Facilitating Repeat Intracarotid Injections in Mouse Models by a Novel Injection Site Repair Technique

All the steps outlined below are in compliance with our protocol, which follows guidelines established by and was approved by the Institutional Animal Care and Use Committee at The University of Texas MD Anderson Cancer Center. 1. Preparation of the surgical table and mouse for surgical procedure Preparation of the surgical table and mouse for surgical procedure (refer to Figure 1A,B) Arrange the dissecting microscope and light source in front of the isoflurane vaporizer with an active scavenging system or place on a downdraft table. Place the electric heating pad on the dissecting microscope base (or preferably under the microscope if the base is solid and can transmit heat) and cover with a sterile surgical drape. NOTE: All surgical tools and drapes should be sterilized by autoclaving prior to use and all disposables should be sterile and individually packaged. Sterile gloves should be used throughout the setup and procedure and changed when necessary to maintain sterility. Prepare the bed (4-5" long x 2" wide) on the surgical drape using vinyl lab tape. Cut a 1.5 inches x 1.5 inches square of gauze, roll it tightly, and place the rolled gauze under a 3 inch piece of vinyl tape at the "head" of the bed to form a pillow (head tilt with the use of a pillow allows for greater extension of ventral neck area). Lay 4 inch pieces of surgical tape along the sides of the bed (on top of the vinyl tape). [If using isoflurane, place an anesthesia nose cone with weight (or tape) attached near the head of the bed. Adjust it for specific placement once the mouse is anesthetized and restrained.] See Figure 1A,B. Using a needle driver or heavy forceps, tear off small pieces of cotton from a cotton swab and roll into balls of various sizes from 0.5 to 1 mm in diameter (8-10 cotton pieces per mouse). Keep the cotton on the surgical drape near the head of the bed. Using sterile instruments, cut 6-0 suture thread into 1 cm pieces (3-4 pieces per mouse). Place the prepared suture lengths on a sterile drape. Prepare a 1 mL syringe for administration of buprenorphine or other suitable analgesic (as approved by the Institutional Animal Care and Use Committee [IACUC] protocol). Sterilize all surgical tools according to IACUC standards prior to placing the tools in the sterile field. Preparation of mouse for the surgical procedure (refer to Figure 1C-F) Anesthetize one mouse following methods approved by IACUC for major survival surgeries (use either isoflurane 1%-4%, based on the sensitivity of individual mice, or a cocktail of 10 mg/mL ketamine, 1 mg/mL xylazine at 100-200 mg/kg (ketamine) of body weight). Clip the fur or depilate if necessary. Administer 0.5-1.0 mg/kg buprenorphine ER (Extended-release) by subcutaneous injection 30 min prior to the start of the surgical procedure. Position the anesthetized mouse so the pillow rests under the neck (Figure 1C). The pillow helps to extend and support the neck when used with the anesthesia nose cone. If using the ketamine/xylazine cocktail, place a weighted tooth bar or similar tool in the mouth, behind the incisors to tilt the head and extend the neck. Restrain the forelimbs using the surgical tape previously placed along the sides of the surgical bed (Figure 1D). Adjust the position of the mouse under the microscope so that the ventral surface of the neck is in view and adjust the magnification to comfortably observe the surgical site (Figure 1E). NOTE: The magnification of the dissecting microscope should be adjusted by the surgeon to their comfort level for each step. Apply artificial tears using a sterile cotton swab. Disinfect the surgical site by alternating circular swabs of betadine or chlorhexidine and alcohol three times per disinfectant (Figure 1F). Confirm the depth of anesthesia by ensuring the mouse does not retract its leg in response to a toe pinch. Monitor the respiratory rate and ensure that the mouse is not gasping, as this is an indication of excessive anesthesia when using isoflurane. If necessary, adjust the oxygen and isoflurane flow rates to reach appropriate anesthetic depth and even respiration. 2. Surgical procedure (Figure 2, Figure 3, Figure 4, Figure 5, Figure 6, and Figure 7) Primary incision and dissection Begin by making a 1 cm longitudinal, midline incision, starting just cranially to the manubrium (protruding lump at the cranial end of the sternum) and continuing over the trachea using a sterile scalpel blade or similar instrument following the user's approved IACUC guidelines (Figure 2A). Insert the tip of the closed scissors into the incision and gently open it to perform blunt dissection of the subcutaneous connective tissue, separating the two salivary glands. With the fine forceps, gently pull the right salivary gland through the incision to rest on the aseptically prepared skin surface or retract the salivary gland laterally using the blunt hook retractor (Figure 2B,C). If the exteriorized gland appears dry or tacky, moisten with sterile saline. Continue blunt dissection of the connective tissue until the sternocleidomastoid and digastric muscles are visible (Figure 2D). NOTE: The muscular triangle formed by the trachea/sternohyoid (sh) muscle, sternocleidomastoid (sm) muscle, and the digastric (dg) muscle (caudal belly) will be used to locate the right CCA and the carotid artery bifurcation in the protocol. In general, the smaller omohyoid (oh) muscle can also be seen lying transversely across the CCA (Figure 2D); however, this muscle varies in size and it is not uncommon for the omohyoid muscle to be absent altogether in young or small mice. CCA isolation Using the angled tip forceps, continue careful dissection of the connective tissue (by opening the tips of the forceps) near the caudal end of the muscular triangle to expose the common carotid artery, jugular vein, and vagus nerve. NOTE: The common carotid artery is the largest blood vessel adjacent to the trachea and can generally be easily identified at the base of the muscular triangle (Figure 2E, arrow). Be extremely careful with the fine forceps around the blood vessels as the tips can easily nick the vessels resulting in excessive and potentially lethal bleeding. Continue careful dissection of the connective tissue around the portion of the common carotid artery from the base of the muscular triangle up to the omohyoid muscle. Use the small, sterile cotton balls to control any minor bleeding and absorb secreted fluids from the salivary glands as needed. Carefully dissect away connective tissue to separate the CCA from the vagus nerve. Take special care to minimize handling of and damage to, the vagus nerve, easily identified as the thick, white, nerve bundle adjacent to the CCA (Figure 2F). CCA preparation Once the CCA has been completely mobilized from the sternocleidomastoid muscle, and where visible, up to the omohyoid muscle, place a 1 cm piece of 6-0 suture on the aseptically prepared skin of the mouse’ sternum (for easy retrieval) and pass the angled tip forceps under the CCA (taking care to isolate CCA from the vagus nerve and jugular vein) (Figure 3A). If the omohyoid muscle is not present, clear the area around the CCA sufficiently to be able to place sutures on the CCA, cranial and caudal to the proposed injection site, and to insert the needle. With the fine forceps in the left hand, pass the suture to the angled tip forceps, grasping near the end of the suture. Gently pull half of the length of the suture under the CCA with the angled tip forceps (Figure 3B). Repeat this process with a second suture, parallel to the first suture (Figure 3C). Loosely tie each suture around the CCA, but do not tighten the knots or restrict blood flow (Figure 3D). External carotid artery isolation and preparation Using the angled tip forceps, carefully remove connective tissue at the cranial end of the muscular triangle, cranial to the omohyoid muscle, to locate the CCA and the bifurcation into ECA and ICA (Figure 4A). NOTE: The ECA angles toward the midline and is slightly more superficial, while the ICA angles laterally and moves deeper into the neck. Take special care to prevent damage to the hypoglossal nerve (HN) crossing over the ICA just above the bifurcation. Carefully clear away connective tissue from all sides of the ECA near the bifurcation. Once enough connective tissue has been cleared away from the ECA, place a piece of suture on the aseptically prepared skin on the mouse’s sternum and pass the angled-tip forceps under the ECA. With the fine forceps in the left hand, pass the suture to the angled tip forceps in the space between the ICA and the ECA and gently pull half of the length of the suture through (Figure 4B). Loosely tie the suture around the ECA, but do not tighten the knot. NOTE: It is important that enough connective tissue be cleared from the ECA that you can grasp the suture with the angled tip forceps without also inadvertently grabbing connective tissue surrounding the arteries, causing damage to the arteries when the suture is retrieved. Needle and syringe preparation NOTE: For this step, the injection can be performed with a straight needle, which allows for the syringe to rest against and be stabilized by the body of the mouse (Figure 5A). Alternatively, the injection can be performed using a needle that is bent near the tip, allowing the syringe to be held like a pencil with the hand resting on the surgical table (Figure 5B). Both techniques work well, and the choice of technique is a personal preference. To prepare the bent needle, hold a 33 G, ½ inch needle with the bevel facing up and grasp the tip with a sterile needle driver (Figure 5C). Bend the needle approximately 30-40° directly toward the bevel (Figure 5D). Fill the syringe with the appropriate volume of the solution to be injected (be sure to account for the cavity space of the needle if the solution for injection is not drawn up through the needle to load the syringe). Attach the needle and remove any air bubbles. Ensure that the meniscus of the solution is visible at the bevel of the needle. Intracarotid injection Tighten the knot of the suture around the ECA. Next, slide the lower suture on the CCA down toward the sternocleidomastoid muscle as far as possible and tighten the knot. Ensure that the upper suture on the CCA remains loose until after the injection. Place a sterile cotton ball into the edge of the cavity to absorb secreted fluid and blood during the injection. Holding the syringe in the right hand and the fine forceps in the left hand, bring the needle to the artery immediately above the lower suture on the CCA. With the fine forceps, gently pull the loose end of the lower suture in a caudal direction to place a low level of tension on the CCA (Figure 6A). Insert the needle into the CCA just past the bevel and slowly release tension from the suture (Figure 6B). NOTE: The artery has substantial blood flow and will likely bleed into the surgical cavity as the needle is inserted. However, once the needle is inserted past the bevel, the artery will form a seal around the needle and the bleeding will stop. Continuous bleeding with the needle in place indicates that the needle is not far enough into the artery (the gap from the bevel allows blood to flow) or that the needle has been pushed through the back of the artery. It is important now to hold the needle extremely still to keep from tearing the artery or allowing the needle to slip out. Continuously monitor animals with hemorrhage events, and if bleeding is significant (such as when an animal loses coloration, appears cyanotic, becomes cool to the touch, or any other humane endpoint described in the animal use protocol), it is necessary to apply humane endpoints. Use the left hand to push the syringe plunger to inject the solution very slowly (Figure 5B, no less than 15 seconds to inject 100 µL of solution). Do not remove the needle when finished injecting the solution. To prevent backflow, with the fine forceps in the left hand, grasp the upper suture on the CCA (still loosely tied) by the knot and lift to kink the artery (Figure 6C). Remove the needle, set the syringe aside, and pick up angled-tip forceps with the right hand. Keeping the kink in the artery, tighten the knot in the upper suture on the CCA (Figure 6D). NOTE: At this point, there should be no additional bleeding into the surgical cavity. Following this step, there are two alternative procedures as described in steps 2.7 and 2.8 below. If the CCA is to be ligated, follow step 2.7. If the CCA is to be repaired at the injection site, follow the modification listed in step 2.8. Ligation of CCA In cases that do not require that circulation be restored to the CCA, leave the artery ligated, leaving both sutures tightened on the CCA. Trim the suture ends and confirm that the suture knots are fully tightened. Proceed to close and analgesia procedures described in step 2.9. Alternative to ligation of CCA-Injection site repair and restoring circulation Using sterile cotton balls, absorb any residual blood within the surgical cavity. Locate the injection site on the CCA (Figure 7A) and determine the number of sutures needed to close. Irrigate the injection site and lumen of the isolated area of the CCA thoroughly to remove coagulated blood. Using the angled-tip forceps, grasp the 9-0 suture needle by the needle body close to the swage. While using the fine forceps to support the artery from the opposite side, place a single suture in the CCA by penetrating the arterial wall approximately 1-1.5 mm lateral to the injection site, perpendicular to the artery. Hold the artery with the fine forceps, opening the injection site and passing the needle and suture through the right and left sides individually. Alternatively, hold the artery, gently pressing the sides together with the fine forceps, passing the needle, and suture through both sides of the artery with a single bite. NOTE: With either technique, be careful that the needle and suture do not penetrate the back wall inside the artery as this will close off the lumen when tightened. Close the injection site with a surgeon's knot performing an instrument tie with the fine forceps and angled-tip forceps, using a minimum of four throws (Figure 7B). In general, use a single simple interrupted suture to close the injection site of a 33 G needle. To restore circulation using the fine forceps, untie and remove the suture around the ECA, followed by the upper suture on the CCA (Figure 7C,D). Next, slowly loosen the lower suture on the CCA but do not immediately untie. Confirm that the injection site is closed sufficiently to prevent major bleeding with restored blood pressure and flow (Figure 7D). ​NOTE: If major bleeding occurs at the injection site, the lower suture on the CCA can quickly be re-tightened and the injection site can be adjusted or re-sutured if necessary. Remove the upper and lower CCA sutures (Figure 7E). Go to step 2.9. Closing and analgesia Reposition the salivary gland in the cavity and close the incision with three simple interrupted sutures using a sterile suture pack. Remove surgical tape restraints, and allow the mouse to recover from anesthesia on a heating pad. NOTE: Follow-up doses of analgesic should be given according to the frequency and dose specified in the user’s approved IACUC protocol. It is recommended a healing duration of one week between injections. If repeated injections are needed, repair of the injection site can be used after the first injection and the same surgical procedure can be followed for subsequent injections. The subsequent injection can be administered into the CCA cranial to the repaired injection site as repeated suturing to repair the same injection site will likely lead to scarring and blood clots.