animal welfare. To our knowledge, this study is the first to evaluate cisapride''s potential to treat opioid-induced GI stasis in rabbits.Previous methods used to measure GI transit times in rabbits require training to interpret data and the use of expensive techniques such as Doppler ultrasonography, computed tomography, MRI, or wireless motility capsules.^5,9,22 In our current study, GI transit times were measured by using an established method involving biologically inert, barium-filled, radiopaque spheres administered via orogastric tube and quantified through serial radiography of the feces of each animal. Little specialized training is required to administer the spheres and to interpret results. The efficacy of this technique for measuring GI transit time was previously validated in horses.¹⁷ This method was used previously to document the effects of buprenorphine and methylnaltrexone on GI transit time in rabbits.^17,18,26Based on published data, we hypothesized that buprenorphine at 0.03 mg/kg SC every 8 h would reduce GI motility in female rabbits as evidenced by increased GI transit time and decreased fecal output when compared with a saline control group.¹⁸ Furthermore, we hypothesized that buprenorphine would decrease food and water intake when compared with a saline control group.¹⁸ For our second aim of the study, we hypothesized that cisapride alone at 0.5 mg/kg every 8 h would increase GI motility as evidenced by more rapid GI transit time when compared with a saline control group. The buprenorphine and cisapride dosing regimens selected for this study are commonly cited in many veterinary drug formularies.^6,10,23,24 Finally, we hypothesized that coadministration of cisapride and buprenorphine would alleviate the opioid-associated side effects of GI stasis by increasing GI motility, such that we would find no significant differences in GI transit time, fecal output, and food and water intake when comparing treated and control rabbits.     

Injection Reactions after Administration of Sustained-release Meloxicam to BALB/cJ,C57BL/6J,and Crl:CD1(ICR) Mice

The sustained-release formulation of meloxicam (MSR) is a compounded NSAID that may provide pain relief for as long as 72 h after administration. MSR injection-site skin reactions have occurred in several species but have not previously been observed in mice. We investigated the development and progression of localized skin reactions after a single injection of MSR in Crl:CD1(ICR), C57BL/6J, and BALB/cJ mice. Each mouse received a subcutaneous injection of MSR (n = 60), standard-formulation meloxicam (MEL; n = 24) or saline (control; SC; n = 24) and was scored daily according to a 5-point system for erythema and mass characteristics. Mice were euthanized at either 7 or 14 d after injection and underwent postmortem analysis. MSR-treated mice had more erythematous and mass reactions than did MEL and SC mice. Mass lesions developed in 49 MSR mice (82%; 95% CI, 70% to 90%), 5 MEL animals (21%; 95% CI, 7% to 42%), and 1 SC mouse. MSR-treated BALB/cJ developed erythematous lesions less frequently than similarly treated Crl:CD1(ICR) or C57BL/6J. Lesions often were ventrolateral to the injection site. The median times to the appearance of mass and erythematous lesions were 2 d and 3 d, respectively. Histologically, the erythematous and mass reactions correlated with necrotizing to pyogranulomatous injection-site panniculitis. Inflammation severity scores at 7 and 14 d after injection were greater in the MSR-treated group than the other 2 groups. No strain- or sex-associated differences emerged except that inflammation severity scores at day 14 were higher in Crl:CD1(ICR) females than males. The character of the inflammatory response in MSR-treated mice did not differ between 7 and 14 d after injection, indicating that MSR-induced inflammation is slow to resolve. The ventral migration and delayed onset of MSR injection-site reactions could result in their being attributed to another cause or not being identified. Researchers and clinicians should be aware of the potential for slowly resolving injection-site reactions with MSR.Abbreviation: MEL, standard-formulation meloxicam; MSR, sustained-release meloxicam; SC, saline control

NSAID are commonly used for analgesia in laboratory animals.^4,10 Standard-formulation meloxicam (MEL) is an NSAID that can be provided to mice via several routes (for example, subcutaneous, oral) and is typically administered every 12 to 24 h,¹⁰ but recent studies have shown that, due to rapid clearance, mice require more frequent dosing. A formulation that prolonged the dosing interval could be particularly beneficial.¹⁰ According to the manufacturer, sustained-release meloxicam (MSR) is a compounded formulation that might provide as long as 72 h of analgesia after a single subcutaneous dose in rats and mice.¹⁸ MSR maintains higher plasma drug concentrations for longer durations than MEL in mice, although not for a full 72 h; thus MSR may be preferable when analgesia is warranted for extended periods.⁹ A sustained-release formulation would reduce the amount of handling stress, risk of postoperative injury, and labor needed in animals undergoing an analgesic regimen.^4,9,10Previously published reports focus on drug efficacy and have not assessed or characterized potential adverse reactions to MSR in rodents or other species, despite reactions after the administration of standard formations of meloxicam.^12,13 However, Hispaniolan parrots and cynomolgus macaques are reported to have adverse reactions at MSR injection sites.^1,7 A sustained-release formulation of the opioid buprenorphine has caused erythematous lesions in several species,³ including mice² and rats.⁶ Despite reports of injection-site reactions to sustained-release formulations, this possible complication is not mentioned in the data sheet or information sheet for MSR.¹⁸ Elements of the proprietary vehicle, described by the compounder as a biodegradable liquid polymer matrix,¹⁸ may stimulate the immune system or mechanically irritate or damage tissues. In addition, previous studies have been conducted within the prescribed 72-h drug efficacy range, such that potential injection-site reactions initially arising beyond 72 h may not be observed or linked to MSR injections.^1,2,7,9,13 Regardless of the exact etiology, adverse reactions pose a threat to the assurance of animal welfare and potentially could interfere with research outcomes, particularly when increased inflammation may alter study results.We hypothesized that adverse injection-site reactions could be influenced by strain or sex and that these reactions could vary histologically in time, acutely or chronically. To test these hypotheses, we designed this study to characterize injection site reactions after treatment with MSR by assessing the presence, frequency, and severity of MSR injection-site reactions in males and females of 3 commonly used inbred and outbred mouse (Mus musculus) strains (BALB/cJ, C57BL/6J, and Crl:CD1(ICR). These strains have unique genetic backgrounds, which could influence reaction propensity. The C57BL/6J strain was selected because it is a widely used mouse strain for models of disease and is known to be predisposed to skin sensitivities, such as developing ulcerative dermatitis.^4,8,14 Crl:CD1(ICR) are outbred animals and may have more robust or variable immune responses, owing to higher genetic diversity.The goals of the current research were to determine the extent to which MSR leads to localized tissue reactions at injection sites, determine the characteristics and progression of any reactions, and assess whether strain-associated differences occur. These findings may aid veterinarians and researchers in drug selection.     

Effects of Cling Film Draping Material on Body Temperature of Mice During Surgery

General anesthesia induces many systemic effects, including thermoregulatory impairment and subsequent perioperative hypothermia. Due to the animals’ small size, monitoring and maintaining body temperatures in laboratory rodents during anesthesia is important for successful surgical outcomes and prompt anesthetic recovery. Draping materials have the potential to aid in thermal support during surgical anesthesia. In this study, rectal and surface (infrared) temperatures were measured in C57BL/6 mice under isoflurane anesthesia every 5 min for the duration of a 35-min sham surgery. In addition to placement on a circulating water bath, mice (n = 6/group) were draped with commercial cling film (CF; Press''n Seal, Glad, Oakland, CA), a conventional paper drape (PD), or no drape (ND) during surgery. Results demonstrated that CF-draped animals had significantly higher rectal temperatures than nondraped animals. Furthermore, surface temperatures of CF-draped mice were considerably higher than those of both paper-draped and undraped animals. The data indicate that cling film is an effective material to help minimize hypothermia in mice and potentially in other laboratory rodents requiring general anesthesia.

Surgery and anesthesia introduce many challenges, especially in veterinary medicine, due to the diversity of species. One major challenge during general anesthesia involves changes in an animal''s thermoregulatory ability.^1,14 Body temperatures in mice and rats fall significantly during anesthesia if no thermal support is provided.^29,30 Hypothermia occurs due to drug-induced alterations to the thermoregulatory center, inadequate circulation, and a loss of body heat to the environment from evaporation, radiation, conduction, and convection.⁷ Mice are particularly susceptible to hypothermia, due to their large surface area per gram of body weight, which permits significant physiologic changes in response to fluctuations in the ambient temperature.³¹ Covering the animal''s body with towels, drapes, or blankets to reduce the area exposed to the environment can minimize heat loss.^6,7,13 Placing the animal on an insulated surface can limit conductive heat loss. In larger animals, warmed fluids can be given perioperatively, heated anesthetic gasses can be administered, and heated blankets and heat packs can be applied to body surfaces to provide exogenous heat.^1,7 Safer and more practical methods for rodents are circulating water heating blankets, thermal gel packs, and warming lamps, which are commonly used for thermal support during anesthesia.^5,14 Addressing all of these factors can contribute to maintaining normothermia during anesthesia.Risk of mortality is elevated during anesthesia and in the postoperative period, including in rodents.^1,13 Hypothermia induced by anesthesia can negatively affect rodents by altering vital parameters such as heart rate and blood pressure and delaying anesthetic recovery.^3,5,12,19 These risks require careful selection of an appropriate anesthetic protocol and careful monitoring of the patient throughout anesthesia until full recovery occurs. Strict anesthetic monitoring and the use of supplemental heat devices have been shown to reduce the likelihood of complications, improve overall postoperative recovery, and reduce mortality associated with surgical procedures.^1,7,15,16 However, due to these species’ small size, monitoring equipment must be specialized and is often costly. Cost-effective and practical alternative equipment and materials would facilitate monitoring and care of rodents.Various draping options are available for rodent surgery, and their use is vital for both sterile technique and heat retention. Traditionally, paper draping material has been a popular option, because it is relatively inexpensive and can be autoclaved together with surgical instruments.^15,16 Some institutions have adopted varying methods and types of draping, including no drape and paper draping. Commercial cling film (CF) has been used as draping due to its low cost, ease of use, and sterility straight out of the box.⁹ Our study team sought to evaluate the effects of draping material on intraoperative thermoregulation in mice by measuring rectal temperature (modified rectal probe) and surface temperature (infrared device) during a 35-min laparotomy procedure, with both temperature devices chosen for affordability and availability. We hypothesized that mice draped with CF would maintain a higher intraoperative body temperature under general anesthesia than would mice with traditional paper drapes or no drape.     

Comparison of Two Hair Removal Methods in Sprague–Dawley Rats (Rattus norvegicus)

Rats commonly undergo surgery for research purposes. However, the effects of different methods of hair removal on wound healing and surgical site infections (SSI) in rats has not been evaluated. The current study evaluated 2 hair removal methods, clipping with an electric clipper and using a depilatory agent, and their effect on wound healing and SSI. Swabs for bacterial culture were obtained on Day 0 just after hair removal, after aseptic skin preparation, and on Days 1 and 3 before conducting skin biopsies to assess bacterial load and recolonization. Full-thickness punch biopsies were taken for histopathologic evaluation on Days 0, 1, 3, 7, and 10. The surgical incisions were assigned an ASEPSIS score on Days 1 and 3. The data revealed that the bacterial load was significantly higher with the depilatory method as compared with the clipper method, but only on Day 1. The histopathologic evaluation found no significant difference in wound healing between the 2 methods. Although the ASEPSIS score was significantly higher for the clipping method than for the depilatory method on Day 1, both techniques were equivalent by Day 3. We conclude that both hair removal methods are safe and efficacious components of aseptic technique in rats.

Sprague–Dawley rats are often used in surgical procedures for biomedical research and training. The Guide for the Care and Use of Laboratory Animals²⁰ requires the use of aseptic technique when performing survival surgery on any species. One purpose of aseptic technique is to reduce or eliminate the bacterial load on the animal prior to the start of surgery to prevent the introduction of bacteria into the sterile surface below the skin.^2,30,42 Insufficient or inappropriate skin preparation may result in surgical site infections (SSI). SSI can delay or compromise wound healing.^1,34 Aseptic technique requires the preparation of the surgical site on the animal by removing the hair such that skin damage, abrasions or other dermal injuries are avoided, followed by cleaning the skin with topical antiseptic compound.^6,30Traditionally, hair is removed from surgical sites because it harbors bacteria and prevents thorough cleansing of the incision site. Hair removal also facilitates visibility of the surgery site and removes a potential foreign-body that may result in SSI.^10,22,43 The 3 most common hair removal methods are shaving with a razor, clipping the hair with an electric razor, and using a depilatory agent. In human patients, recommendations are that the hair not be removed unless visualization is needed or the hair would interfere with the surgical site or postsurgical bandaging.^5,43 If hair removal is necessary for humans, the recommendation is to use either clippers or a depilatory agent. Using a razor has been shown to traumatize the skin, resulting in higher rates of SSI.^22,28,35,36 Some research studies indicate that a depilatory agent is a better method, as it is efficient, atraumatic, and safe to use on or around wounds. However, it can cause a transitory lymphocytic reaction, and some individuals may have a sensitivity reaction.^1,17,22,24 In mice, hair removal with either clipping or depilatory agent resulted in acceptable healing.²⁵ In Wistar rats, the use of a depilatory agent did not affect the healing of a dorsal flap.³ Despite these findings, no studies have compared the effects of clipping and a depilatory agent on the prevalence of SSI and on wound healing in rats.Our facility''s standard practice is to remove hair with clippers using a no. 40 blade. However, this approach leaves a short stubble of approximately 1 mm.²² For surgical procedures that require a smooth skin, using a depilatory agent appears reasonable; however, little information is available on the effects of a depilatory agent on rat skin, SSI, and wound healing. We hypothesize that using a depilatory agent as a hair removal method in rats will reduce bacterial counts, dermal trauma, and SSI as compared with using clippers.     

Behavioral and Reproductive Effects of Environmental Enrichment and Pseudoloma neurophilia infection on Adult Zebrafish (Danio rerio)

Recent studies have shown beneficial effects of environmental enrichment (EE) for zebrafish, while infection of zebrafish with the common pathogen Pseudoloma neurophilia has negative effects. This study investigates the effects of P. neurophilia infection and EE in housing and breeding tanks on measures of behavior, growth, and reproduction. Zebrafish were socially housed and were either infected, P. neurophilia-infected (PNI) (n = 12 tanks), or SPF for P. neurophilia (SPF) (n = 24 tanks). Fish were housed with or without EE, which consisted of placing plastic plants in the tanks; sprigs from plants were placed in half of the breeding tanks for half of breedings, alternating breeding tanks without EE weekly. Behavioral testing included the Novel Tank Diving Test (NTT) and Light/Dark Preference Test (LDT) conducted prior to breeding. At the end of the study, biometric data were collected. Histopathology and molecular analysis for common diseases in fish confirmed that SPF fish remained SPF and that fish from all PNI tanks were infected. PNI fish produced significantly fewer eggs and had lower body weights and lengths than did SPF fish. Fish with EE had longer body lengths, than did fish without EE, and male fish had longer body lengths than female fish. The biometric results and reproductive measures show that SPF fish exhibited better growth and suggest that EE in housing tanks could improve fish growth. The behavioral test results were inconclusive regarding whether infection status or EE altered anxiety-like behavior. Our results support other recent studies showing negative effects of P. neurophilia infection on zebrafish.

The use of laboratory zebrafish (Danio rerio) has expanded dramatically in recent decades from an animal model for developmental genetics into one that is employed in a wide variety of research areas, including: neuroscience, animal behavior, cancer, tissue regeneration, immunology, infection, aging, and toxicology research.^9,20 Many of these research areas use adult zebrafish, which are more likely to be affected by spontaneous disease than are embryos and larvae.⁸ Coinciding with the expanding popularity of zebrafish in research is the prevalence of microsporidiosis, caused by the microsporidian parasite Pseudoloma neurophilia.^23,36 P. neurophilia is one of the most common pathogens detected in laboratory zebrafish,^23,27 causing both subclinical infections and progressive clinical disease in heavy infections.^7,11 Clinical signs of P. neurophilia infection include emaciation, spinal deformities, reduced growth, altered behavior, decreased fecundity, and increased mortality.^7,11,28,29 Zebrafish with clinical signs of P. neurophilia can often be visually identified by their low body condition score,⁶ scoliosis, and resultant swimming abnormalities and should be removed from experiments and breeding stock.P. neurophilia is an obligate intracellular parasite with a tropism for neural tissue. P. neurophilia is transmitted both vertically and horizontally.²⁶ Some evidence indicates that surface disinfection of embryos is not completely effective at preventing transmission of the parasite,^10,11 making its exclusion from laboratory animal facilities challenging. In addition, no treatment is currently available for microsporidiosis, which makes elimination of established infections difficult. However, rigorous pathogen screening protocols have helped establish lines of laboratory zebrafish that are SPF for P. neurophilia.¹⁴Naturally occurring clinical and subclinical infections in animals are well established to cause confounds and high variability that can lead to invalid or misinterpreted experiments and an accumulating body of experimental evidence has begun to characterize the adverse effects of P. neurophilia and other pathogens on zebrafish research.^{2,13,15,21,22,24,28,29} However, subclinical PNI zebrafish often go undetected, and thus may be enrolled in, and potentially compromise, biomedical research projects. In addition, uninfected zebrafish may acquire the infection during the course of an experiment, potentially creating undesired effects on research outcomes. In one study, PNI zebrafish were found to have a significantly reduced habituation to a startle response as compared with both exposed negative tankmates and unexposed control zebrafish.²⁹ In other studies, PNI zebrafish displayed more freezing behavior or altered shoaling behavior relative to uninfected control zebrafish.^21,28 These findings suggest that microsporidiosis could initiate a stressed or anxiety-like behavioral phenotype, which could confound behavioral research findings.²⁸ Previous reports have suggested that the effects of P. neurophilia infection may vary with sex and genetic background.^5,24 A retrospective study evaluating histopathologic specimen from the Zebrafish International Resource Center (ZIRC) found a 12.4% higher prevalence of P. neurophilia infected males in routine health cases and a 7.5% higher prevalence in males with clinical cases.⁵ Another study found that when zebrafish from the AB line that had an endemic P. neurophilia infection were subjected to stress or treated with cortisol, their mortality rate significantly increased over a 7 wk period, while no mortality was observed in the TL line of zebrafish.²⁴ However, experimentally-infected TL zebrafish weighed 27% less than control TL zebrafish, indicating that although mortality rate was not affected in the TL line, the growth rate was affected.²⁴ Larval zebrafish appear to be particularly susceptible to P. neurophilia infections, leading to the suggestion that infection could compromise developmental neurotoxicity testing and other development studies.¹⁵ Thus, the exclusion and elimination of P. neurophilia from laboratory zebrafish colonies would be beneficial to research.To date, methods for providing environmental enrichment (EE) are not well-established for many aquatic species,¹² and zebrafish may often not be routinely provided with EE. However, a growing number of studies demonstrate that EE influences zebrafish behavior in certain situations.^{8,16,18,19,32,33} For example, for zebrafish exposed to unpredictable chronic stress, access to EE for 21 or 28 d attenuated both behavioral and chemical responses to stress.¹⁹ While socially housing zebrafish is the default at most research facilities, zebrafish occasionally require single housing for experimental or other husbandry or clinically-related purposes. In cases where zebrafish are individually housed, the addition of an artificial plant as EE has been shown to decrease anxiety in behavioral measures, compared with single-housed fish in tanks without EE.⁸ In breeding tanks, zebrafish appear to prefer spawning in a grass-enriched environment, as evidenced by a higher number of viable embryos produced in breeding tanks containing EE.¹⁶ In a separate study, EE did not affect fry survivability.³³ The authors of that study concluded that further studies are warranted to identify optimal EE strategies that promote species-typical behaviors and reduce stress and anxiety-like behaviors in laboratory zebrafish.Multiple behavioral assays have been developed to assess behavior in zebrafish, including two tests of anxiety-like behaviors - the novel tank diving test (NTT) and the light:dark preference test (LDT). Based on geotaxis, the NTT uses the zebrafish''s instinctual escape response to seek protection in an unfamiliar environment by diving, freezing, and reducing exploration in the vertical water column space.³ As zebrafish gradually acclimate to a novel environment, increased exploration into the test tank''s top half normally occurs.³⁵ Anxiolytic drugs, such as fluoxetine and diazepam, increase the time zebrafish spend at the top of the NTT tank.³¹ In contrast, anxiogenic stimuli increase time spent at the bottom of the tank, immobility, and erratic movements.³¹ The LDT is based on the zebrafish''s overt scototaxis, or innate preference for dark rather than brightly lit environments.¹⁶ Zebrafish exposed to anxiogenic substances or stressors spend more time in dark areas of a test tank than do control zebrafish.¹⁶ While both the NTT and the LDT test for anxiety-like behaviors in zebrafish, one study evaluating the use of these 2 tests suggested that for a comprehensive assessment, the best approach is to use both NTT and LDT tests.¹Our goal in this study was to identify the effects of P. neurophilia infection and EE on anxiety-like behaviors and reproductive performance in adult AB line zebrafish. Juvenile PNI and SPF zebrafish were socially housed with or without EE in their tanks. After 5 wk, zebrafish were behaviorally tested in the NTT and LDT and then bred for 6 wk, alternating whether EE was provided in the breeding tank or not each week. At the end of the breeding experiment, all adult zebrafish were euthanized, measured, and weighed, and samples were submitted for analysis by histopathology and real-time PCR to confirm PNI or SPF status. We hypothesized that P. neurophilia infection would increase anxiety-like behaviors in NTT and LDT behavioral assays, would not be significantly impacted by EE, and would have a negative impact on fecundity. We hypothesized that no significant difference in anxiety-like behaviors would be present between fish that were group-housed with EE or without EE and that a significant difference in reproductive performance would be found between breeding tanks that contained EE and those that did not.     

Effects of Pair Housing on Patency of Jugular Catheters in Rats (Rattus norvegicus)

Chronic vascular access devices are widely used in a variety of species for repeated blood sampling or substance administration. Jugular catheters are commonly used for studying addiction-related behaviors in rats. Rats with catheters have historically been individually housed for the duration of the study to prevent cage mates from damaging the catheter. The 2 goals of this study were to determine 1) the effects of pair housing on catheter patency and 2) the effects of pair housing on catheter patency of rats in a study of opioid self-administration and cue-induced reinstatement of opioid-seeking behavior. The latter study also represented an opportunity for experimental refinement as it evaluated the temporary use of a barrier that allowed for pair-housed rats to be physically separated. Male Heterogeneous Stock (HS; n = 24) and Sprague–Dawley (SD; n = 121) rats were allocated to either single- or pair-housed condition. To assess the effect of social housing on catheter patency, rats (HS, n = 24; SD, n = 36) were monitored in their assigned housing condition for one month, with scheduled evaluation of catheter patency and structural damage. To examine the effect of social housing on catheter patency during a study of opioid self-administration and cue-induced reinstatement of opioid-seeking behavior, rats (SD, n = 85) were monitored in their assigned housing condition with similar routine patency evaluations. Catheter patency rates between single- and pair-housed rats were not statistically different in the first experiment, and pair-housed animals were successfully maintained on an infusion study in the second experiment. The use of a barrier between pair-housed rats after surgery allowed continued social contact with no observed adverse effects. These results suggest that, pair housing is a viable option for rats with chronic vascular implants, and may improve their wellbeing by allowing them to display species-typical social behaviors.

The widespread support of species-appropriate social housing in biomedical research has increased in the recent years and, as has the number of publications that can be found citing social housing as a key variable within their investigations.^13,28,29,31 Regulations state that appropriate social interactions are essential to the wellbeing of laboratory animals, and that single housing of social species should be the exception.¹² Further guidelines state that a disrupted or restricted social environment can negatively affect animals of all ages and may lead to maladaptive behaviors such as self-injury and stereotypies.¹¹The benefits and behavioral effects of social housing in rodents, and rats in particular, have been reported in previous literature and provide an important foundation for research into the feasibility of socially housing animals used in behavioral research.^2,5,27,32 Rats are a naturally social species,²⁷ and social housing in the laboratory does not impede critical behaviors such as feeding.⁵ Moreover, the beneficial effects of social housing include reduced anxiety-like behavior² and attenuated motivation to self-administer rewarding substances such as sucrose and cocaine.³²Some situations still require that rats be single-housed in the laboratory. Such situations include health concerns, social incompatibility, and specific research paradigms. Findings from previous literature indicate that single housing, if not properly accounted for, may significantly affect the interpretation of results.^{1,7,9,10,16,17,19,28} Single housing can be stressful¹⁷ and can alter gut microbiotia,⁷ both of which may induce anxiety-¹ and depressive-like behavior.¹⁶ These conditions may manifest as weight gain,²⁸ poor grooming, inactivity, learned helplessness²⁰ and social avoidance.¹⁹ Social isolation also reduces normal behavioral repertoires, such as ultrasonic vocalization,¹⁰ and increases consumption of rewarding substances such as sucrose and cocaine.⁹Although social isolation may introduce an experimental variable and adversely affect research outcomes, a limitation of pair housing in the research setting is concern that a cage mate may damage an implanted device. One example of this is the study of addiction-related behaviors using self-administration paradigms. In these cases, rats with indwelling jugular catheters self-administer controlled substances over an extended period of time. Rats participating in these studies are often individually housed in order to prevent damage to the catheter, which must remain patent for the duration of the experiment. Limited information is available about the incidence of cage mate-related catheter complications and the methods used to socially house rats with chronic vascular implants.The development and refinement of appropriate practices to socially house rats with chronic implants are important factors in optimizing animal wellbeing and successful research outcomes. The principles of replacement, reduction, and refinement (3Rs) were developed several decades ago to promote the humane use of animals in research.^22,26 With this in mind, we studied whether social housing was a feasible experimental refinement for rats participating in an opioid self-administration and reinstatement study. Rats used for this investigation were already part of our laboratory''s rat colony and not needed for experiments for meeting the laboratory''s primary research goals. The 2 goals of the current study were to determine the effects of social housing on catheter patency (Experiment 1) and on catheter patency of rats participating in an opioid self-administration and cue-induced reinstatement study (Experiment 2). Opportunities for refinement were identified throughout the study, one of which was an additional aim for Experiment 2. We examined the use of a ‘buddy barrier’ for pair-housed rats participating in the opioid self-administration study. For this study, we hypothesized that the long-term patency rates of catheters in individually housed and pair-housed rats would not be different.