rats?

Pharmacological Treatment

A number of analgesic options are available. Refer to your institution's veterinary staff for treatment recommendations. Generally you should consider the use of local anesthetics, opioids, and non-steroidal anti-inflammatory drugs (NSAIDs). The opioids are controlled drugs and may be dispensed from an animal facility pharmacy. Some commonly used analgesics are: Opioid: Buprenorphine hydrochloride NSAIDs: Meloxicam Carprofen Flunixin Non-NSAID Acetaminophen derivatives

Detecting Clinical Signs of Pain and Distress

Signs of pain and distress in rodents are not easy to detect because of their: Small body size Tendency to conceal outward signs of pain and distress Habit of hiding or freezing when disturbed Nevertheless, signs of pain or distress can be detected in rodents by carefully observing subtle changes in behavior. The ability to properly assess pain and distress in rodents requires: Knowledge of normal rodent behavior and appearance Systematic approach to observing clinical signs in rodents The image shows rats with sleek hair coats moving around their cage. Normal feces are present in the bedding. The rats appear relatively normal from this top view. However, the rats in the far left upper corner and the rat in the lower left corner should be checked a little more carefully as they are hidden and perhaps may be head-pressing, which is a sign of distress.

Elicit a Response to Your Presence

2. Cage Wire Lid Off Lift the cage wire lid to elicit a response to your presence. This disturbance may prompt the animals to move about the cage. Examine the animals' behavior, gait, and hair coat. Normal rats and mice are inquisitive and explore their cage perimeter. The image below shows rats that appear alert, inquisitive, and well socialized. They have clean hair coats and are interested in who is on the other side of their cage.

examine the animal

3. Hand Restraint Examine (and treat) an individual mouse or rat by gently restraining the animal. You can move the animal to a separate examination box for detailed clinical inspection.

Systematically Monitoring for Pain and Distress

A best approach to minimizing animal pain or distress is to systematically monitor animals after a procedure or when illness is expected. How often the animals should be monitored depends on the: Severity of the animals' condition Expected rate of change in the animals' status Impact of the procedure on the animals At a minimum, all animals should be evaluated once daily. However, the nature of the procedure and condition of an animal may dictate that the animal be assessed multiple times a day. As mentioned on the previous screen, smaller mammals may experience physiologic changes such as chilling and starvation faster than larger animals. Therefore, rodents may require more frequent monitoring than larger animals. Some situations may require hourly or even continuous monitoring during critical periods in which rapid change in an animal's condition would be anticipated. This course offers a systematic daily approach for assessing clinical signs of rodent pain and distress. Some clinical signs may require assessment at a greater frequency to focus on parameters of particular relevance to the specific model and to provide the animals with appropriate intervention to minimize pain/distress.

performing a critical exam

A clinical exam should include: Observations of animal behavior, appearance, and posture to assess: Signs of pain or distress Clinical condition and homeostasis Measurements of clinical parameters, e.g., body temperature, clinical chemistries

common parameters

A common approach to assessing animal appearance and behavior is through observation of the following parameters. Tip: It is helpful to have blank forms to use as "score sheets" to enter and track each parameter assessed. (More on this at the end of this course.) Activity level hypoactivity (hunched, huddled, lethargic), hyperactivity, restlessness, lack of inquisitiveness Attitude arousal, depression, awareness of surroundings Behavior, spontaneous vocalization, self-trauma, isolation from cage mates. These observations are made without disturbing the animal Behavior, provoked vocalization, hiding, aggressiveness, minimal response. Note that these observations are made when the animal is disturbed or even prodded. Body condition emaciation, missing anatomy Food and fluid intake, elimination of feces and urine Fur and skin unkempt or greasy or dull fur; porphyrin staining around eyes and nostrils; cyanotic, pale, or congested mucous membranes or skin (ears, feet, tail); skin lesions; soiled anogenital area Eyes clarity/condition of lens, cornea; position of globe (e.g., sunken in orbit or protruding); condition of eyelids, encrustation Posture hunched back, tucked abdomen; prostrate; head tucked down Locomotion gait, ataxia, lameness, action of each limb, position of tail when ambulating Neurological tremor, convulsion, circling, paralysis, head tilt, coma Vital signs respiratory distress (open mouth breathing, pronounced chest movement) Other clinical parameters that are relevant to your study presence and status of tumors, infection, or surgical wounds

Scoring Systems for Clinical Exam Data

A defined scoring system of clinical parameters is a valuable aid for monitoring animal morbidity. The clinical parameters and scoring standards should be appropriate for the animal species and the disease model. This system can facilitate the decision to intervene to allay an animal's pain/distress, e.g., to administer treatment or euthanasia. If appropriate clinical parameters are not known for a particular disease model, you can perform a pilot study on a small number of animals to: Characterize the relevant clinical parameters; Define the time course of the disease and related critical events; Refine the endpoints; and Determine the timing and frequency for animal monitoring.

Practical Recommendations

A practical approach to using analgesics in rodents is to prepare a batch of doses for a population of animals over the period of a study. First calculate the total fluid volume required to dose all animals. Then make a solution of the analgesic at a concentration that will deliver the desired dose per aliquot administered. This approach can be used to medicate the animals with analgesic only or it can be used as a combination with hydration therapy. Remember to adjust the analgesic concentration according to whether the fluid aliquots will provide for hydration therapy or not.

abnormalities

Abnormal mice or rats may huddle in their cage, or they may fail to move around and explore their cage. In addition, rats may vocalize when approached. Inspect an animal's mode and speed of movement. Observe the tail position when the animal moves. Is the gait (how it walks) awkward? Observe how all limbs move while walking. Does the animal teeter or stumble? Is the animal's back hunched and abdomen tucked while walking? Is the tail held stiff and upright? Or does the tail drag? Tip: Observe a cage of normal animals for a comparison.

Physical Exam for Clinical Condition

After assessing the animals' appearance and behavior (discussed in the previous lesson), you should conduct a physical exam using methods that are appropriate to the species and experimental model. Performing a clinical exam on rodents is somewhat limited compared to larger animals due to the greater difficulty in venous access and the smaller sampling size of biological fluids. Nevertheless, specific methods and equipment for rodents allow a clinical exam to provide information on animal well being. In the image, the rats appear distressed. The investigators on this study believed that this was normal for day one postoperatively because the animals were moving. However, one can see head-pressing, no evidence of grooming, and porphyrin staining in these rats. One rat (bottom) does not move his tail in a normal way. A physical exam of this animal revealed low body temperature, hind limb weakness, anemia, pain, and weight loss.

nutritional support

Animals recovering from surgery develop a negative nitrogen balance as do human surgical patients. Young rodents are especially vulnerable to starvation because they lack long term fat and glycogen stores. Rodents typically have a reduced food (and water) intake 1-2 days after surgery. Low food intake may be more severe and more prolonged if animals are experiencing pain and distress (e.g., if pain alleviation is inadequate). Returning animals to a physiological plane that is as close to normal as possible is nearly always consistent with the scientific objectives of the study. Thus, the impact of surgery on the experimental model should be minimized. Nutritional support (as well as fluid and electrolyte therapy) is important for enhancing an animal's recovery after surgery. Nutritional support can also be important for nonsurgical studies in which morbidity and reduced food intake occurs. If you have included weight loss as a humane endpoint, you can actually generate false negative findings simply by failing to provide adequate nutritional support during the peak impact of a study. This is detrimental in research on interventions designed to help animals overcome sickness. Some examples of nutritional support include: Peanut butter Fresh fruit Baby rice cereal High protein or high fat drink (detailed on next page)

Tumors: Assessment

Develop an approach that evaluates both the general effects of cancer, e.g., inappetance, and the specific problems related to the type and placement of the tumor. Assessment of the clinical condition of a tumor-bearing rodent largely depends on characteristics of the tumor's biology, such as: Tumor growth rate Invasion Distension Ulceration Metastasis Production of cachectic factors The body systems most likely affected by the tumor should be identified and examined for clinical signs of illness. Therefore, the tumor model will determine the clinical signs to be monitored. Examples: Superficial tumors - ulceration, swellings Intracranial tumors - neurological signs Ascitic tumors - abdominal distension, dyspnea Although clinical signs may be anticipated, as related to the tumor biology and location, be mindful that unexpected signs may also occur.

Body Temperature: Assessment

Due to their large ratio of body surface area to mass and high metabolic rate, rodents lose body warmth at a faster rate than do larger animals. Conventional thermometers are not practical for use in rodents and can cause stress if used in unanesthetized rodents. In studies of toxicology, sepsis, diabetes, or whenever morbidity is expected to be high, investigators may consider the use of implantable microchips to track body temperature (as well as to identify an animal) without the need for animal manipulation. Microchips can be injected under the skin using conventional restraint or light inhalation anesthesia. Check with your institution's veterinary staff for information on purchasing a microchip system. (The noise of the microchip reader can frighten a rodent. Consider placing the chip in the animal's rump as opposed to the neck.) Body temperature is also a useful adjunct in the monitoring of humane endpoints in rodents because a reduction in temperature of sufficient magnitude can be a reliable predictor of death. Body temperature measurements may guide the decision of when to euthanize an animal, which will end or prevent unnecessary pain/distress and allow for the antemortem harvest of fresh body tissues for histopathologic or other analysis.

example

Even though this mouse is eating, he has a terribly rough hair coat, mottled appearance, squinted eyes, and is underweight and hunched.

summary

Good science requires good animal care. Animals that are in poor condition, discomfort, or pain are poor research subjects if such problems are extraneous to the objectives of the research. The impact on the animals' physiology can alter the outcome of the research data. In these cases, animal well-being supports the integrity of the research. In studies where animal morbidity is an expected outcome of the procedure (i.e., in a disease model when clinical symptoms are manifested), humane experimental endpoints should be established that do not conflict with the scientific objectives. The use of humane endpoints often benefits research by allowing the pre-mortem collection of biological samples. Using pre-established endpoints can avoid spontaneous death that results in loss of tissue due to post-mortem autolysis. The strategies described here for assessing animal well-being and pain or distress are guidelines that can assist you in developing animal assessment methods that are appropriate for your experimental procedures. Alleviation of pain and distress in animals is not achieved solely by the use of analgesics. Experimental procedures offer many opportunities for enhancing the animals' well-being by the refinement of procedures to reduce the severity of injury or stress and by the provision of supportive care. Many such refinements were described in this course. Using a system to assess animal well-being will help document the improvements in technical procedures and the benefits from supportive care.

Oral Analgesics

If injections are not necessary (i.e., for hydration therapy), you may consider offering an analgesic orally. A common approach is to add the analgesic (usually an opioid) to a gelatin treat, such as grape jelly, jello, or various commercial doughs or gels. (Rodents may prefer berry flavors and may avoid artificial citrus flavors.) Your veterinary staff will be familiar with these techniques. When administering a medicated treat, it is important to be sure that the intended animal (and not cagemates) eats the whole dose.

Important Considerations

If there is concern whether an analgesic may interfere with the experiment, conduct a pilot study to determine whether the analgesic may affect the study or not. Note: The pilot study must be approved by your IACUC first. An important consideration in the use of analgesics is to reassess the animal for pain as the analgesic effect wanes. Perform a clinical exam for signs of pain to determine if another dose is needed. For information on a record-keeping system, refer to the next lesson, Documentation of Post-Procedure Care.

overhydration

In conditions of diuresis and low specific gravity, urine may be collected for measuring urine specific gravity on a refractometer. Since rodents often urinate when picked up, you can be ready with a tube to collect a sample. You may also gently express the bladder. To locate the bladder, gently palpate the caudomedial abdomen while the animal is hand-restrained. The bladder will feel like a pea-sized structure. Be careful to avoid traumatizing the bladder! Excessive force will cause the bladder wall to hemorrhage, and blood will appear in the urine. A clinical refractometer is an inexpensive hand-held device that measures specific gravity and total protein. Rodent urine typically has a high specific gravity and so a small animal instrument should be used rather than one designed for humans. Although here, too, rodent urine specific gravity is likely to be above the scale. Therefore, the use of a refractometer will be more useful in conditions associated with diuresis and low specific gravity. Commercial urine dip sticks also measure urine specific gravity as well as urine creatinine, blood, leukocytes, protein, ketones, pH, and bilirubin.

quantifiable characteristics

In conducting a physical exam, use quantifiable characteristics whenever possible. These can be tracked over time and compared to a starting baseline or to normal, untreated animals. Such measurements are not only helpful for clinical assessments, but they can also be useful when compiling research data and writing manuscripts. Later in this course, simple record-keeping methods will be discussed to help utilize this information. You may evaluate: Behavior Body weight Surface lesions (wounds, masses) Hydration status Body temperature (telemetric methods) Blood parameters (Blood collection can be difficult/stressful in mice; may be used to confirm disease or failed treatment.)

example

In the image below, the mice are huddled. The mouse on the left has piloerection and a poor body condition. This animal has a generalized loss of muscle mass, making the spine prominent. One can palpate along a mouse's back and pelvic area to determine the extent of loss in the muscle mass.

investigator responsibility

Investigators are responsible for minimizing pain and distress in research animals by: Judicious use of anesthetics and analgesics Refinement of experimental techniques Implementation of best practices Implementation of humane endpoints Two critical components in the refinement of experimental techniques are: Monitoring animals for pain and distress, and Using interventions for reducing pain and distress. Federal animal welfare laws, regulations, and policies mandate the scientist's responsibility for the humane care and use of animals in research. A concise description of the requirements for the humane care and use of laboratory animals is given in the U.S. Government Principles for the Utilization and Care of Vertebrate Animals Used in Testing, Research, and Training.

Causes of Animal Pain and Distress

Investigators should be familiar with the causes of animal pain and distress. Pain and distress may be caused by spontaneous or experimentally-induced disease or injury. Many other factors may contribute to an animal's distress or discomfort, including extreme homeostatic challenges. Investigators should try to minimize pain/distress to an extent that is possible and compatible with experimental objectives. Wherever possible, pain/distress should be eliminated. Changes in the following parameters may cause or be associated with animal pain or distress: Body temperature Hypoxia Edema Blood electrolytes, e.g., hyperkalemia Dehydration Environment Caging Cage mates Lighting Humidity Noise Vibration Environmental temperature Note: Smaller mammals experience physiologic changes such as starvation (due to high metabolic rate) and chilling (due to large ratio of body surface area to mass) faster than larger animals.

fluid and electrolyte balance

Maintaining normal homeostasis is greatly dependent on osmotic pressure between tissue spaces. Fluid and/or electrolyte imbalance resulting in dehydration or edema may produce discomfort and add to pain and distress resulting from other causes. Also, animals in pain and distress are likely to have reduced fluid and food intake and so may develop dehydration secondarily. Rodents commonly become dehydrated due to experimental procedures that affect their water intake. Therefore, scientists and caregivers must be able to assess and control hydration. Performing the exam: Observe the animals' behavior. Rodents that are dehydrated may be sluggish. Assess the animals' appearance. Useful indices of hydration: Skin turgor: To assess skin turgor, tent the skin. Grasp, lift, and twist a fold of skin over an animal's back and watch the skin fall downward into normal position. Compare the response in a normal animal. In a dehydrated animal, the skin is less elastic and may remain tented longer and return more slowly to normal position. Hair coat Eye clarity Shape and position of the eye within the orbit Blood may be collected (in rats) for measuring total serum protein and electrolytes.

Supporting the Integrity of Research Data

Maximizing the humane care and use of laboratory animals and minimizing confounders of experimental variation are mutually complementary objectives of research animal management. Both support the integrity of the research data. Achieving humaneness in animal research depends upon the control, and whenever possible, the reduction of animal pain and distress. Minimizing pain and distress also reduces the impact of these extraneous factors on the research, i.e., as sources of non-experimental variation.

body weight assesment

Measuring body weight is a quick way to determine whether an animal is eating and drinking. Body weight changes are a sensitive indicator of rodent health, and a baseline weight measurement allows monitoring of the experiment's impact on the animal. Reduction in body weight may reflect starvation, dehydration, or a combination of both. Failure of young animals to gain weight is equivalent to a loss of body weight. Therefore, body weight changes should be interpreted in terms of both actual loss of weight and lack of expected growth. It is helpful to compare body weights of treated animals with those of normal controls. The body weights of mice and rats can vary dramatically depending on stock or strain. Refer to the weight curves on each strain or stock available from the animal vendor. In addition to measuring body weight, you should assess body condition. (This was briefly mentioned in Lesson 6, Appearance and Behavior: Assessment.) Rodents can be assessed for emaciation or cachexia (body wasting) by examining and palpating the lumbar spine and iliosacral areas. A scoring system can be applied to the progressive loss of fat and muscle mass to gauge the severity of emaciation. Approaches for nutritional supplementation will be described in this lesson. For treatment of hydration, refer to Lesson 9.

Body Condition Assessment and Scoring

Mice and rats can also be assessed for body condition which is one indicator of health status. For some situations, such as tumor studies, it may be more accurate to assess an animal's body condition, rather than its overall weight. Endpoints or other intervention criteria may be based on a score of body condition. Scales have been developed to standardize this process.

Treatment, continued

Normal maintenance volumes of Lactated Ringers Solution, or 0.9% saline, or glucose-saline can be injected in amounts of 1-2 ml/25 g mouse and about 5-10 ml/ 250 g rat per day. The subcutaneous administration of these volumes may begin prior to a study and should continue after the procedure, with multiple doses of smaller amounts, separated by several hours, throughout the period of expected morbidity. Therapeutic fluids should be warmed prior to injection because fluids administered at room temperature will chill the animal. Fluids can be loaded into syringes and kept warm in rodent support areas. Analgesic treatments may be combined with daily fluid administrations (for hydration therapy). For convenience in treating multiple animals, you can figure the total fluid volume needed for the study and add the appropriate amount of analgesic to a concentration that will deliver the desired dose in each aliquot administered. For more information on medications, refer to Lesson 12, Alleviation of Pain and Distress: Pharmacological Treatment.

Monitoring and Treatment

Previous lessons have discussed practical methods for conducting a clinical exam on rodents to assess morbidity, pain, and distress. This lesson addresses the documentation of exam findings and treatments. A records management system aids in documenting the status of your animals over time. And in cases when multiple staff take turns monitoring the animals, a record system facilitates good communication among all persons involved in the care and use of these animals. The image below shows cages of rats that recently underwent surgery. The cages have been affixed with a "watch" card so that it is easy to find these cages on the rack when a person enters the room. Each cage card also corresponds to a 5" x 8" medical record in a desk just outside the room. (There is a fresh orange for added nutrition; be careful not to leave fruit in a cage more than 12 hours as it will spoil.)

Treatment of Imbalance of Fluids and Electrolytes

Rodent discomfort and morbidity can be minimized with: Adequate administration of fluids Monitoring for clinical dehydration Providing supplemental fluids during experimental studies where there is predictable morbidity is often helpful for optimizing well-being in rodents. There are two common approaches for maintaining hydration status: Administering fluids proactively without assessing hydration status, based on the assumption that most animals in the study will have a similar degree of dehydration. Assessing hydration status and then formulating a fluid dosage to normalize hydration. This approach customizes the treatment for each animal and avoids over-hydration.

Scoring Systems: Guidelines

Some practical guidelines in developing a scoring system are to: Identify the clinical sign or signs that can be used to recognize the need for immediate euthanasia. Over successive studies, be mindful that the scoring system may need refinement. New and useful assessment methods may become available. Or, the clinical profile may change and require assessment by other parameters. Collect data on the numbers of animals that are euthanized vs. die unexpectedly. These data will help you refine the scoring system to minimize the number of animals dying from the experimental procedure without benefit of euthanasia. Publish your scoring system so that others may refine their methods based on your work. A scoring system can be incorporated into the health record. This composite record can track the animals' clinical profile and document the administration of treatments. The next page offers an example of a scoring system for clinical parameters.

Specific Examination Procedures

Specific physical exams may be added to the list on the preceding page to facilitate the detection and monitoring of illness, pain, and distress that result from your study procedures. For example: A neuromuscular exam can be conducted with simple techniques to measure hindlimb or forelimb strength and neurological deficits. Abdominal palpation (gentle) of the abdomen may detect pain due to peritonitis. (In rats, listen for vocalization or grunting or breath-holding by placing the animal close to your ear.) The following pages describe a systematic approach for a typical physical exam. Methods to treat abnormalities are included in this discussion.

High-Protein and High-Fat Diet

Stimulating appetite to increase food intake is helpful to promote a more rapid recovery in rodents as in other species. Something that tastes different and better than the normal everyday diet may be appealing to rats and mice and so may stimulate their appetite. Although some studies may have restricted nutrient requirements, the provision of a homemade or sterile commercially prepared supplement can be helpful to increase food intake and to maintain homeostatic controls such as caloric intake, electrolyte balance, and insulin/glucagon ratio. Commercial diets formulated specifically for rodents recovering from a surgical procedure may be used for balanced nutrition and fluid source, e.g., Surgical Transgel® (Charles River Laboratories). In addition, peanut butter has been used to tempt rodents to eat. This is an example of a high-protein and high-fat diet, which may coax an inappetant rodent to eat. It can be prepared as follows: 1 cup hot water 1 package raspberry gelatin 30 ml STAT VME High Calorie Liquid® (by PRN) 20 ml Pediasure® (by Abbott Laboratories) 2 scoops Designer ProteinTM (by Next Proteins International) Blend well. Pour into ice cube trays. Refrigerate. Feed the above diet at a rate of: 1/4 cube per rat per day 1 cube per cage of mice (5) per day

porphyrin staining

Stressed mice and rats commonly display "red tears" or porphyrin staining, which is a discharge from the Harderian gland in the orbit. Porphyrin staining may be seen on the nose, around the eyelids, or on the medial aspect of the forepaws which become stained through grooming of the face. Affected rodents may also fail to groom or they may have piloerection of the hair coat (giving a spiky appearance to the hair). The image below shows a mouse with porphyrin staining around the eye. Swelling around the eye and muzzle may indicate that these areas are irritated and that the animal has traumatized them by scratching.

Tumors: Pain and Distress

The growth of solid or ascitic tumors produces pain and distress in rodents just as in humans and other animals. Examples: Pain is associated with distension of overlaying tissues and ulceration of involved skin. Tumors that impinge on joints can impair body movement and locomotion and can restrict the animal's access to food and water. Growth of a tumor (any type) may cause the animal not to eat and lose body condition.

Alleviation of Pain and Distress

The detection and alleviation of pain or discomfort in rats and mice have been discussed in this course. The effective recognition of pain and distress should not rely on a single clinical observation but rather on a composite of signs and measurements that together reflect animal well-being in terms of pain or distress. In the image, a rat is shown 36 hours after a neurosurgical procedure. He has porphyrin staining or "red tears" around his eyes, nose, and medial forepaws. His incision appears swollen and painful. He has not been grooming. This animal should receive treatment to alleviate his pain and distress.

A Collaboration of Scientists and Veterinarians

The enhancement of the well-being of animals in experiments is often best accomplished through a collaboration of scientists and veterinarians. This team approach capitalizes on diverse perspectives for: assessing the animal response to the experimental procedures, and for arriving at a strategy of humane interventions during a study. Because the behavior of an animal model may be difficult to predict, ongoing efforts are often necessary to refine the supportive treatments used. A dynamic collaboration between scientists and veterinarians, involving continuing observations of the animals, will be most productive for developing humane interventions that are beneficial for the scientific outcome of an animal study. The image below shows normal Sprague Dawley rats on the day sutures were taken out of their head incisions. They appear comfortable. Rats normally sleep stretched out like this with their bodies in contact with one another. These animals have clean haircoats and appear well-groomed.

Appearance and Behavior: Observations

The first step in assessing clinical signs of pain and distress is a gross inspection of rats or mice for abnormalities in appearance and behavior in their home cage. This assessment takes only a few minutes for the practiced observer. Note that very young pups should be observed very carefully in order to avoid upsetting the mother and causing pup rejection. 1. From the Cage Exterior Routinely inspect the rodents through the top and sides of the cage. Get in the habit of removing the cage from the shelf and looking through all sides of the cage. Signs of distress may be missed in animals on lower or upper shelves because of low lighting or difficult access. Baby mice and rats can be inconspicuous within piles of bedding or nestboxes. The rats in the photo below are not having problems after surgery. They are sleeping the way one would expect and they appear comfortable. They are clean, have normal hair coats, good color (skin and mucosa), and normal vital signs.

example

The image below shows a nude mouse with an implanted tumor. The mouse has reached a humane endpoint in the experiment because of the tumor's size and because the tumor has become necrotic and ulcerated. This mouse was euthanized.

example

The image below shows a rat following a neurosurgical procedure. Although he is fairly clean and there is no staining around the eyes (porphyrin staining described later in the Course, he is displaying a hunched posture. The hunching of the back is a symptom of abdominal pain that is typically seen in quadrupeds. His head is held down and his coat is beginning to have a spiky appearance. This rat was euthanized and found to have an intestinal ileus from the use of chloral hydrate

Strategies

The results of the systematic clinical exam described in this course should be documented in a study record for animal health. (See the next lesson.) When animals are found to be in pain or distress, appropriate individuals should be contacted (i.e., veterinary staff and investigators). Determining the appropriate response involves a team approach with both scientific and veterinary input. A strategy to manage the adverse effects of the experimental procedures should be addressed in the protocol. Possible treatments may include the administration of analgesics, antibiotics, warmth, fluid therapy, nutritional supplements, etc.

Treatment of Hypothermia

There are many practical ways to provide temperature support to rodents, either individually or in cages. Select the following links for examples of practical approaches for conserving body warmth in rodents during an experimental procedure: Click links individually to show or hide information below each link, or use these buttons to open or close all links at once. Show All Hide All Insulated pouch or wrap Insulated pads Using warming pads Chemical warming pads (often too hot) Circulating water warming pads Electrical heating pads (use of human-grade pads is strongly discouraged) Warming racks Heat lamps (use is strongly discouraged) Monitoring area temperature For animals recovering from anesthesia, body temperature may remain low beyond the time the animals begin to ambulate. Therefore, it is best to keep them warm until their activity has returned to normal. In addition, if recovering animals are warmed within a cage, offer an area for escape from the heating device. This will allow recovering animals to leave the heated area for a cooler part of the cage if they become too hot.

Scoring Systems: Examples

Two example scoring systems are presented below. There are links to sample score sheets illustrating scoring standards. Example 1: For postoperative monitoring of rodents in surgical models, from Lab Animal 29:5, 40-45, May 2000. In this example, five parameters are used: Attitude Porphyrin staining Gait and posture Weight Food intake All parameters are rated on a 3-point scale of 0.0, 0.1, and 0.4. Score standards are defined for each parameter. A total score equal to or greater than 1.0 indicates the need for veterinary attention. Example 2: For clinical assessment of rodents with experimental autoimmune encephalomyelitis (EAE), from Animal Welfare Information Center Bulletin, 10(3-4):1-2, 20-22, 1999/2000. In this example, five grades of clinical signs of EAE are characterized and intervention actions are prescribed.

Components of a Record System

Typically, there are three components to a record system. Cage identification system: Cages to be monitored should be flagged to help an observer quickly locate the cages to be checked among all others in the animal room. Once the animals are no longer being treated, a different cage flag can be used to indicate the need for a later recheck of these cages. Colored stickers, hanging tabs, or index cards may be used. Consider a color-coded system for distinguishing the type of monitoring. Health record: A health record is used to document the clinical observations and physical exam findings. Records may be maintained for individual animals or a cage of animals. Consider using an index card, which can be kept in the cage card-holder throughout the monitoring period. A scoring system for clinical signs can be incorporated into the health record to provide both an efficient way to track the animals' clinical profiles over time and to compile numerical data useful for scientific purposes. For more information, refer to the next page, Scoring Systems for Clinical Exam Data. Record archive: It is helpful to archive animal health records so that they are accessible for routine use, for example in a procedure area adjacent to an animal room. The archive can be organized with a section of current cases. Staff who monitor the animals should routinely check this file before entering the room. This system allows for animals to be checked on a frequency that is appropriate to the condition - daily or weekly, for example.

Tumors: Endpoints

Unless otherwise approved by the IACUC, animals should be euthanized before they become moribund or die due to tumor load. Also, animals should be euthanized before the tumor mass becomes excessive, ulcerates, or impairs the animal's bodily functions or behavior. The criteria for endpoints in tumor development should be established in the animal protocol. These are generally a combination of: Tumor mass or burden (many institutions have specific tumor size guidelines) Body condition, e.g., cachexia (emaciation) Impairment of body functions, e.g., gait Ulceration Unless other arrangements have been made, it is usually the investigators' responsibility to euthanize animals that have reached their endpoints. Investigators should monitor their animals often enough so that endpoints are never passed. Allowing animals to go past their endpoints can be considered a protocol violation and may be acted upon by the IACUC if it occurs with any frequency.

Hypothermia

When under general anesthesia, rodents lose heat very rapidly. A mouse can lose 1 degree of body temperature per 5 minutes. A best practice is to use methods for conserving body heat during a procedure that will induce hypothermia, such as anesthesia and surgery. These methods are the provision of a heat source, thermal insulation, or a combination. Caution! Warming devices should provide gentle heat only (maximum of 40 °C or 104 °F). Having a high ratio of body surface area to mass, rodents on a heat source heat up as quickly as they lose body heat when chilled. They can readily overheat when high temperature heating systems are used, causing animal injury or death. The image below shows a rat with burns of the ears from over-utilization of a heat lamp. Burns can occur when a heat lamp is positioned too close to the animal. Heat lamps are generally discouraged because it is difficult for animals to escape the heat when they are too hot.