Diseases of mice
Additional sources of information
The laboratory mouse, Mus musculus, belongs to the order Rodentia and family Muridae. The mouse is probably the most genetically and biologically characterized mammal in the world. Mice were first used for the study of respiration in the 17th century. Later in the 19th and 20th century they were bred for their coat color, and subsequently characterized genetically. Most of the common laboratory strains were developed in the early 1900s.
The mouse has short hair, a long naked tail, rounded erect ears, protruding eyes, a pointed snout and five toes on each foot. Mice come in a variety of colors. Insert picture of a mouse, get those cool ones.
Mice have a pair of incisors and three pairs upper and lower of molars. Molars are permanently rooted while the incisors have an open root and grow continuously. Due to this continuous growth of the incisors mice can have problems with overgrown teeth when the upper and lower incisors do not meet properly (malocclusion). Malocclusion can be hereditary or follow trauma, disease or inappropriate diet and/or soft food. There is no permanent cure for overgrown teeth the only treatment is to trim the teeth every 2-3 weeks, if malocclusion persists. Insert pictures.
Mice have a large horseshoe-shaped Harderian gland deep within the orbit. Secretions from the gland contain varying amounts of reddish-brown porphyrin pigment depending on the physiologic state, age, strain and sex of the mouse. The amount of secretions increases during stress and appears as 'red crusts' around the eyes and nostrils.
Males 20-30 g, Females 18-35g
|Birth weight||1-2 g|
|Heart rate||310-840 beats per minute|
|Respiratory rate||80-230 breaths per minute|
|Blood volume||7-8%, 1.5-2.5 ml|
|Urine volume||0.5-1 ml per day|
|Allergens||Dander, urinary protein|
Mice are communal animals, which live in a very hierarchical society. Within these groups they will aggressively defend their territories. Mums with newborns will likewise aggressively defend their pups and territories. Mice are nocturnal animals but adapt to their environments. While mice are timid, they may bite. Most of mouse behavior is pheromone driven.
Mice are communal animals with a social hierarchical system. This hierarchical system creates a situation whereby the dominant animals often barber (chew off the hair) those of lower rank. This behavior is especially evident among females and is more common in some strains than others. Subordinate animals may have their whiskers, trunk or flank hair removed. There are some suggestions that it may be a cooperative and/or learnt behavior. It appears to be of no sequelae for the health of the mice. Separating the dominant mouse often leads to rise of another dominant mouse. It is important that barbering is differentiated from other causes of hair loss or skin problems e.g. mites, fungi or bacteria by a veterinarian.
Mice have two distinct cervices and uterine bodies. There are separate urethral and vaginal openings. There is a vaginal closure membrane, which is lost at puberty. The inguinal canal remains patent throughout life. Mice have an os penis or os clitoridis associated with external genitalia.
Mice have a 4-5 day estrous cycle, divided into characteristic phases: proestrus, estrus and metestrus. The stage of the estrous cycle can be determined by vaginal cytology. Ovulation occurs at the end of metestrus. Mating leads to formation of a vaginal plug. Plugs persist for 16-24 hours and may last as long as 48 hours. Pregnancy lasts 19-21 days. Females will build a nest prior to parturition if opportunity is provided. Birth usually occurs at night with 10-12 pups being born. Stretching and hindleg extension are usually signs of impeding birth. Babies are born either head or tail first (breech). The female usually eats the placenta. Delivery lasts 1-4 hours, if labor persists call a veterinarian (5-3713). There is a fertile postpartum estrus. Maternal antibody is transferred to the fetus in utero and to the newborn via colostrum.
The young are born incompletely developed (altricius). They are born hairless and their eyes open after 10-12 days. Young are weaned after 21 days at which time they are 10-12 g. Puberty is attained at 4-6 weeks. Breeding onset is 7 weeks and breeding life is 7-9 months. It may be preferable to replace breeders when they are 6 months old.
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Mice are generally fed a diet containing low fiber (5%), protein (20%) and fat (5-10%). Feed may be pelleted or powdered. The pelleted feed is supplied as regular, breeder, certified, irradiated or autoclavable. Mice are usually supplied feed free choice and they eat 4-5 g a day (12 g/100 g body weight/day). Water is supplied free choice and they usually drink 3-5 ml a day (1.5 ml/10 g body weight/day). Water may be supplied using a bottle or automatic waterers, and may be further treated by reverse osmosis, ozone, ultraviolet radiation, hyperchlorination or acidification. Picture caging, types of feed, feeders, watering systems
Mouse rooms are usually maintained at 30-70% relative humidity and a temperature of 18-26ºC (64-79ºF) with at least 10 room air changes per hour. The mice are housed in standard shoebox cages with or without filter tops. Filter tops prevent cross contamination of mice limiting the spread of disease and keep facilities clean. Cages with filter tops may have a slightly higher temperature, relative humidity, carbon dioxide and ammonia than the room air. Microisolator tops provide even a higher level of protection than bonnet type filter tops, since they seal better. Static cages as described above are usually changed one to two times a week depending on cage density and housing style. In ventilated cages air is forced into the cage at up to 60 air changes per hour. This keeps the cage dry and reduces build up of ammonia and carbon dioxide. In such situations cages are changed once every 1-2 weeks. Ventilated cages may be kept positive or negative to room air depending on the study being performed.
Mice are usually provided with some kind of bedding in the shoebox cages. Bedding can be paper, wood shaving, wood chips or corncob. In very rare instances mice are housed on wire floors.
Mice should always be clearly identified on cage cards indicating protocol number, strain, sex, age, supplier, investigator and contact person. Procedures performed on the animal should be clearly indicated. Individual mice can be identified using ear punches, ear tags, tattoos, fur dyes, indelible mark on tail or microchips. Picture with standard ear numbering.
Sex is determined using the anogenital distance. Males have a greater (1.5-2 times) anogenital distance than females as well as a larger genital papilla. In neonatal males the testis may be visible through the abdominal wall. Conspicuous bilateral rows of nipples are visible in females at about 9 days of age. Absence of testicles is not a useful criterion for sexing since the testis is retractable into the open inguinal canal throughout life.
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Mice should be acclimatized to handling (gentling) to reduce stress. Always talk quietly, move hands slowly and handle them frequently. Mice should be handled at the base of the tail using your fingers or with forceps. Transfer the mice to a firm surface and apply a scruff hold to the loose skin between the ears with your fingers and forefingers while maintaining a grip on the tail. Do not pull the skin too tightly as the mice can choke, too loose a hold will allow the mouse to turn its head and bite. This hold allows you to examine the under belly and perform other procedures. A variety of restraint devices are available to assist in handling mice.
An adult mouse has a circulating blood volume of about 1.5-2.5 ml (6-8% of the body weight), however in older and obese animals this may be lower. Up to 10% of the circulating blood volume may be taken on a single occasion from a normal healthy animal on an adequate plane of nutrition with minimal adverse effect. Always make sure the animal has recovered safely from the procedure and give warm isotonic fluids. This volume may be repeated after 3-4 weeks. For repeat bleeds at shorter intervals, a maximum of 1% of an animal's circulating blood volume can be removed every 24 hours.
Blood can be collected from several sites in the mouse including tail vein, saphenous vein, retro-orbital sinus, brachial vessels, vena cava or cardiac puncture. Always ensure complete hemostasis before returning the mouse to its home cage.
It may be necessary to warm the tail by exposing it briefly to a heat lamp or placing it in a bowl of warm water. The mouse should be restrained in a device for the collection. Blood can be collected from the tail vein (and artery) by making a snip in terminal =3 mm of the tail with a scalpel or sharp scissors. Stroke the tail gently with thumb and finger to enhance flow of blood into the collection vial. Because of the thermoregulatory function of the tail no more than the distal 3 mm should be taken at a time. At the end of the collection apply pressure to the cut end with a gauze bandage and ensure that blood has completely stopped flowing before returning the mouse to the cage. A small nick can also be made at side of the tail 0.5 -2cm from the tail base to collect blood. A fine gauge needle introduced through the skin at a shallow angle can be used to withdraw blood from the tail vein. Apply a tourniquet around the base of the tail to aid in the collection. A butterfly catheter with only about 5 mm of tubing attached to it (rest cut off) may be used instead of a needle and syringe.
Restrain and extend the hind leg applying gentle downward pressure above the knee joint. This stretches the skin making it easier to shave and immobilizes the saphenous vein. Wipe the shaved area with alcohol or sterile lubricate gel and use a 25-gauge needle to puncture the vein (Vein is highlighted by the dark line in the picture below). If done correctly a drop of blood forms immediately at the puncture site and can be collected in a microhematocrit tube. Gentle pressure over the puncture site or relaxation of the restrainer's grip is usually sufficient to stop the blood flow. The scab at the puncture site can be rubbed off at a later date to allow additional blood collection.
The retrorbital sinus is a system of dilated venous channels at the back of the orbit. Blood can be collected form this area in anesthetized mice using a microhematocrit tube. There should be no movement of the head during the procedure. Pressure down with the thumb and forefinger just behind the eye and pull back on the skin to allow the eyeball to protrude. Position a microhematocrit tube along the inner corner of the eye (medial canthus) beside the eyeball. Insert the tube gently but firmly through the conjunctiva towards the back of the eye along the orbit. Rotate the tube gently as you proceed. Blood should flow freely, if, the tube is properly inserted. Tilt the head slightly downward to improve flow. After collecting the blood withdraw the tube and apply pressure on the closed eyelids to stop any bleeding. Remove excess blood with gauze. Complications include damage to the eye and surrounding tissues.
Blood can be collected from the brachial plexus as a terminal procedure in deeply anesthetized mice. Make a cut through the skin at the side of the thorax into the angle of the forelimb (axilla) to expose the axillary vessels. Transect the vessels and allow blood to pool into the pocket created by tenting the skin. Aspirate the mixed venous arterial blood is into an appropriate receptacle.
Vena cava and abdominal aorta
Blood can be obtained from the posterior vena cava or abdominal aorta in a deeply anesthetized mouse following laparotomy. Approach the vessel at a shallow angle using a fine gauge needle attached to a small syringe. This is a terminal procedure.
Up to 1 ml of blood can be obtained from the heart of a deeply anesthetized mouse in a terminal procedure. The most common approach is to lay the mouse on its back and insert a 25 to 30 gauge needle attached to a 1ml syringe just behind the xiphoid cartilage and slightly left of the middle. The needle should be introduced at 10-30 degrees from the horizontal axis of the sternum in order to enter the heart. Alternatively approach the heart laterally immediately behind the elbow at the point of maximum heartbeat.
Materials to be administered to mice can be given orally e.g. in water or feed or injected systemically through a variety of routes. The average daily consumption of feed and water for an adult 25 g mouse is 3-5 g and 4 ml respectively. The following volumes can be injected into mice safely: 2-3 ml subcutaneously, 0.05-0.1 ml intramuscularly (0.03 ml per site), 0.05 ml intravenously, 0.1-0.3 ml into the stomach and 2-3 ml intraperitonealy. Intramuscular injections are usually not recommended in mice because of the small muscle mass. A fine gauge needle should be used to make injections in the anterior thigh muscle. It is good practice to use a new needle each time you perform an injection.
Oral gavage is performed using a ball ended feeding needle. Estimate the distance that the needle needs to be inserted into the mouse (usually from the nose to the first rib) and mark it on the needle. Restrain the mouse with the head and body extended as straight as possible to facilitate introduction of the gavage needle. Introduce the needle in the space between the left incisors and molars, and gently direct it caudally toward the right ramus of the mandible. The mouse usually swallows as the feeding tube approaches the pharynx, facilitating entry into the esophagus. If the animal struggles or appears to be in respiratory difficulty withdraw the tube and begin all over again. Once the desired position is attained, inject the material and withdraw the syringe. Monitor the animal after the procedure to ensure that there are no adverse effects.
Subcutaneous injections are usually made into the loose skin over the neck or flank using a fine gauge needle. Insert the needle 5-10 mm through the skin before making the injection. Lack of resistance to the injection is indicative that you are in the right location. Check for leak back especially if a larger volume is injected.
Intraperitoneal injections are usually made in the lower right quadrant of the abdomen. The mouse is restrained with its head tilted lower than the body to avoid injury to internal organs. After swabbing the lower right quadrant with alcohol, a fine gauge needle is introduced slowly through the skin, subcutaneous tissue and abdominal wall. Withdraw the syringe plunger to ensure that you are not in the bladder or intestines. If nothing is withdrawn inject the material slowly. If you accidentally enter the bladder or intestines withdraw and discard the needle and syringe.
Intravenous injections are usually made into the dorsal tail vein. Warm the tail by immersing it in warm water or placing the animal under a heat lamp. The tail vein is easier to see in non-pigmented mice. A fine gauge needle should be used for this procedure.
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|Acute pain||Chronic pain|
|Anorexia (no fecal pellets)||Decreased body weight|
|Decrease in appetite (few fecal pellets)||Reluctance to move|
|Rubbing or scratching surgical site||Change in behavior|
|Biting or shaking affected body part||Poor grooming|
|Vocalization||Change in bowel or urinary activity|
|Restlessness||Rough hair coat|
|Porphyrin discharge (red-brown pigment around eyes and nostrils)|
For a detailed discussion of pain relief in mice refer to module 2. Generally opioids e.g. buprenorphine or non-steroidal anti-inflammatory agents e.g. acetaminophen, ketoprofen, caprofen, ibuprofen are used to relieve pain. Drugs can be administered in water, in jello, as oral drops, by gavage or injected. Drugs administered in water may be broken down in water, or insufficient quantities may be taken due to poor solubility in water or palatability problems.
In general inhalant anesthetics are safer than injectable anesthetics. Halothane and isoflurane are the safest ones to use. Methoxyflurane is no longer available. Use of ether at Johns Hopkins University is subject to restrictions due to safety concerns. Ketamine and xylazine is a common injectable anesthetic combination. Sodium pentobarbital can be used, but it has a narrow safety margin and is associated with a prolonged recovery period. For details on anesthetic techniques refer to the rodent surgery module.
Euthanasia in mice is most often performed by carbon dioxide asphyxiation or overdose of an anesthetic agent. Use of cervical dislocation or decapitation in absence of deep anesthesia must be scientifically justified. All individuals performing euthanasia must be properly trained. Individuals must also ensure that animals are dead before the carcass is disposed. Exsanguination or opening the thoracic cavity will ensure death. AVMA Panel on euthanasia report
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Diseases of mice are usually handled as a herd (colony) health problem rather than on an individual animal basis. The goal is to prevent introduction of a disease into a colony rather than to treat animals after disease outbreak. Disease prevention is practiced by institution of a disease surveillance (sentinel) program based on serological and microscopic diagnosis of problems in a representative sample of animals. Due to the widespread movement of animals all over the world with advent of genetic manipulation of animals, the possibility of introducing disease agents in a colony has markedly increased. The expanded use of genetically modified and immunocompromised animals greatly exacerbates the problem. Furthermore the practice of transplanting tumor material into mice provides a portal where these agents can be introduced into animal, especially if the tumors are not screened for adventitious infectious agents. Some important mouse diseases are discussed below to draw attention to the need to adhere to practices recommended by the veterinary staff to avoid these diseases.
Pinworms (Syphacia and Aspicularis) inhabit the intestine (cecum, rectum, colon) and have a direct lifecycle. The eggs are particularly resistant and survive for a long time in the environment. The disease is usually subclinical being marked in weanlings and immunocompromised animals. Symptoms include poor body condition, rough hair coat, reduced growth rate and rectal prolapse. Infection with pinworms has a negative impact on gastrointestinal, growth, behavioral and immunology studies.
Mites affect the skin of mice and up to 100% of the animals may be affected. Affected animals are scruffy, pruritic (itchy), loose hair and have scratch wounds, which can become infected with bacteria. There are changes in the immune responses of affected animals.
This is a viral disease of mice that affects multiple organs. Weanlings are important in maintaining the disease in a colony. Outbreaks result in widespread deaths in neonates and occasionally weanlings, with or without diarrhea. Mouse hepatitis virus causes a wasting disease and high mortality in immunocompromised animals. Usually 100% of the animals are infected. This disease wreaks havoc in a colony, with disruption in research especially in oncology, transplantation, immunology, gastroenterology, metabolism and transgenic technology.
This viral disease is the worst nightmare in a mouse colony. There have been recent outbreaks at several facilities in United States associated with the injection of mouse sera or tissue culture material containing mouse sera into mice. The disease produces massive die off in adult mice and amputation of the limbs (ectromelia) in surviving animals. Pox lesions (mousepox) appear on the skin. There is conjunctivitis, hair loss, as well as swelling of the liver, spleen and lymph nodes. Ectromelia causes high mortality and drastic measures including depopulation are usually taken to eliminate the disease.
This is a fungal disease affecting a wide range of laboratory animals and humans. The organisms are primarily localized in the lungs but may also involve other organs including the eyes, skin etc. It causes a slowly progressive chronic pneumonia with weight loss and eventually death in a large number of immunocompromised animals. The disease has a severe negative impact in research involving immunocompromised animals, pulmonary function and immunology.
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Mouse Genome Informatics
Mouse Phenome Database