General Surgery

Many behavioral neuroscience studies involve surgery on the animal. Surgery can allow the investigator to make strong statements about specific systems in specific regions of the nervous system. With lesions, an investigator can look at the necessary and sufficient properties of various regions of the brain. With a cannula or electrode implantation, the investigator can investigate various was of stimulating or suppressing activity in a certain region of the nervous system. Other surgeries allow for chronic implants for monitoring body systems, or delivering drugs over a long period of time. Finally, surgeries can used to control levels of hormones in the animal’s body (e.g., castration, ovariectomy, adrenalectomy, etc…).

Surgery is a serious endeavor, which should only be performed by, or under the close supervision of, a trained professional. Close and careful supervision is important to ensure that the animals are well treated. Listen to instructions and advice of your trainer, and leave the room if you feel uncomfortable. Surgery is stressful to watch and experience. No one will fault you if you need a time out. Please respect the surgeon and the animal. Do not get in their way or distract them. Be aware that there are very strict laws that regulate the use of animals in research, and these laws may stipulate how surgery may be performed, and by whom it can be performed. Before using an animal in research, and certainly before performing surgery on an animal, you should check institutional regulations as well as local and federal laws that are applicable to your project. The guidelines in this book conform to the guidelines of the Canadian Council of Animal Care and the policies of the University of Calgary, but may not be sufficient for your institution.

Surgical Area

The surgery room should be well ventilated with appropriate scrubbers on the ventilation system for any gaseous anesthetics in use. Ideally, it should be in an out-of-the-way place with minimal traffic and noise. The area should have a door which should be kept closed to prevent the unprepared and uninitiated from witnessing something they would rather not see. The surgical field should be made of a non-porous surface which can easily be cleaned and disinfected, such as stainless steel. Adequate lighting should be available, ideally in the form of a boom-mounted or a fiber-optic lamp. This will allow you to adjust the angle of the light source. One advantage of a standard incandescent lamp is that it will keep the surgical area warm. A homeothermic surgical pad can be used to help keep the body temperature near the normal physiological level during the surgery, as the animal’s ability to thermoregulate during surgery is compromised by the anesthetic. An area removed from the surgical area should be provided for preparing the animal for surgery (injections and hair clipping). This helps keep the surgical area sterile.

Standard supplies which are always handy include: 0.9% saline, 70% ethyl alcohol, distilled water, gauze pads, paper towels, and cotton swabs. Of course, many specific tools will be needed for each surgery, but having the above mentioned supplies at hand is a good idea. The surgeon should be wearing a clean lab coat whenever working with animals. When there will be an open wound, gloves and a facemask are essential. After washing your hands with a disinfectant soap, don a pair of latex gloves. I have found for stereotaxic surgery that it is best to wash and put your gloves on after you get the animal in the stereotaxic frame, but before you make the initial incision. The animal’s fur seems to stick to the latex, which makes the task of getting the animal in the ears bars that much more arduous. So long as you put your gloves on and disinfect the incision site before making the incision, sterility can be maintained.

Oxygen and Ventilation

Maintaining the animal on oxygen will help with recovery. As most anesthetics decrease respiration, so increasing the percentage of oxygen in the air which the animal breaths will help maintain an adequate level of oxygen in the blood. For rat surgery, the anesthetic machine should be an open system as opposed to the more typical Magill circuit used on larger animals. (the Magill circuit has a valve, and the animal must have sufficient lung capacity to force this valve open to expel the exhaust air, and rats do not have sufficient lung capacity). Many open circuit systems have optional scavenging units which create an active draw circuit and collect the exhaust anesthetic. The O2 flow should be set at 1L/min, with the isoflurane set to 2%. An animal will begin to look blue (cyanosis) if its blood-oxygen level drops too low.

Tools The tools required will depend on the particular surgery being carried out. The following is a list of commonly used tools:

Scalpel blade and holder: The blade used should be sterile, and either a #10 (curved) or #11 (straight edge). Never touch the blade with your fingers, use a clamping device to hold the blade when placing it on the holder.
Hemostats: These locking tools are classically used clamp off blood vessels (hence their name) but are indispensable for grasping and holding tissue. Because they lock closed, they can act as a third hand.
Forceps: These are ideal for grasping and manipulating tissue or equipment. They come in various sizes, curved or straight, tissue-grasping or normal.
Needle holders with scissors: these will hold suture needles and cut suture thread.
Scissors: For cutting tissue only.
Spatula: For separating bone and muscle, or for delivering and shaping dental acrylic.
Suture needle and thread: There are many different types of each available. Smaller is better for small animals. Monofilament sutures are preferred as braided silks have spaces that may harbor microorganisms. Catgut or absorbable sutures should be used for internal sutures, while a more durable thread can be used for external sutures. If the suture is in a place when the animal can chew on it, a hidden stitch should be used, or even better, surgical staples (Michel clips).
Surgical drill and bits: For drilling bur holes for the placement of implants or anchoring screws.
Jewelers’ screws and screwdrivers.
Dental Acrylic
All tools should be cleaned prior to surgery. A bead sterilizer is a good choice, and a few minutes at 220°C will kill most pathogens. Alternatively, a disinfectant bath of gremaphine, Zepherian’s solution or 70% ethyl alcohol are quite good at cleaning the tools and will kill bacteria in minutes, although bacterial spores will survive such treatment (Waynforth, 1980).
For completely aseptic sterile tools, more extreme measures are needed. Tools can be autoclaved (180°C, 15 lbs/square inch for 15 minutes). However, many facilities are not well enough equipped to have multiple sets of tools such that you can have a separate autoclaved set for each surgery (particularly if you do six or seven a day). Cleaning your tools with a bristle brush and clean water, followed by sterilization in the bead sterilizer or an antiseptic solution is sufficient for most rodent surgeries. If you find that your animals are getting infections, then obviously you’ll want to re-evaluate this step, and proceed with more stringent sterile cleaning of your tools.
Tools should be left to air dry at the end of the day after a final wash. They should be placed on clean paper towel and hemostats and scissors should be left open to air in drying and prevent rusting. A small tray is often helpful for transporting the tools from the sink to the surgical table. If you are autoclaving your tools, make sure they are encased in sterile packaging so that they their sterility can be maintained until you need them.

Pre- and Post Anesthetics

Various drugs can be used as a pretreatment prior to surgery. The use of any of these depends on the particular surgery and anesthetic being used. Be aware that drugs do interact, and in some cases certain combinations should be avoided. As well, animals will require various other medications during or following surgery. They will be described below as well.

Analgesics

Many anesthetics have no analgesic properties beyond those of obtained during general anesthesia. Although it seems like they should go hand in hand, they do only while the anesthetic is having an influence. As well, any tissue damage early in the surgery can lead to central sensitization, which may lead to a stronger sensation of pain after surgery (Yashpal, Katz, and Coderre, 1996). It has been suggested that a pretreatment with an analgesic will minimize pain when emerging from the anesthetic. Such an analgesic will have an effect prior to tissue damage (i.e., incision), and will ideally, still be active when the anesthetic has worn off. Such a pre-anesthetic would go a long way towards minimizing suffering. The major hurdle in this case is that some analgesics can have detrimental effects when given before surgery (e.g., some may interact with many anesthetics to decrease respiration, while others may impair) so careful, informed, evidence-based decisions should be made in consultation with a knowledgeable veterinarian. Butorphanol and Buprenorphine are two safe opiates that are partial agonists at the µ-opioid receptor and thus cause less respiratory depression than full agonists such as morphine, but yet produce excellent analgesic effects. The ideal situation would be where pain is manaded through multimodal analgesia, where an opiate, an anti-inflammatory, a local anesthetic and a general anesthetic are used in combination to abrogate pain through a variety of mechanism (Lichtenberger and Ko, 2007) A strong opiate should be given post-operatively. Buprenorphine is ideal as only a very small dose (0.1-0.5 mg/kg) is needed, and its duration of action in rats is much longer than the other opiates (~ 12 h). Other drug which work well include Butorphanol (0.05-2.0 mg/kg), Morphine (10 mg/kg) or Codeine (25-60 mg/kg) (Jenkins, 1987). µ-receptor agonist (such as morphine) may be a better choice than -agonists (such as Butorphanol and Buprenorphine) post surgically for management of somatic pain associated with skin and deep tissue damage (Lichtenberger and Ko, 2007). -agonists appear more suited to visceral pain (Lichtenberger and Ko, 2007). Buprenorphine has been reported to induce abhorrent eating behavior in rats, whereby the rat will consume its sawdust bedding, leading to damage of the gastrointestinal tract (Jacobson, 2000) so care should be used with this drug, particularly following a gastrointestinal surgery. Non-opiate drugs are available, and can be given in the following doses: Aspirin, 100 mg/kg; Acetaminophen, 110-330 mg/kg; Ibuprophen, 10-30 mg/kg (Jenkins, 1987; Flecknell, 1984). These generally provide more mild analgesia than the opiate analgesics, but are relatively safe and long acting (Lichtenberger and Ko, 2007). For long-term pain management, it has been suggested that animals be treated for 48-72 hours after surgery (Flecknell, 1984). This presents a problem for a lab with a small staff doing surgery on a large number of animals. One approach has been to deliver drugs in drinking water. Cooper, DeLong and Gillett (1997) determined that acetaminophen in drinking water up to doses of 600 mg/kg/d did not provide any measurable analgesia. Buprenorphine (2.9 mg/kg/d) did work in their paradigm (paw withdrawal reflex) when delivered in drinking water. Both drugs did work when delivered parentally. Based on these results, acetaminophen, if used should not be given in the drinking water, as it is unlikely to provide any pain relief. Cooper, et al. (1997) hypothesize that the concentration of acetaminophen which would be needed in the drinking water to provide analgesia would make the water unpalatable and thus would decrease the amount of water consumed, and thus would decrease the amount of drug consumed, leading to insufficient analgesia. This is a concern as water consumption is generally decreased after surgery. Cooper, et al., (1997), and Flecknell (1984) recommend the use of Buprenorphine for pain management as its effects are long lasting. Lidocaine can be used as a form of analgesia (it is a topical anesthetic, which acts by blocking voltage gated sodium channels, and thus prevents action potentials). A 5% (0.5 g in 10 ml) solution in sterile saline should be used, and can be applied with a cotton swab, or syringe. It is recommended that it be injected prior at the surgical site prior to an incision. As well, it can be applied prior to suturing a wound. Another topical anesthetic which could be used is Benzocaine (20% solution).

Antibiotics

“An ounce of prevention is worth a pound of cure.” Clean, aseptic surgeries are the best way to prevent infection. If a procedure appears to be infection prone, antibiotics can and should be given. Baytril is a common broad spectrum veterinary antibiotic. It can be given S.C. 12 hours prior to surgery, and again 12 hours after surgery (I.P. delivery is not recommended, as this could lead to rapid metabolism by the liver, decreasing the duration of action). This pretreatment prevents any bacteria from colonizing the wound. A dose of 5 mg/kg is recommended for rodents. A good topical antibiotic for use on wounds is the gel form of chloramphenicol, which can be used in conjunction with Baytril. A small amount can be placed on a cotton swab, and then be applied to the wound. For long term treatment, tetracycline can be used. Tetracycline is supplied in a drink crystal form in different concentrations. Treatment lasts 7-10 days. Fresh tetracycline drinking water should be supplied at least every second day.

Anesthesiology

Anesthesia is defined as a reversible, drug-induced loss of awareness and sensation. There are many choices when it comes to anesthesia. Each technique has its own benefits and disadvantages. Your choice of anesthetic will depend on the type of surgery and the species being used. All surgeries must be performed under complete general anesthesia. Under no circumstance should an animal ever have the conscious experience of surgery.

Assessing depth of anesthesia. There are four stages of anesthesia (Trevor and Miller, 1992). The first is Analgesia, where the subject experiences analgesia and followed by amnesia. The next stage is Excitement. Excitement is characterized by delirious behavior. Struggling, incontinence and vomiting can occur, although this is not a concern in rats, which cannot vomit. The reason for the appearance of this stage is that cortical function is depressed, and thus refinement of motor control no longer exists. Amnesia is experienced in this stage. The third stage is Surgical Anesthesia, which is the target stage for our purposes. It is characterized by the loss of many reflexes (described below). However, anesthesia does not mean absence of all movement, simply the lack of awareness and sensation. For example, the corneal reflex is maintained. The fourth stage is Medullary depression, which is characterized by the loss of the corneal reflex and the cessation of respiration, followed soon after by death. When assessing the reflexes, it is best to hold the animal in you hand. This will allow you to feel any response by the postural skeletal muscles.

Tail flick Pinching the tip if the tail elicits a response in the animal, which may be withdrawal of the tail or movement of the limbs and torso. Eyelash reflex Tickling the anterior corner of the eye elicits an eye blink. Pedal reflex Pinching the toe or foot elicits a response in the limb being pinched. The animal will withdraw the foot and may curve its back to better withdraw the limb. Ear reflex The animal will shake its head when the ear is pinched with your fingernail. I find this the most conservative test, as responses in the first three reflexes may be absent when this one is maintained. Complete absence is not necessary, but it should be substantially depressed. N.B., the ear can be moved merely by the shaking of you hand while you pinch. It is important to distinguish between the reflex and movement caused by touching the ear. Corneal reflex When the cornea is swabbed with a saline soaked cotton swab, the animal will blink its eye. This response should never be absent unless the animal has been too deeply anesthetized. However, it is not necessary to check this reflex. While its presence may be comforting to the surgeon, it may cause trauma to the cornea. Furthermore, there is often little that can be done if it is absent.

Common anesthetics Sodium Pentobarbital (Somnotol) This barbiturate was probably the most commonly used anesthetic for rodent surgery, but has fallen out of favor due to the fact that proper dosing can be challenging, leading to frequent overdoses. When used properly, it induces deep anesthesia. It is easy to deliver (i.p.), acts quite quickly (5-10 min), and anesthesia lasts for about 60 minutes. It acts on the GABAA receptor, which inhibits neural activity. The reticular system, which maintains consciousness, is turned off, and thus a deep state of anesthesia is reached. This system controls many important functions, such as respiration, and thus it is possible to overdose an animal. The major disadvantage is its EXTREMELY narrow therapeutic index. That is, the effective dose is quite close to the lethal dose. Care must be taken to carefully weigh the animal prior to delivery of the aesthetic, as well as to carefully fill the syringe. A dose of 40mg/kg is recommended for rats (CCAC, 1993), although a survey of the published literature suggests that this may be insufficient, and doses of ~65mg/kg may be needed. For hamster, a dose of 90mg/kg is recommended (CCAC, 1993), but again, higher doses are often used in the literature (100-120mg/kg) for some stains (i.e., LVG hamsters require more sodium pentobarbital than the HSD hamsters). Given high individual variability, care should be taken when using this drug, and it is better to underestimate the dose needed and then titrate up to surgical anesthesia. For some animals, the dose provided will not be sufficient to achieve anesthesia. This is due to individual variability. A supplement of ~10mg/kg can be given 10-15 minutes after the initial dose. A second supplementary dose can be give after another 10-15 minutes if anesthesia is insufficient. More than two supplementary doses of Somnotol should be avoided is possible, as it is possible that one or more of your injections was placed in a tissue which from which the Somnotol would be absorbed slowly, and hence a large quantity could be present, despite the lack of anesthesia. This could lead to an unexpected overdose. Commonsense should dictate whether or not a supplementary dose is necessary. A topical anesthetic or gaseous anesthetic may be sufficient if the animal regains a degree of consciousness near the end of the surgery. S.K. Wixson and colleagues (1987a,b) evaluated pentobarbital, ketamine and Fentanyl-droperidol (Innovar-Vet) for their respective advantages and disadvantages. Pentobarbital was found in their study to have many undesirable properties, including total lack of analgesia during sedation. Although they had no animals overdose from pentobarbital, they ranked it second worst for mortality, behind Innovar-vet that had a 6% mortality rate at one dose! They also ranked pentobarbital second worst for depth of anesthesia, a finding likely due to the fact that they used a dose of 40 mg/kg, far below the more typical dose of 65mg/kg. This low dose may also have been responsible for the amount of responsiveness to the noxious stimuli.

Ketamine Ketamine is a dissociative anesthetic that is similar to PCP. It interferes with the excitatory effects of glutamate at the NMDA receptor. Being quite lipophilic, ketamine reaches the CNS quite quickly and anesthesia is obtained within 5 minutes. Ketamine is a wonderful anesthetic that has a large therapeutic index. It is very difficult to overdose an animal with this anesthetic, as it actually stimulates the cardiovascular system and does not depress respiration. Often ketamine is co-administered with a tranquilizer such as Xylazine. For rats a dose of 90mg/kg Ketamine and 5-10mg/kg Xylazine is recommended (CCAC, 1993). Anesthesia lasts for about 60 minutes. For hamsters, some labs have found success with doses of 120 mg/kg Ketamine, 20mg/kg Xylazine, and 0.02mg/kg Acepromazine, although the effects are quite variable in this species.

Isoflurane Isoflurane is a gaseous anesthetic. It is a good anesthetic for short surgeries, or maintenance of anesthesia induced by injectable anesthetics. Simple procedures such as castration, ovariectomy, or abdominal implants may be most easily performed this way. The animal can be placed in an induction chamber and Isoflurane can be introduced or, alternatively, a low dose of Somnotol can be used to induce the animals prior to introduction of Isoflurane. It should only be used in a well ventilated room, and/or with a proper scavenging device. For induction, 2-5% Isoflurane mixed with oxygen is appropriate, while 0.5-2.5% is appropriate for maintenance. Recovery is extremely rapid if isoflurane is used alone. This makes it quite a desirable anesthetic for simple procedures. There is little risk of overdose with isoflurane when used this way. An active draw anesthetic machine should be used for rodents. The flow rate of oxygen should be set at 1 liter/min, while the amount of isoflurane can be controlled by the knob on top of the vaporizer. Although a single dose of an injectable anesthetic is typically sufficient for anesthesia in rodents for surgeries lasting up to 60 minutes, Isoflurane is a good emergency anesthetic should the animal emerge from anesthesia near the end of the surgery. It is undesirable to give extra injectable anesthetics near the end of a surgery, given that an animal can overdose easily, and this will prolong recovery. Isoflurane is a simple and quick alternative.

General Techniques

Aseptic surgery and the sterile field Minimizing the risk of infection required the proper implementation of aseptic technique and the maintenance of a sterile field during the surgery. Anything in surgical area can harbor microorganisms, and should be cleaned with a disinfectant or sterilized with an autoclave. Anything not sterilized should be considered non-sterile. Two major sources of microorganisms during the surgery are the surgeon and the animal. Since neither of you can be autoclaved, disinfectants and sterile barriers should be used. It is important to be aware that anything that enters a wound should be sterile, and that anything this object touches from the moment it is sterilized until the completion of the surgery should also be sterile. Anticipate what you will need to handle during the surgery, and make sure that it is sterile before you touch it. For instance, while surgical blades and syringe needles inside their packaging are sterile, the packaging itself is not, and therefore handling the packaging would contaminate you. For stereotaxic surgery, the sterile field can be established after the animal is in the frame, as manipulating the animal into place requires handling its body. After the animal is in place, wash your hands, put on sterile gowns, masks, hairnets and gloves. Place sterile drapes over the animal except for the area to be incised. At this point, only touch sterile items, or surfaces that have been disinfected. Have a sterile platform/area for surgical tools.

Preparing the animal. Once the animal is anaesthetized, the area of the body which will be incised must be prepared. Small animal clippers should be used to shave the area of interest. Care should be taken near sensitive tissue (i.e., ears, eyes and vibrissa). The following tips may be useful:

Hold the stationary part of the clipper against the skin. Holding the moving cutting blade against the skin will cause abrasions.
Hold the clippers tilted forward at about a 45° angle.
Hold the skin taut, especially for hamsters, which have loose skin.
Perform the shaving away from the surgical field. After you have finished clipping, take the animal to the surgical area and disinfect the area. Use a small gauze pad or cotton swab soaked in a disinfectant (Xenodine, Germaphine, 70% alcohol) to clean the area. Start in the center of the area and work your way outwards, scrubbing firmly. A cotton swab works well here, as you can rotate the shaft between your fingers such that the swab is rolling away from you, as you draw the whole thing towards you. This really cleans all the dirt, oils and hair clippings from the area.
If the surgery is prolonged (as it is during stereotaxic surgery) the eyes should be covered with ophthalmic ointment, mineral oil or Vaseline to prevent them from drying out.

Incising Using a 25 gauge needle, inject a small amount of lidocaine (2-5%) along the incision line. Allow about a minute for this to take effect. For making the incision, a #10 blade would be most appropriate for most small animal surgery. A #11 blade may be useful for abdominal surgery, but the cutting technique would be different than that described here (although the present technique would work for abdominal surgery as well). To make an incision, hold the scalpel like a pencil. Pull the skin taut and press the scalpel through the skin (a slight rocking motion of the blade may help). When you are through the epidermis, the skin can be grasped with your free hand immediately adjacent to either side of the blade. Draw these fingers away from the cutting edge, pulling them apart as you go, in a “Y” like motion, where the blade is at the intersection of the “Y” and the path of your fingers starts at the intersection and follows either arm of the “Y”. The stem of the “Y” represents the path the incision will follow. This technique is a little tricky, and a simple incision where the blade is drawn towards you will suffice. For this technique, it is best to use the ring finger on the hand holding the scalpel, and the index finger on the non-dominant hand to hold the skin taut as the blade is drawn towards you. The incision should be performed in a single smooth motion so that the incision has a smooth edge.

Suturing There are various options available to the surgeon when it comes to close the wound, each option having advantages and disadvantages in various situations. The classic technique involves the use of a suture needle and thread, although metal wound clips and super-glue are becoming more popular. There are many types of needles available, in different lengths, diameters and curvatures. They differ also in how they cut skin. Atraumatic needles have a point which resembles a classic sewing needle and have a circular cross-section, whereas the traumatic needle has a triangle cross-section with cutting edges along the shaft of the needle in addition to its point. The atraumatic needle is used for soft tissue (e.g., the intestinal wall), while the traumatic needle is used for tough tissue (e.g., skin). Needles are either supplied with an eye through which to thread the suture material, or with pre-bonded suture material. The advantage of the pre-bonded suture material is that the hole created in the tissue is smaller than with the treaded needle. Suture material comes in various thicknesses. Size is indicated by a number, with a smaller initial number indicating a thicker suture material. 3-0 to 5-0 is a good size for rodents. There are various types of material which can be used. The synthetic absorbable suture material is known as vicryl. This material should be used for sutures which are to left inside the animal, such as suturing the peritoneal wall. Non-absorbable sutures are either made of silk, or some synthetic material such as polypropylene (e.g., Prolene), or nylon (e.g., Ethilon). Silk comes as either twisted or braided. The twisted silk appears to be the most supple of all the non-absorbable suture material, although unbraided seems better for wound healing. Generally, suture material is left in place in experimental animals, unless infection occurs. The animal will eventually groom out the sutures, but the wound would be healed by the time this occurred. If the sutures are irritating to the animal, they should be removed as soon as the wound is healed. There are many different types of suture styles. One thing which they all should have in common is being closed with a square knot (also known as a wreath-knot). To tie one by hand, you would hold a thread in each hand, pass the left thread over the right one, and twist it through the loop formed (completing on half-hitch), then pass the thread which is now on your right over the left and twist it through the loop formed (completing a second half-hitch). You will be left with the threads on each side passing out of the loop side by side. A third half-hitch can be made by repeating the first step. This will strengthen the knot. This is not how we will tie the knots, but you can do this to understand how the thread will lie. Ideally, this technique should be completed without touching the needle or thread with your hands, but when learning, the animal will appreciate speed rather than “proper technique”, provided that you are fairly clean when doing the suturing. It takes practice to master the proper technique, so don’t expect to master it on your first try. To make a simple knot, you should grasp the threaded needle by the shaft with a pair of hemostats or needle drivers. Make sure that they are locked closed, as this will make it easier for you to manipulate the needle, as you won’t have to worry about maintaining pressure on the needle while moving it. If the needle is curved, have the curve pointed upward. Using forceps, grasp either one or both edges of the wound to hold it still while you use a flick of the wrist to force the suture needle through one or both layers. Doing both edges of the wound at the same time is faster, but you will have greater control over where the needle passes if you do one layer at a time. In some cases it is simply too hard to do both at the same time. Ideally you’ll want a suture every 1½ to 2 mm along the wound, with the thread passing through the skin about 1-2 mm from the incision edge. One the needle is through both edges of the incision, grasp it with another pair of hemostats or needle drivers, and disengage the first pair. Draw the needle and suture material through the skin until only about 4 to 5 cm of thread remain on the side of entry. If the needle is a threaded rather than a bonded kind, clamp both the thread entering and leaving the eye with the needle drivers. If you were to hold the needle, any attempt to manipulate the suture material would result in you simply sliding the needle off of the suture material. The suture material should now be head in hemostats in your non-dominant hand. Hold the other pair of hemostats in your dominant hand horizontal over the animal, perpendicular to the wound. Pass the suture material from proximal edge of the free-hemostats over top of it down and around the distal edge. Then use the free hemostats to grasp the free end of the suture material, pass the loop of suture material off of the end of the hemostats and cross your arms to tighten the knot. Let go of the short end of the suture material, and hold the free hemostats (located in you dominant hand) horizontal gain, and this time reverse the direction of the suture material, starting with the distal edge, over top, and under the proximal edge. Again grasp the free end of the suture material, and pass the loop over the end of the hemostats. This time, pull your hands apart, each to its respective side. Repeat the initial step to create a third throw, thereby strengthening the knot. The excess suture material should be cut down to 3-5 mm in length. Too long and there will be too much for the animal to catch and rip, while too little could lead to the knot unraveling. Alternatives exist to this simple knot. A continuous stitch is completed more rapidly, but is inappropriate if an implanted cannula is emerging from the skin. One major disadvantage of the continuous stitch is that if it is broken or torn, the whole stitch will unravel, while an interrupted stitch would only lose the stitch that was damaged. In some cases, sutures are not appropriate, and wound clips should be used. Rodents will chew on any visible thread and can easily re-open a fresh wound. This is an issue for many location, such as abdominal surgery. In this case, wound clips would be best. The two edges of an incision are held together with special forceps and the clip is applied with a special tool that bends the metal and causes it to bite into the tissue. These clips should be removed (again, with a special tool) after one week. For mice, thread or clips may cause too much tissue damage. Their skin is very thin and fragile. Cyanoacrylic glue is the most efficient option. The two edges of the incision are brought close together and a thin bead of glue is run along the wound. The glue dries quite quickly, and care must be taken not to glue forceps to the animal. In addition, glue should be used sparingly to prevent it from running into the eyes, ears or other vital areas.

Post operative care A quiet area should be available for recovery. This area should be warm, and easily accessible to watch the animal. Depending on how invasive the surgery, you may want to leave the animal in recovery for a couple days so that you can monitor for infection and quality of recovery. An animal emerging from an anesthetic will have an impaired ability to thermoregulate due to loss of muscle tone. There are many ways to keep the animal warm. We typically place its recovery cage near a heat lamp during surgery so that it is already warm when the animal is placed in the cage. Rather than leaving the directly under the heat lamp when the animal is recovery, which may lead to the animal overheating, the cage is placed to one side, such that it is kept warm. Wrapping the animal in paper towel, like a burrito, will help it remain warm, much like a blanket would keep you warm at night. Place some food pellets into the recovery cage so that the animal does not have to stand to eat, a task that can be difficult while under the influence of a barbiturate or opiate. For surgeries that are very invasive, a palatable, high calorie mash can be provided. Use equal parts sucrose and ground rodent chow, mixed with vegetable oil until it has the consistency of cookie dough. Animals should be monitored regularly (hourly) until they are conscious, and then periodically (daily) until they are fully recovered. Analgesics should be given based on the dosing regimen for the specific drug that is being given, typically every 6-12 hours until recovery is complete. Animals should be monitored for any sign of pain (decreased appetite, lethargy, aggressiveness) or infection. Confirmation of urination and defecation should also be observed.