Common surgical procedures in rodents

Common Surgical Procedures in Rodents P. L. Foley Office of Animal Research Education and Compliance, University of Virginia, Charlottesville, VA, USA. In: Laboratory Animal Medicine and Management, Reuter J.D. and Suckow M.A. (Eds.). International Veterinary Information Service, Ithaca NY (www.ivis.org), Last updated: 1-Jun-2005; B2515.0605 Table of Contents • Introduction • Adrenalectomy • Bile Duct Catheterization • Cesarean Rederivation • Jugular and Carotid Catheterization • Ovariectomy • Pinealectomy • Portal Vein Catheterization in Rats • Skin Grafting • Vasectomy or Orchiectomy • Special Considerations for Embryo Implantation Surgery in Mice Introduction This chapter describes surgical procedures commonly performed in rodents in biomedical research. The previous chapter, Principles of Aseptic Rodent Survival Surgery, provided information necessary to performing surgery successfully, such as aseptic technique, pre-, peri-, and post- operative care concerns, and anesthesia. Therefore these details are not provided in this chapter, which focuses on the actual surgical procedures. Nonetheless, careful attention to sterility, thorough understanding of appropriate methods of anesthesia in rodents, and an understanding of concerns specific to care of rodents during and after surgery are imperative to the success of these techniques. It is also critical that the surgeons have experience in the use of surgical instruments and basic surgical technique. Practice with suturing techniques and anatomical dissections should be performed on inanimate objects and postmortem animals respectively prior to beginning experimentation on live animals. In most cases, the techniques in this chapter are described as they would be performed in a rat. When the differences in technique for mice are significant, such as catheter sizes, these are discussed within the relevant section. Surgical procedures are listed in alphabetical order. The final section focuses on special considerations for surgery in generation of genetically engineered mice. This section was contributed by Patricia A. Brown and Shelley Hoogstraten-Miller. Adrenalectomy The adrenal glands are small, pink organs located near the anterior pole of each kidney. The anesthetized rat should be placed in ventral recumbency and an area on the mid-dorsum (thoracolumbar junction) shaved and prepared for aseptic surgery. A midline incision 1 - 2 cm long is made just caudal to the peak of the animal’s dorsal hump (Fig. 1). Hemostats or blunt-tipped scissors are inserted subcutaneously through the incision and used to bluntly dissect the connective tissue down both sides a short distance (about 1/3 of the distance down the lateral abdominal wall). The skin incision is then pulled laterally to one side to expose the muscle just caudal to the last rib and a small incision made in the muscle to enter the peritoneal cavity; this incision is just large enough to retrieve the adrenal gland back through the incision. On the left side, the spleen should be visible directly underneath or slightly to the left of the incision (Fig. 2). Curved forceps are inserted through the incision into the peritoneal cavity and used to hold the spleen laterally. A second pair of forceps is used to move the incision over the gland, which is usually surrounded by adipose tissue just in front of the kidney. Once the gland is located, it is manipulated by grasping periadrenal fat and exteriorized (Fig. 3). This may require freeing up some of the fascial connections between the kidney and adrenal gland. Care should be taken not to grasp the gland itself because tearing of the gland may result in leaving functional residual tissue within the abdomen. Clamp the vessels at the base of the adrenal gland with both forceps. The forceps are then used to tear away the gland and its surrounding tissue. The tissue stump is then returned to the abdomen. No hemostasis is necessary as bleeding should be minimal. On the right side, an incision is made in the abdominal muscle similar to the left side, but the liver must be moved cranially to view and retrieve the adrenal gland. Closure of the abdominal muscle may be necessary if the incision is large (greater than 3 or 4 mm), such as in larger rats. If needed, one should use a single interrupted suture with 4-0 absorbable suture. The skin incision should be closed with either a wound clip or using 1-2 simple interrupted sutures with a non-absorbable suture. Figure 1. Location of incision for adrenalectomy on dorsum of a rat Figure 2. A large incision is present for demonstration purposes to illustrate the location of the left adrenal gland in relation to the spleen and kidney. The arrow points to the adrenal gland nestled in adipose tissue just cranial to the anterior pole of the kidney. Figure 3. Exteriorized adrenal gland. Post-operatively, replacement of mineralocorticoids and corticosteroid is necessary to maintain homeostasis. A standard 0.9% saline or a saline/dextrose solution (0.9% saline and 10 g/L dextrose) can be administered instead of drinking water. Corticosteroid replacement can be achieved with hydrocortisone acetate (0.1 mg twice daily by subcutaneous injection in rats). Bile Duct Catheterization The bile duct runs from the hilum of the liver through the pancreatic tissue to the duodenum, where it is surrounded by the muscular sphincter of Oddi. The duct is approximately 1 mm wide in the rat and moderately translucent. There is no gall bladder in the rat. Substances excreted by the liver into the bile duct may be reabsorbed in the gut, and carried back to the liver by the portal vein as part of the enterohepatic circulation. A dissecting microscope can be useful for successful catheterization of the bile duct. The anesthetized rat is placed in dorsal recumbency, prepared for aseptic surgery, and a midline abdominal incision is made starting at the xiphoid cartilage and extending caudally about halfway down the abdomen (approximately 4 cm long in a rat). A technique, along with excellent diagrams, is described in Waynforth et al., [1]. This technique involves arching the back up using rolled gauze, to facilitate visualization of the bile duct. For exteriorization of the catheter and chronic access, a needle with an interior diameter just large enough to slide the catheter through is passed through the skin at the side and towards the back of the rat, and into the abdominal cavity taking care not to puncture the gut. The needle is used to exteriorize the catheter. The duodenum and a small part of the intestine is pulled out and placed on a saline-moistened gauze pad. The course of the bile duct through the pancreatic tissue should now be visible (Fig. 4). The major lobes of the liver must be moved cranially against the diaphragm with a retractor or with a moistened gauze pad. One can identify the portion of the bile duct near the hilum of the liver. In order to collect pure bile without contamination of pancreatic juices, the duct should be catheterized near the hilum. Forceps are used to carefully clear the bile duct of surrounding connective tissue for a distance of approximately 1 cm. The surgeon can then pass a doubled-up length of suture under the bile duct and cut to create two separate threads. One suture is tied to create a ligature just cranial to the pancreatic tissue to obstruct bile flow. A single hitch should be thrown with the second suture strand about 0.5 cm cranial to the first ligature, closer to the liver (Fig. 4). Figure 4. Bile duct near the hilum of the liver, with ligatures pre-placed prior to cannulation. A polyethylene (PE-10) or silicone catheter, 0.28 mm inside diameter (ID) and 0.61 mm outside diameter (OD), is cut so that a short beveled point is obtained. The bile duct near to the first ligature is held with fine forceps and partially transected with micro-scissors or punctured with a 23 gauge needle. The catheter is introduced into the duct towards the liver and advanced until several mm past the cranial suture. This suture is tightened and the knot completed to secure the catheter in the bile duct. One can then tie another throw around the catheter with the caudal suture. Bile should be visible within the lumen of the catheter. The catheter is then exteriorized by passing it through the end of the needle and passing the needle back out of the animal’s skin, bringing the bile duct catheter with it. The catheter should lie in the abdominal cavity with some free play to allow free body movement once the animal regains consciousness, and without twisting or kinking. The liver and intestines are then returned to the abdomen. The flow of bile in the catheter is confirmed before suturing the catheter to the skin. The catheter can be secured to the skin by use of tape placed around the catheter, which is then sutured to the skin using simple interrupted sutures, or by use of a silicone disc/button which is slid over the catheter and sutured to the underside of the skin. The abdominal incision is closed in routine fashion. If the catheter is to be used for acute studies only, then the rat can be restrained in a "Bollman-type" restrainer or similar device during the collection period when the animal has recovered from anesthesia. For chronic collection studies, the catheter will need to be protected, either by use of a tethering device and harness or by attachment of the catheter to a vascular access port and placement of the port in a subcutaneous pocket on the dorsum. The difficulty with chronic collection studies is that gravity significantly aids in flow, and the collection system must take this into consideration. Cesarean Rederivation Cesarean rederivation is often used to obtain pathogen-free mouse or rat pups from a mother that is either infected with a known pathogenic agent or is of unknown health status. In this situation, extra precautions must be taken to ensure that cross-contamination does not occur. These include dipping the mother in disinfectant, such as dilute iodine, immediately after euthanasia, and using a second set of sterile instruments for opening the uterus from those used to open the skin. If survival of the dam is not necessary, it is best to euthanize it by cervical dislocation without any anesthesia to avoid cardiovascular and respiratory depression of the pups. This procedure should be performed as close as possible to the expected time of parturition to increase the likelihood of producing viable pups. For mice, this should be performed on Day 19 or Day 20 (with day of vaginal plug observation being Day 0), depending on the strain. For rats, Day 20 is typically used. It is also important to have suitable foster mothers available. The foster mothers should: • Ideally be of a stock or strain that typically has strong maternal instincts. Most outbred stocks do well. With regard to inbred strains, BALB/c mice are typically good mothers. • Foster mothers should have newborn litters preferably no more than 2 - 3 days old at the time of rederivation. • Using a foster mother with pups of different coat color from those pups being rederived will allow the two sets of pups to be easily distinguished once they are mixed together. The euthanized or anesthetized pregnant animal is placed in dorsal recumbency on a sterile surface, and a long ventral midline incision made from the xiphoid process to the pubis. Care should be taken when opening the abdominal wall to not accidentally enter one of the uterine horns, as these are often lying immediately dorsal to the abdominal musculature. One horn should be exteriorized and placed on gauze soaked with warm saline. The horn is carefully opened along its entire length using scissors, on the side opposite the placental discs. Working with one pup at a time, forceps should be used to ligate the umbilical blood vessels between the placental disc and the mother’s uterus. The fetus is then pulled away with its amniotic sac and placenta still attached. The pups should be handed to an assistant who can then remove the amniotic sacs and provide postpartum care to the pups. After all pups are removed from one horn, the forceps can be removed. If the blood vessels are large (e.g., in rats in advanced pregnancy) they should be ligated prior to removal of forceps; otherwise, gentle pressure with gauze or a cotton-tipped applicator will provide sufficient hemostasis. Once the pups have been removed from one horn, one should quickly proceed to the other horn and repeat the procedure for removing the pups. If the dam is intended to survive from this procedure, the abdominal muscle can be closed in standard fashion with simple interrupted sutures using an absorbable suture, and the skin closed with non-absorbable suture or staples. An audiovisual step-by-step instructional guideline of this procedure in mice is available online. For non-survival procedures, an alternative method is to make a cut across the caudal vagina and dissect out the entire uterus from the mesentery. The entire uterus is then placed in a dish with disinfectant, then removed onto a sterile absorbent surface for removal of the fetuses. The pups should be gently dried and stimulated until they are breathing well and gain a healthy pink color indicating good tissue oxygenation. Gently squeezing the tail or a paw, or providing oral stimulation using a cotton tipped applicator should evoke a response from a healthy full-term pup. The pups should be transferred to the foster mother as soon as possible. Removal of the natural litter from the foster mother shortly before the procedure, and gently rubbing the two litters together along with some of the nesting material from the home cage should provide sufficient olfactory masking of the new pups to avoid rejection. Depending on the total number of pups in the two litters, some of the natural pups may need to be removed. A total of 8 to 12 pups will stimulate good milk production without overburdening the mother. Jugular and Carotid Catheterization Vascular access is often needed to either infuse a substance, obtain repeated blood samples, or to monitor blood pressure. This may be done as an acute procedure where the experiment takes place over the period of one or several hours and the animal is euthanized while under anesthesia, or as chronic cannulation to allow for vascular access over a prolonged period of time (days to weeks). Chronic catheterization requires that utmost attention be paid to sterility and the prevention of thrombus formation which obstructs cannula flow. Infections are the leading cause of early failure for indwelling catheters. The type of catheter used also has significant implications on biocompatibility and long-term patency. The most common materials used are polyurethane and silicone. Polyurethane or polyethylene both have good rigidity which makes vessel entry easier, but may induce more inflammatory response from the luminal surface. Silicone is more flexible and has been shown to be more resistant to infection and cause less tissue trauma within the vessel but is harder to work with. Newer catheters, such as Renathanetm (Braintree Scientific Inc. Braintree, MA, USA), Hydrocathtm (BD Medical, Franklin Lakes, NJ, USA) and CBAStm-coated catheters (Carmeda AB, Stockholm, Sweden) have anti-thrombotic coatings and seem to also resist infections better than standard non-coated catheters. Use of retention beads is also recommended. These serve as anchoring points for the catheter in the tissue, and can be "home-made" using Silastic glue, or purchased with the catheter as either movable or fixed rings on the catheter. There are numerous references available addressing catheter properties, effective flushing solutions, and ways to minimize the complications of infection and catheter obstruction, so this topic will not be addressed in any detail here. For all vascular catheterizations, the catheter should be pre-filled with heparinized saline solution (50 IU/mL), and is usually attached via a 23G needle to a syringe similarly filled with no air bubbles. For rat jugular veins, a 3Fr (OD 0.94 mm; ID 0.51 mm) catheter works well. In mice, catheters are either tapered to a small diameter or the smaller intravascular portion (1.2Fr; OD 0.41 mm, ID 0.23 mm) of the catheter is inserted into a larger diameter tubing (3Fr) for the extravascular portion. Venous access can be achieved via the jugular vein, femoral vein, or tail vein. Similarly, arterial access can be obtained by catheterizing either the carotid or femoral artery. The techniques are largely similar and differ primarily in approach and method of securing the catheter outside the animal. Catheterization of the Rat Jugular Vein The anesthetized rat is placed in dorsal recumbency with its head towards the surgeon. The right external jugular vein is most often used as it feeds directly into the right atrium.The tip of the catheter should ultimately lie at the junction of the precava and right atrium. An incision approximately 1.5 cm long is made in the skin on the right ventrolateral aspect of the neck above the clavicle. In mice, the procedure at this point may be more easily performed using a dissecting microscope or loupes. The right external jugular vein is dissected free of surrounding fascia with the caudal most landmark being where the vein courses underneath the pectoral muscle (Fig. 5). The bifurcation into the maxillary vein and the linguofacial vein may be visible rostrally. Once there is a cleared section of vein (5 - 10 mm) it can be stabilized by passing two loops of 4-0 silk suture underneath the vein. This is most simply done by passing a small forceps underneath the vein, grasping a doubled piece of suture in the teeth of the forceps, then drawing the suture back underneath with the forceps. The suture is then cut into two pieces, one of which is moved to the most cranial (anterior ligature) aspect and the other moved caudally (posterior ligature). After filling that portion of the vein with blood, the anterior ligature is tied tightly to occlude blood flow returning to the heart. The posterior ligature is tied with a single loose half-hitch and can be used to create tension on the vessel during venotomy and catheter insertion (Fig. 6). Some surgeons like to place a flat instrument or metal strip underneath the vein during venotomy to provide stabilization and a rigid surface against which to catheterize. A venotomy is made posterior to the anterior ligature, using either fine iris scissors, micro-scissors, or a needle tip. The venotomy can be widened by inserting the tips of Dumont #5 forceps and spreading gently. Figure 5. Jugular vein visible after blunt dissection of surrounding tissue. Figure 6. Jugular vein with two sutures in place. The anterior suture is tied to occlude blood flow. Angled Dumont forceps can be used to keep the venotomy site open while the catheter tip is introduced and advanced towards the heart using a fine pair of forceps to grasp the catheter and push it gently forward. A commercial catheter introducer can also be used for this procedure. The distance that the catheter must be advanced to place the tip in the right atrium will depend on the size of the animal and should be pre-determined in a pilot study. For infusion studies, a shorter distance of insertion is adequate, as long as good blood flow is obtained at the site of placement. Typically, for a rat weighing 250 - 300 g, a distance of 32 mm (starting 3 mm cranial to the pectoralis major muscle) will place the tip at the entrance to the right atrium. In mice, th