• Fix
  • Neuter (a term generally reserved for males)


Spay refers to the removal of the ovaries – with or without the uterus – rendering the animal sterile.

Ovariectomy is the removal of the ovaries, without removal of the uterus or cervix.

Ovariohysterectomy is the removal of the ovaries and uterus, all or in part.

Clinical Signs

Intact female, the presence of ovaries and a bicornuate uterus.


  • To prevent unwanted pregnancy if combining with male cage-mates.
  • To reduce/prevent hormone dependent mammary tumors and pituitary tumors.
  • To prevent uterine/ovarian masses and infection.

May observe the following:

  • Enlarged, firm abdomen.
  • Foul odor from vaginal orifice.
  • Unusual discharge from vaginal orifice (e.g., discolored, blood tinged mucus [unrelated to normal delivery of pups], drops or streaks or frank bleeding).
  • Prolapse tissue from vaginal orifice.
  • Dystocia (difficulty or inability of uterus to contract and relax in effort to birth young).


The purpose for doing an ovariectomy or ovariohysterectomy in the female rat is to prevent pregnancy in the event of being housed together with male rats, or to prevent/reduce/treat cystic ovaries, hormone dependent mammary and pituitary tumors, and uterine diseases (tumors, pyometra) that are influenced by the release of hormones from the ovaries. Ovariohysterectomy is also performed in cases of dystocia to save the life of a pregnant doe, or in cases of uterine prolapse.

Although female rats may be spayed at any age, studies indicate that the spaying of female rats at a younger age (3 to 4 months) appears to be more effective at reducing and preventing hormone induced mammary tumor and pituitary tumor growth than if a female is spayed later in life (reference to ovariectomy at 90 days: Hotchkiss, 1995). In addition, since the majority of uterine diseases (tumors) tend to occur in the aged female rat, doing a complete spay at an early age (such as at 3 to 4 months) can be preventative.

The following sections provide the rat owner with instructions on how to prepare their female rat for the day of surgery, as well as the type of care, preparations, procedure(s), and recovery the veterinarian may follow. There are also sections with home care recommendations and additional comments that may be helpful.

Assessment and Checkup Prior to Surgery

The rat should be thoroughly evaluated for any signs of respiratory disease (as well as heart disease in older rats). This is particularly important since rats are obligate (i.e., preferred; allowing them to scent while eating) nasal breathers, having their soft palates permanently engaged around the epiglottis. Therefore, any respiratory secretions that are already present and thick or increased due to infection may become more thick and sticky while the rat is under anesthesia, as a result of decreased respiratory rate and movement of secretions. This could potentially make them more susceptible to respiratory arrest if these secretions or other causes block the airways. (Girling, 2013) *Note: although rats are identified as obligate nasal breathers, they do have a patent pathway from mouth to lungs for endotracheal intubation.

For rats with a history of respiratory illness, or for those rats that are elderly and where the pain and stress of a procedure can further suppress the immune system, it is advisable for the veterinarian to place the rat on an antibiotic with coverage for mycoplasma as a prevention to relapse. The antibiotic should be started a few days prior to surgery and continued until the surgical incision site has healed. *Note: spays in elderly rats should be limited to response to clinical problems with the reproductive tract, not done as an elective to prevent potential mammary tumors. Tumor preventative spays need to be done early in life, optimally 3 to 4 months of age, to be effective.

Ideally, a baseline minimal screening (hematocrit, total protein, blood glucose, blood urea nitrogen, urine specific gravity) may be advised preop. However, there is a wide range of normal lab values reported in rats, and depending upon the procedure and age of the rat – unless the rat has a previous health history – it may not always be necessary. (Bruce & Olsen, Ovariectomy, ovariohysterectomy and orchidectomy, 1986)

Preparation Day of Surgery

It is not necessary to “fast” rats prior to surgery. Rats do not vomit. Providing food and water up to the time of preop prepping (just prior to anesthesia being given) helps to prevent the rat from becoming dehydrated, or suffering ketoacidosis/hypoglycemia during surgery and recovery. *Note: some surgeries such as gastrointestinal surgery may require withholding food for 2 to 3 hours prior to the procedure, but water should be made available until just prior to anesthesia.

The pet rat should be brought in a small one level cage or carrier where the rat can be easily observed from all sides of the carrier by the veterinarian and staff. It is helpful to attach a small tag to the carrier with your rat’s name, sex, age, color/markings, type of surgery, as well as your name and a phone number where you can be reached. Also note whether or not the rat can be handled safely. Some rats may become frightened around unfamiliar humans, such as: semi-wild rats, or rats that may have been abused and rescued.

Provide non-ravel cloth (e.g., fleece or t-shirt type material) for bedding, preferably white or light-colored, so that signs of urination or bleeding are easily visible. In addition, provide an extra clean set of bedding if the rat is expected to wait awhile before the surgery (the bedding may become soiled).

A filled water bottle and the rat’s food – including fresh fruit and small treats – should accompany and be accessible to the rat. Note that the water bottle may need to be lowered during the recovery period, if the rat cannot sit up comfortably. In addition, it is advisable to include high caloric foods (e.g., pasta, porridge, oatmeal, farina, etc.) in the diet during a few days leading up to the day of surgery. Doing so can aid metabolic function, reduce hypoglycemia, and may improve healing time.


It is imperative to obtain an accurate bodyweight for correct dosing of anesthetics and analgesics.

Making sure the rat is well hydrated helps to prevent dehydration throughout the perioperative period. Note that the maintenance fluid volume for rats is approximately 100 mL/kg/day. Warmed, IV or SQ, crystalloid fluids (e.g., lactated Ringers or Hartmann’s solutions or normal saline) may be required to maintain hydration perioperatively.

Since rats possess serum atropine esterase, which rapidly hydrolyzes atropine, administering atropine for secretion reduction or for its vagolytic effects may not be that effective. In cases where secretion reduction is necessary, and depending upon the procedure, the veterinarian may elect to use glycopyrrolate. (Bruce & Olson, 2008; Harkness, et al, 2010 )

Preemptive analgesia should be used cautiously in rats. Although it may be desirable to reduce effects of pain upon arousing in recovery, and provide the ability to reduce the percentage of gas anesthetic when giving an opioid such as buprenorphine prior to the anesthetic, both opioids and particularly NSAIDs (such as Metacam or carprofen) are best used postoperatively when blood pressure is more stable and the side effects less. *Note: that the use of NSAIDs alone does not require reduction of gas anesthetic. (Flecknell & Richardson, 2009; Bennett, 2012)

Providing oxygenation to the rat prior to intraop (along with providing oxygen intraop and postop) will significantly help prevent hypoxemia and hypocapnia.

Minimizing the area of fur shaved at the site for incision, prepping with warmed surgical scrub such as chlorhexidine and avoiding use of alcohol prep, and providing absorptive towels to prevent the rat from becoming wet will help to decrease development of hypothermia.


Oxygen should be continued through to recovery to prevent hypoxia from developing.

It is important for the veterinarian to be prepared for emergencies prior to anesthetizing the rat. Any emergency drugs should be prepared and ready, at the correct dosage.

While some anesthetic agents are not appropriate for use in rats due to an increased risk of anesthetic death, the following are those with the widest margin for safety:

    Inhalation Anesthetics

    Isoflurane or Sevoflurane are considered the safest and most effective anesthetics for use in rats. The advantage is that each of these inhalation agents permits a quicker induction and recovery, being only minimally metabolized in the liver with the majority being eliminated through exhalation. This is particularly helpful in older rats or those rats with subclinical hepatic issues. While these inhalant anesthetics do have the effect of hypothermia and drying of respiratory mucous membranes, these effects can be countered by the use of a recirculating warm water pad or warm air blanket and monitoring the rat’s temperature, and keeping the rat hydrated, respectively. (Flecknell & Richardson, 2009 ; Girling, 2013)

    Injectable Anesthetics

    When administering injectable anesthetic agents those that allow for a specific antagonist to be used when the agents must be reversed quickly provide a better margin of safety. Agents such as xylazine and medetomidine can be reversed with atipamezole. Using either xylazine or medetomidine in combination with Ketamine allows for lower dosage of the drugs to be given, decreasing unwanted side effects. IV or IP administration is preferred over IM or SQ. IV administration allows the effects of the drugs to be seen quickly and for the dosing to be adjusted. Since these substances are irritating to tissue, giving them SQ or IM can result in nerve irritation and/or muscle necrosis at the injection site, as well as self-mutilation, if care is not taken when injecting. Injecting into thigh muscle should be avoided for this reason. A disadvantage of injectable anesthetics is that reversal and recovery take longer because they are required to be metabolized before being eliminated by the body, and the fact that they can cause severe hypotension. (Flecknell & Richardson, 2009)

The eyelids of rats do not always close completely under sedation. Placing ophthalmic lubricant on the eyes will prevent corneal desiccation (i.e., severe drying out of the cornea).

It is important to prevent hypothermia from developing. Anesthetic agents suppress the body’s ability to regulate temperature. Rats physically have a large body surface area to volume ratio and when inactive due to being sedated or anesthetized they can rapidly become hypothermic. When performing surgery on the rat it is therefore important that it be performed on a recirculating warm water pad, or warm air blanket. In addition, draping the rat as soon as possible while in surgery can decrease heat loss. (Mayer, 2013) While warm water bags and heated pads regulated to 30 to 35 degrees centigrade (85 to 95 degrees Fahrenheit) and covered by a towel before placing the rat on it may also be used, this should be done carefully so as not to cause burns to the sedated rat. The temperature should also be carefully monitored throughout the surgery and recovery to ensure that the rat does not become overheated. Hyperthermia in the rat can be just as deadly as hypothermia.


The ovaries found on each side of the abdomen are associated with the caudal pole (slightly below) of the kidneys and are embedded in fat. Each ovary is attached to its own uterine horn of the bicornuate uterus via an oviduct, which is a long, convoluted, tightly packed Fallopian tube. In the female rat the uterine horns enter separately but open directly into a single cervix, connecting the uterus and vagina. A single artery and vein run along the entire length of the medial side of each ovary and uterine horn.

Types of procedures that may be performed are the following:


    A dorsal approach is most commonly used for an ovariectomy in the rat. The anesthetized rat is placed in a ventral (sternal side down) recumbency (laying) position, and the fur clipped (shaven) along the spine from the mid thorax to the sacral region. A longitudinal incision is made from the second to fifth lumbar vertebrae. The skin is then retracted from one side to the other to remove each fat-embedded ovary through the same skin incision. Care is taken when removing the ovaries to avoid the spleen, liver and kidneys. (An alternative method is to do a transverse flank incision on each side – caudal to the last rib and ventral to the epaxial muscles – to remove each ovary.) A ligating suture is placed on the ovarian vessels to prevent hemorrhage, although this is usually slight in an ovariectomy. The pedicle is then transected distally to the ligated vessels, and the ovary and oviduct are removed. With either the single midline incision or the bilateral incisions the ovaries and entire oviduct should be removed intact to avoid having any portion or small pieces remain. This prevents reimplantation and the retention of estrus activity.


    For an ovariohysterectomy the anesthetized rat is placed in a dorsal (place rat on its back) recumbency (laying) position, with the tail towards the veterinarian. A small ventral midline incision (approximately 2 to 4 cm) is made through the skin and muscle, from the umbilicus to the pubis anterior to the urethral opening, taking care not to cause damage to viscera (i.e., intestines, urinary bladder). The uterine horns are then traced back cranially (towards the head of the rat), where the ovarian pedicles and ovarian vessels are ligated and then transected distally to the ligatures. The uterine horns, oviducts and ovaries are gently freed, and the uterus and uterine vessels are ligated cranially to the cervix – which helps to prevent urine reflux into the abdomen (this may occur if urine is allowed to pool in the vagina and leak back into the surgical site when the uterine horns are removed) – before being transected distally to the ligature there. The small portion of the uterus, along with the uterine horns, oviducts and ovaries, are then removed. (Waynworth & Flecknell, 1992; Harkness,
    et al, 2010 ; Bennett, Rodents: soft tissue surgery, 2009; Bennett, Soft Tissue Surgery, 2012)

Closure in both types of procedures can then be done using the same types of materials. The muscle wall is closed with 4/0 UNDYED VICRYL absorbable suture material that is tissue friendly in rats, producing the least incidence of suture reaction. Chromic gut is not advised due to its high tissue reactivity. Skin may be closed with either non-absorbable sutures or autoclips or with surgical adhesive. *Note: it is important not to draw sutures too tightly in order to prevent tissue strangulation; and when using surgical adhesive to avoid getting it on surrounding outer skin as this can lead to self-trauma by the rat.


Case histories


  • Fig. 1a: Ovariectomy in 1-year-old female rat (Octopus). *Warning graphic. Includes necropsy photos.*
  • Fig. 1b: Ovariectomy in 1.5-year-old female rat (Cassowary). *Warning graphic.*
  • Fig. 1c: Ovariectomy in 2-month-old female rat (Goose)

Surgical pictorials


  • Fig.Spay 1 Routine ovariohysterectomy (spay) in young adult female rat


    Female Rat Reproductive Anatomy

  • Fig.3a: Female rat reproductive anatomy and description

Postop Recovery

To reduce any concern with regard to hypothermia, the rat should be dried with a soft towel of any remaining wet drainage from the surgery and be placed in a quiet, warm enclosure (e.g., incubator), on non-ravel, non-loop, white or light colored cloth to monitor for any signs of hemorrhage (bleeding).

Continue oxygen and supplemental heat and monitor for and correct any signs of hypoxia, hypothermia and/or hyperthermia.

Turning and repositioning the rat approximately every 10 minutes – until its crawl reflex returns – will help to stimulate respirations during recovery and reduce the chance of pulmonary congestion developing. (Bruce & Olson, 1986; Girling, 2013)

If recovery is prolonged, warmed, IV or SQ, crystalloid fluids (e.g., lactated Ringers, Hartmann’s solution, Normal saline) should be continued or instituted, to prevent dehydration, until the animal is drinking and eating on its own.

Once the righting reflex returns the rat can be placed in a small cage or carrier, again on non-ravel light-colored cloth or light colored paper towels, with provision for continued warmth and monitoring.

Encourage oral food and water intake upon being recovered from anesthesia. Small treats, fresh fruit or baby foods may be more welcome to the rat before moving to pelleted or block food.

Monitor the rat to ensure that the ability to urinate and defecate has returned.

Managing pain in the rat with analgesics, such as with the use of buprenorphine and/or with NSAIDs such as meloxicam or carprofen, can help aid in recovery and reduce the chance that the rat will bother the incision site.

Antibiotics should be continued or initiated if the rat has underlying illness, or in the event that aseptic technique has not been maintained, or in the case of developing infection.

Discharge Instructions

Pain medication such as Metacam (which is an NSAID) and buprenorphine given SQ is recommended to be given postop for 3 to 5 days as directed. Inform the owner of the time when the rat may next receive pain medication.
Metacam alone is not enough to control deep abdominal pain, and may take up to 24 hours to reach full effectiveness. If the owner is not capable of giving SQ injections of buprenorphine at home, then give a post op injection of buprenorphine at the clinic, and dispense Metacam for use at home. The dose of oral buprenorphine needed to significantly increase pain threshold is so high, it is not practical to give at home, nor is it cost effective enough to dispense. (Comparative Medicine, Vol. 51, No. 1, February 2001) However, buprenorphine is highly effective given SQ.

*Note: that pica has been known to occur in the rat with the use of opioids such as buprenorphine and that it is primarily dose related. Upon discharge, in the event that the rat is receiving an opioid for pain control, instruct the pet owner to monitor for pica type behavior (i.e., chewing and ingesting of non-food substances) and have the pet owner remove the rat from any bedding source that could be chewed and ingested and to provide additional fluids in the rat’s diet until the drug has been eliminated from the rat’s system. Contact veterinarian if behavior becomes pronounced.

Provide and continue antibiotic if necessary.

There are no diet restrictions. Free access to food and water should be provided. Offering additional flavorful softer foods, treats and fruits will encourage the rat to return to a normal appetite and provide additional fluids. In the event that the rat has stopped eating or drinking fluids, or has not defecated or urinated by the morning following the day of surgery, notify the veterinarian!

For the first 24 to 72 hours (or as directed if there are complications) after arriving home the rat should be housed in a one level cage, alone, on paper towels or non-ravel cloth (e.g., fleece or t-shirt type material) as bedding. This will enable better monitoring of the incision site, fluid intake and appetite, and if the rat is passing urine and feces.

Following the first 24 to 72 hours, if the rat is stable, the rat can return to supervised visits with cage-mates. Housing with cage-mates may be resumed when assured that they will not cause injury to or bother the rat’s incision site.

Monitor the incision site closely for infection or wound disruption.

Return to the veterinarian for a wound check, and to have sutures or wound clips removed, in 10 to 14 days.

A wound wrap, as explained and demonstrated by L.Pulman LVT,LATg, here:, may be required in the event that the rat has attempted or attempts to chew at the surgical site or sutures. An Elizabethan collar should only be applied as a last resort since this can prevent the rat from drinking from a water bottle, or holding its food when trying to eat.

Nursing Care at Home

  • Contact the vet if any of the following are observed:
    • Swelling, redness, bleeding, disruption (reopening) of the surgical site, or if there are signs of pain unrelieved by pain medication, noted by reduction of appetite, decreased drinking, hunched posture, rapid breathing, repeated tremors or abdominal stretching, aggression in an otherwise non-aggressive rat, or signs of pica (see below).
    • If there is absence of passing feces or urine, or if the rat is not eating or drinking by the morning following the day of surgery, or if there are signs of increased weight loss, unusual rapid weight gain, or lethargy.
    • It is important to be aware that pica (i.e., chewing and ingestion of non-food substances) may occur where pain has not been adequately addressed in the rat, or as a dosage related side effect of having been given an opioid such as buprenorphine. In this case, remove the rat from any bedding source that might be chewed and ingested, and provide additional fluids in the diet until the medication has passed through the rat’s system. Contact the vet if the behavior becomes pronounced.
  • Provide a hospital cage for the first 24 to 72 hours upon arriving home, or longer if there are concerns that the cage-mates may groom the sutures or surgical site. Supervised visits with cage-mates can be resumed 24 hours after returning home if there are no complications. Housing with cage-mates may be resumed when assured that they will not cause injury to or bother the rat’s incision site.
  • Provide clean bedding daily, such as fleece or soft t-shirt type material or ink-free paper towels. Avoid using material such as terry cloth type towels that can ravel. Also avoid litter-type bedding until the incision has healed, to prevent the chance of wound contamination or infection.
  • Provide additional warmth to maintain body temperature within normal limits. It is essential that the rat does not become overheated or dehydrated. The rat should also be able to move away from the heat source if it becomes uncomfortable. If the rat is moving minimally, extreme care must be taken to keep the heat low and stable. The following are examples of warming products that may be used:
    • Isothermic products that are heated in the microwave, such as a SnuggleSafe®, may be used. Make sure to follow the product directions carefully and wrap it in a towel before placing it in the cage. The SnuggleSafe® will provide heat for 12 hours before needing to be reheated. Other similar types of products may vary in reheat time. Check the directions for the individual product.
    • If using a heating pad (good for long term use) use only the low heat setting, put a thick towel in between the pad and the cage bottom, and place it beneath a corner of the cage. Avoid use of heating pads with Aquarium-type glass tanks where over heating can occur rapidly. Care must be taken that the rat does not overheat!
    • If none of these options are available you can fill a sock with rice and heat it until it is warm in the microwave. An alternative is to fill 2 or 3 one gallon plastic jugs with hot water, place around the outside perimeter of the cage, and drape cage and jugs with towels to contain and direct the heat into the cage.
  • Medicate for post-op pain as directed by the veterinarian.
  • Give or resume antibiotic(s) as directed by the veterinarian.
  • Encourage the intake of oral fluids, such as water, Jell-O water, or electrolyte replacement drinks such as Pedialyte or Gatorade (which can be found in local grocery stores).
    • Please note that Pedialyte is only good refrigerated for 24 hours after opened, but can be frozen as ice cubes and thawed as needed.
    • Note: a juicy type of fruit also provides an additional fluid source in the diet.
  • For 24 to 72 hours post-op, feed iron-rich foods to prevent anemia (e.g., cooked beef/chicken liver, cooked beef/turkey/oysters, scrambled or hard-boiled eggs) and additional high caloric foods, along with continuing the rat’s usual diet.
  • A body wrap may be required if the rat will not leave the surgical site alone. If needing to reapply see directions for post op wrap by L.Pulman LVT,LATg, here:
  • In the event of dried or excess drainage, the incision site may be cleaned with a moistened Q-tip (swab), using warm water or normal saline. Avoid saturating the incision site.
  • In the event a crust develops over the healing incision site the area of crust can be softened by applying a non-Vaseline ointment, such as Aloe gel or a Calendula lotion, that is water soluble and pH balanced.


  • Post-op pain relieved
  • Incision site remains free from infection
  • Appetite and fluid intake normal
  • Abnormal weight loss prevented
  • Feces and urine output normal
  • Incision heals normally
  • Quality of life improved
  • Comfort and quality of life increased where spay has been performed in cases of disease/disorder or emergency.

Prevention of complications

  • Assess the rat’s health and stability prior to the procedure. Use caution with rats that have signs of respiratory disease, are obese or elderly.
  • Refrain from withholding food and water prior to or following the surgery, to prevent hypoglycemia or dehydration in the rat.
  • To prevent hypothermia, monitor the body temperature throughout the procedure and postop, by using warm water circulating mats or heated pads at a controlled temperature. Note: monitor the body temperature closely as hyperthermia can be just as deadly to the rat as hypothermia.
  • Determine the appropriate anesthetic agent for use in the rat, based on age, health and type of procedure. Note: reversal agents should be at the ready when using injectable anesthetic agents.
  • Avoid giving injectable anesthetics IM or SQ to prevent the potential for skin and muscle necrosis and sloughing and self-mutilation by the rat due to irritating agents.
  • Administer oxygen perioperatively to avoid hypoxemia and hypocapnia.
  • Ensure that the ligatures of the ovarian pedicles and/or uterus do not slip, causing hemorrhage.
  • Monitor for and correct signs of bleeding and/or fluid loss with warmed crystalloid fluids by parenteral (e.g., IV, SQ) routes as necessary.
  • Prevent wound dehiscence or disruption by managing pain and placing a wound wrap as needed. An Elizabethan collar should be avoided unless absolutely necessary, as rats utilize their hands to eat and need to be able to reach their mouth.
  • Provide a quiet environment to reduce stress and fear.


Pros and cons for the types of procedures:

  • Pro: Least invasive, prevents reproduction, prevents ovarian cysts and masses, reduces/prevents hormone related mammary tumors and pituitary tumor if done early in the rat’s life.
  • Con: Will not prevent uterine tumors, infection or prolapse.


  • Pro: Prevents reproduction, prevents ovarian cysts and masses, reduces/prevents hormone related mammary tumors and pituitary tumors if done early in the rat’s life, prevents uterine disorders, tumors, infection and prolapse.
  • Con: More invasive procedure.
  • Michael Hutchinson, DVM, Animal General, Cranberry Township, PA.
  • Lindsay Pulman, LVT, LATg

  1. Bennett, R. A. (2009). Rodents: soft tissue surgery. In E. Keeble & A. Meredith (Eds.), BSAVA manual of rodents and ferrets (pp. 73-85). Quedgeley: British Small Animal Veterinary Association.
  2. Bennett, R. A. (2012). Soft Tissue Surgery. In K.E. Quesenberry & J.W. Carpenter (Eds.), Ferrets, Rabbits, and Rodents, Clinical Medicine and Surgery (Third Edition ed.) (pp. 373-379). St. Louis: Saunders.
  3. Deerberg, F., Rehm, S., & Pittermann, W. (1981). Uncommon Frequency of Adenocarcinomas of the Uterus in Virgin Han:Wistar Rats. Veterinary Pathology, 18(6), 707-713. Retrieved May 13, 2014, from
  4. Dixon, D., Leininger, J., Valerio, M., Johnson, A., Stabinski, L., & Frith, C. (1999). Proliferative lesions of the ovary, uterus. vagina, cervix and oviduct in rats, URG-5. Guides for Toxicology Pathology. Retrieved May 13, 2014, from
  5. Flecknell, P., & Waynforth, H. (1992). Experimental and Surgical Technique in the Rat, Second Edition. New York: Academic Press.
  6. Girling, S. (2013). Part 1 Small mammals. Veterinary nursing of exotic pets (Second ed., pp. 36-47 and 91-97). Chichester, West Sussex: Wiley-Blackwell.
  7. Harkness, JE., Wagner, JE. (1983). The biology and medicine of rabbits and rodents, 2nd ed. Philadelphia: Lea & Febiger
  8. Harkness, J. & Wagner, J. (1995). Biology and medicine of rabbits and rodents Fourth Edition. Baltimore: Williams & Wilkins.
  9. Harkness, JE., Turner, PV., VandeWoude, S., et al. (2010). Harkness and Wagner’s Biology and Medicine of Rabbits and Rodents. 5th ed. Ames: Wiley-Blackwell.
  10. Hedenqvist, P. (2008). Anaesthesia and analgesia for surgery in rabbits and rats: A comparison of the effects of different compounds (Doctoral dissertation). Karolinska Institutet, Stockholm, Sweden. Retrieved May 21, 2014, from
  11. Hotchkiss, C. (1995). Effect of surgical removal of subcutaneous tumors on survival of rats. J Am Vet Med Assoc, 206(10), 1575-9. Retrieved January 1, 2009, from the Medline database.
  12. Martin, L., Thompson, A., Martin, T.,Kristal, M. (2001). Analgesic Efficacy of Orally Administered Buprenorphine in Rats. Comparative Medicine, copyright February 2001, American Association of Laboratory Animal Science, Vol. 51, No. 1, Pages 43-48
  13. Mayer, J. & Donnelly, TM. (2013). Clinical Veterinary Advisor: Birds and Exotic Pets. St. Louis: Saunders
  14. Mayer, J. (2008). Surgical techniques for spaying rabbits and rats. Proceedings of the North American Veterinary Conference: January 18-22, 2003, Orlando, Florida. (Small animalm edition ed., pp. 1854-5). Gainesville, FL: Eastern States Veterinary Association.
  15. Murray, M. (2006). Spays and neuters in small mammals. Proc of the North Am Vet Conf, pp. 1757-9. Retrieved May 13, 2014, from
  16. Olson, M., & Bruce, J. (1986). Ovariectomy, ovariohysterectomy and orchidectomy in rodents and rabbits. Can Vet J, 27(12), 523–527. Retrieved May 13, 2014, from
  17. Nagaoka, T., Onodera, H., Matsushima, Y., Todate, A., Shibutani, M., Ogasawara, H., et al. (1990). Spontaneous uterine adenocarcinomas in aged rats and their relation to endocrine imbalance. Journal of Cancer Research and Clinical Oncology, 116(6), 623-628. Retrieved May 13, 2014, from
  18. Richardson, C., & Flecknell, P. (2009). Rodents: anaesthesia and analgesia. In E. Keeble, & A. Meredith(Eds.), BSAVA manual of rodents and ferrets (pp. 63-72). Quedgeley: British Small Animal Veterinary Association.


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