Direct Vagus Nerve Injection Protocol For Rats

Summary

We present a protocol for direct vagus nerve injection in rats, enabling drug delivery directly into the nerve without post-injection complications. This method applies to preclinical neurological studies involving autonomic nervous system manipulation. It can be used for direct nerve injection for other nerves in rats and other species, with necessary modifications.

Abstract

There is a relative abundance of strategies and methodologies to facilitate drug delivery to the central nervous system. However, drug delivery directly to the peripheral nervous system is less common, with fewer detailed methods publications available to aid researchers. Here, we describe a direct nerve injection method for peripheral nervous system drug delivery, using the vagus nerve as a model nerve. This method can be used in the treatment of autonomic nervous system disorders through targeting of the left vagus nerve, although this general injection method can be extrapolated to injection of other nerves with minor modification. This method explains all critical steps involved in the procedure involving microsurgery in anesthetized adult rats under a dissecting microscope. The use of a tracking dye is described to facilitate the monitoring of injection fidelity in real time. Illustrations of successful and failed injections are provided. If carried out properly, direct vagus nerve injections can be conducted in a safe manner that is well-tolerated by the rat without post-delivery complications. For example, once the surgeons were trained in this method, six out of six rats were successfully injected without any complications. This method of direct nerve injection for preclinical rat studies is capable of delivering agents (including but not limited to gene therapy) to peripheral nerves.

Introduction

The application of the correct method of drug administration is one of the critical factors in achieving successful therapeutic results. Despite the abundance of methods for therapeutic agent delivery to the central nervous system (CNS), only a few methods are reported for peripheral nervous system (PNS) delivery by direct nerve injection. Direct nerve injection, such as injection into the dorsal root ganglia (DRG) in rats, has been tried in preclinical studies for better understanding of pain mechanisms, drug toxicity, gene transfer^1,2,3, and general method development^1,4. Additional reports on direct nerve injection include spinal nerve injection⁴, sciatic nerve injection¹, and vagus nerve injection in rats⁵ and mice⁶. Recently a method for suprachondrial injection has been proposed for the better distribution of therapeutics into the optic nerve head in rabbits⁷.

DRG is considered the ideal location for direct injection of transgene-loaded vectors such as adeno-associated virus (AAV) because of the sensory function of the cell bodies in the DRG². Both surgical and non-surgical methods of DRG injections have been described^1,8. However, controversial conclusions have been found about the consistency of results with the non-surgical method of DRG injection¹. A surgical method involving a partial laminectomy has been suggested to be 100% successful for DRG injection in rats without any alteration in behavior outcomes³, as well as a method involving partial osteotomy in mice⁹. Several studies report the DRG injection methods of drug delivery, which have been used in preclinical gene therapy research in rats and mice^1,2,10. Vector-based gene therapy studies involving localized injections may include the following benefits: decreased off-target expression, reduction in systemic toxicity, and smaller viral loads and injection volumes, decreased risk of immunogenic complications^11,12.

Direct injection method into the sciatic nerve, the longest nerve of the body, has been tried by exposing the right sciatic nerve at the mid-thigh level of a rat. The method used a pulled glass pipette equipped with a microprocesser-controlled injection system to inject a total volume of 10 µL dye with a flow rate of 1.2 µL/min¹. This experiment showed a lack of dye distribution to the level of DRG, and the distribution was mostly limited around the site of injection. Similarly, other methods of direct nerve injections, such as spinal nerve injections, have been tried with dye to evaluate the appropriate amount of injection volume and dye distribution pattern in rats. 2 µL is suggested to be optimum for spinal nerve injection, whereas 3 µL of dye by DRG injection showed the distribution in both dorsal and ventral root ganglia in rats¹. The volume for DRG injection in mice has been reported to be optimum from 1.0 µL to 1.5 µL based on strain and body size^2,9.

Direct vagus nerve injection method was used in rats⁵ and mice⁶ to evaluate the role of neural injury or cellular integrity in transferring human α-synuclein. These two studies, conducted by the same group of researchers, describe a brief method of directly injecting AAV vectors into the left vagus nerve at the cervical region. In rats, the method involved a glass capillary with a 60 µm tip diameter to inject 2 µL vector at a flow rate of 0.5 µL/min with a 5 µL Hamilton syringe. In mice, a total volume of 750 nL vector solution was injected at a flow rate of 160 nL/min using a 36-G blunt steel needle fitted onto a 10 µL NanoFil syringe⁶. These experiments showed that the transgene was delivered and expressed in axons in the pons and midbrain of the rats and mice. Similarly, the dorsal motor nucleus of the left vagus nerve showed positive immunoreaction with transgene. These pieces of evidence illustrate that the direct vagus nerve injection method might be a reliable method in gene therapy where the cellular transduction is extended into several locations of the brain, which project axons through the vagus nerve. However, these methods do not mention the use of any dye to track the injection fidelity.

Here, a method is described for direct injection into the left vagus nerve using nontoxic tracking dyes largely applicable to researchers in preclinical studies. Potential pitfalls that may cause difficulties in drug delivery and the ways to overcome them are discussed. These situations are illustrated with pictures to show what makes the delivery unsuccessful and the way to make it successful.

Protocol

The following protocol is conducted in accordance with institutional ethics guidelines and approval from the Institutional Animal Care and Use Committee (IACUC).

1. Housing preparation

NOTE: This protocol is for adult rats aged at least 2 months. Smaller animals (including mice and younger rats) are possible but are not recommended and will be considerably more difficult.

1. Provide moist chow or any other soft food (veterinary recovery gel) for 48 h prior to vagus nerve surgery. As animals may face difficulty eating after the procedure, introduce these items to the animals before surgery to help them become familiar with the taste of these items and induce eating motivation.
2. House rats individually in a cage at least 1 week prior to surgery, as well as 1 week post-surgery. That will enable them to acclimate to the stress of being alone and prevent scratching of wounds after surgery by a littermate.

2. Preparation of the surgical items and space

1. Make a list of all necessary items ahead of time and double-check that all items are available for the surgery.
2. Sterilize all items with appropriate methods of sterilization before the surgery.
3. Set up the sterile surgical space under the dissecting microscope. Place a heating pad under the microscope to keep the rat warm during surgery.
4. Place all surgical materials on a surgical table on a sterile drape. Keep enough space for surgery under the dissecting microscope. Make sure the microscope's focus is optimum and encompasses the surgical area to visualize the nerve.
5. Place the syringe pump close to the surgical stage so that the length of the tubing is enough to reach the site of injection.
6. Place the surgical tools towards the side of the surgeon's dominant hand.

3. Preparing a mixture of the candidate drug and the tracking dye

1. Ensure the candidate drug is diluted to achieve the required concentration in the recommended diluent such that 5 µL of the drug mixed with dye gives the required amount of the candidate drug to be delivered. A maximum volume of injection by the direct vagus nerve method is recommended to be 5 µL (including tracking dye, see step 3.2).
2. Mix a compatible dye with the candidate drug to enable the tracking of injection fidelity in real-time. For example, a final concentration of either 1% Luxol Fast Blue or 0.002% fluorescein are compatible dyes to use.
   CAUTION: Use the dye at its optimum concentration in the dye and candidate drug mixture. When the concentration of dye in the mixture is too low, it may not be visible to track the injection. When the concentration of dye is too high, it may result in a mixture that is too thick and has high viscosity, which will block the injection needle.

4. Loading the candidate drug and dye mixture into the tubing

1. Connect a 100 µL glass syringe with the plunger removed to a 27 G needle.
2. Connect one end of approximately 45 cm of sterilized polyethylene tubing to the 27 G needle.
3. Take a 3 mL plastic syringe and fit it with a 27 G needle. Load the syringe with about 0.5 mL of 0.9% normal saline.
4. Fill the polythene tubing with the 0.9% normal saline from the open end to fill the full length of the tubing and the whole barrel of the glass syringe until it overflows out.
   NOTE: It is critical to avoid air bubbles getting trapped inside the tubing.
5. Remove and discard the 3 mL syringe and needle.
6. Insert the syringe plunger into the barrel of the glass syringe and push it slightly forward up to the middle of the barrel.
7. Place the glass syringe in the syringe pump.
8. Pull the plunger slightly back using the syringe pump to create an air space of about 1.5 cm at the open end in the length of the tubing.
9. Pipette at least 5 µL of dye and candidate drug mixture with a 10 µL micropipette and drop it onto a sterilized piece of parafilm.
10. Withdraw the candidate drug mixture into the tubing by pulling the plunger back via the syringe pump. Maintain the air bubble between the 0.9% saline and the injection solution, but otherwise, avoid trapping any air bubbles within the injection solution. Mark the level of the injection mixture in the tubing with a marker.
    NOTE: The air gap between the candidate drug-dye mixture and the sterile saline within the tubing is critical to prevent the mixing of the two solutions.
11. Fit the 35 G needle with the tubing. Fix the needle onto the clean surface of the surgical stage with tape so that the needle is not loose, does not move and touch anything else, and remains sterilized.

5. Priming of the injection needle

1. Set the syringe pump program to dispense 0.5 µL/min.
2. Start the syringe pump for about 10-15 s, until the injection solution starts to come out of the tip of the needle. A tiny amount of the mixture at the tip of the needle without a leak at the joint of the tubing and injection needle (or anywhere else along the length of the syringe, tubing, or needle) indicates that the setup is correct.
   NOTE: If a tiny droplet at the tip of the needle is not seen, that indicates a blockage of the needle. Needle blockage may occur due to the high viscosity of the candidate drug and dye mixture, excess air trapped within the system, and the glass syringe is not properly seated within the syringe pump. See Figure 1 for an example of leakage at the junction between the needle and tubing. Priming is critical because it helps to identify if the needle is unobstructed and the injection inside the nerve would pass smoothly.

6. Preparation of rat for surgery

1. Obtain the body weight of the rat to calculate the necessary dose of analgesics. For example, carprofen is used at the dose rate of 5 mg/kg subcutaneously in rats.
   NOTE: Carprofen is recommended to store at 4 °C before injection.
2. Anesthetize the rat with an isoflurane vaporizer using a flow rate of 3%-4% for induction of anesthesia for about 2-3 min. Reduce the isoflurane flow rate to about 1.75%-2% for maintenance of anesthesia.
3. Transfer the rat to a separate table with a setup for hair removal and site preparation. Ensure this table is close to the surgical space.
4. Apply artificial tearing ointment in both eyes of the rat to prevent excessive dryness.
5. Administer analgesics per the approved institutional animal protocol to control pain when the rat comes back into consciousness.
6. Shave the surgical area with a hair clipper at the ventral side of the neck at the cervical region. Shave up to about 2 cm perpendicular distance from the midline on both sides from chin to sternum.
7. Sterilize the area with 70% ethyl alcohol and 10% betadine, wiping the area three times alternatively with an alcohol swab and povidone-iodine. Wipe from the center of the area outward in a circular pattern.
8. Transfer the rat to the surgical stage under the microscope in a supine position. Adjust the focus of the microscope to visualize the area of surgery at the cervical region.

7. Performing rat surgery to inject the candidate drug directly into the left vagus nerve (Figure 2, Figure 3, and Figure 4)

NOTE: This portion of the method will require a second person to assist the surgeon.

1. Place the rat on a heating pad in a supine position under the dissecting microscope associated with a mounted light.
2. Hook the front upper jaw teeth of the rat with a piece of non-absorbable suture and fix the open end of the suture inside the nose cone so that the nostril of the rat is always inside the nose cone throughout anesthesia.
3. Position the rat so its head lies on the left side of the right-handed surgeon (or the right side of a left-handed surgeon). In this position, the front side of the surgeon is perpendicular to the body of the rat. Place a cushion of sterilized gauge under the neck and adjust the angle of the cervical region of the rat so that the cervical region becomes straight. This will make it easier to locate the left vagus nerve and perform the injection.
4. Drape the whole body of the rat with a sterilized drape, keeping the surgical space open.
5. Fix four retractor pins individually with 4 magnetic fixators with elastomers and place the fixators at the four corners of the surgical stage.
6. Inject local anesthetics epidermally at the midline of the cervical region where the incision is made. For example, a mixture of lidocaine and bupivacaine is usually used.
7. Make about a 2 cm long longitudinal skin incision at the midline between the chin and sternum with a scalpel blade in the ventral side of the neck at the cervical region (Figure 3A).
8. Blunt separate apart the cut edges of the skin with sterilized cotton tips.
9. Retract the skin edges in opposite directions from the site of the incision with the help of retractor tips.
10. Separate the facia with cotton tips to go deeper. Push the salivary glands towards the lateral side.
11. Separate the sternomastoid muscles with cotton tips and retract them aside with the pins. As the sternomastoid muscle separation advances wide, the trachea appears in the middle, covered with sternohyoid muscle (Figure 3B).
    NOTE: Be careful not to exert pressure on the trachea.
12. Advance the tissue separation on the rat's left side of the trachea until the carotid sheath appears, which contains the common carotid artery running together with the left vagus nerve.
13. Carefully make a small point hole into the carotid sheath with fine forceps, paying attention so as not to injure the carotid artery under the microscope. Separate the left vagus nerve from the common carotid artery with fine forceps.
    NOTE: The surgical area is anatomically critical. The wall of the carotid artery is thick, slippery, and not injured easily, but the sharp tip of fine forceps may injure it, and bleeding may happen. At the same time, pay attention so as not to injure the nerve.
14. Once the nerve is separated from the artery by the forceps, use a 25 G curved needle fitted to a 1 mL syringe from that hole to "hook" and hold the nerve by the non-dominant hand (Figure 4A).
    NOTE: The tip of the curved needle is made blunt so as not to injure the nerve and artery. The angle of curvature of the needle tip is custom-made to about 90° using a hemostat (Figure 2).
15. With the dominant hand, hold the needle loaded with the candidate drug-dye mixture and gently prick the nerve in the same direction that the nerve runs so that the bevel of the needle is facing up.
    1. For an easy prick, keep the nerve straight by gently pulling using the 25 G hook. Insert the needle into the nerve and advance forward, more than 0.5 cm, while keeping the needle parallel to the vagus nerve. Then, pull slightly back so that about 0.4-0.5 cm needle remains inside the nerve.
       NOTE: The angle of the needle should be shallow enough to penetrate inside the nerve without going through to the other side.
16. There might be occasions when the needle is unobstructed outside the nerve at the time of priming, but the candidate drug-dye mixture does not pass smoothly inside the nerve. To avoid this situation, limit the number of nerve pricks by a single needle not more than 3 times. If the needle gets blunted, the chances of blocking or otherwise poor injection will increase.
17. Make sure the needle is completely inside the nerve. Retain the position immobile and turn the syringe pump on.
    NOTE: Check carefully at the site of injection to see that the candidate drug-dye mixture is not back-flowing out of the nerve.
18. While the injection is proceeding, constantly examine that the infusion is remaining inside the nerve and flowing smoothly. Use the mark made on the infusion tubing to monitor the movement of the drug-dye mixture from the starting point.
    1. If the candidate drug-dye mixture is not flowing smoothly, pull the needle out and repeat the priming step (step 5.2). Then re-insert the needle into the nerve and resume the infusion.
19. The nerve starts to develop the color of the dye (Figure 4B). Continue the infusion at 0.5 µL/min for 10 min to infuse all 5 µL of the candidate drug-dye mixture.
    NOTE: It is recommended to irrigate the exposed tissues with a small amount of warm 0.9% sterile saline as needed to prevent excessive dryness and damage.
20. After the 5 µL has been infused and the syringe pump stopped, retain the needle in place for an additional 1 min to allow all of the candidate drug-dye mixture to spread from the injection site. Remove the needle.
21. Remove the 25 G curved needle. Use sterile cotton tips to gently move tissues back into their original location. Remove the retractor tips.
22. Close the wound with a non-absorbable suture (absorbable suture can also be used). Apply a tiny amount of tissue glue between the stitches if necessary to close the wound perfectly.

8. Post-operative care of a rat

1. Infuse about 3 mL of 0.9% warm saline subcutaneously after the wound is closed. This helps the rats to rehydrate immediately.
2. Place the rat in a heat source until it recovers fully from anesthesia to normal condition. Note anything observed that is abnormal during the process of recovery.
   NOTE: the recovery duration may be longer if the nerve injection procedure takes a long time.
3. Transfer the rat into its home cage with moist chow on the floor of the cage. Make sure water is available all the time. Observe the rat post-operatively for about 2 h to ensure the rat is not feeling cold.
   NOTE: Place the rat in a heat source for additional hours if it feels cold.
4. Monitor the rat at 12 h, 24 h, 36 h, 48 h, and 72 h post-surgery with written notes of observations. Repeat analgesics as per the approved institutional animal protocol to control pain. For example, carprofen is dosed at the rate of 5 mg/kg subcutaneously two times at the interval of 24 h. Consult a veterinarian if pain remains in the rat for more than 48 h.
5. Repeat the subcutaneous injection of about 3 mL 0.9% warm saline at 24 h based on the conditions of the rat.
6. Continue to evaluate the wound. Ensure the wound is dry and is in the process of healing.
   NOTE: Redness, swelling, and painful condition of the wound with inactivity and dullness of the rat can be an indication of wound inflammation and may warrant additional veterinarian consultation. A rat with pain will show a hunched back, ruffled coat, red eyes, and inactive condition. It is usually found in a corner of the cage.
7. Remove the non-absorbable sutures within 2 weeks once the wound is healed.

Results

Six adult rats (3 males and 3 females) were used in the present study. Out of six, one rat was injected multiple times to demonstrate a condition of failure injection that shows a diffusion of dye around the injection site Figure 5A. All the other rats showed smooth injection and nerve staining, as shown in Figure 4B and Figure 5B.

Discussion

The method for direct injection into the left vagus nerve can be done safely and without post-surgery complications in rats. Drug delivery to the vagus nerve can be used to target the autonomic nervous system (ANS). This involves certain critical steps that require practice and a moderate to high degree of surgical skill.

This surgical procedure requires balanced general anesthesia in rats. The surgeon aims to finish the surgery within a shorter duration to limit the exposure of anesthetics f...

Materials

Name	Company	Catalog Number	Comments
1 mL BD Tuberculin Syringe with Detachable 25 G x 5/8". Needle	Becton, Dickinson and Company	SKU:309626	Used to connect with curved needle to pull the vagus nerve and hold it at the time of injection.
0.5% Bupivacaine Hydrochloride Injection	Hospira	NDC 0409-1162-19	Local anesthetics used to anesthetize local tissue.
100 mL 0.9% Sodium Chloride Irrigation USP	Stericare Solutions	Item #6240	Normal saline, used to rehydrate rat and tissue.
20 Blunt, Retractor Tips, 7.5 mm	Kent Scientific Corporation	Surgi 5018	Used to pull apart and hold tissues at the time of surgery.
3 mL BD-Luer-Lok Syringe, Sterile, Single Use	Becton, Dickinson and Company	SKU # 309657	Used to inject saline in rat and fill the saline into the Polythene tubing.
AK-Fluor10%	Akorn	NDC 17478-253-10	Fluorescein dye visible within the nerve. Used to track injection fidelity.
Animal Weighing Scale	Kent Scientific Corporation	SCL 4000	Used to measure body weight of rat.
Ansell ENCORE Perry Style 42 PF Surgical Gloves	Ansell	ASTM D3577	Sterilie glove, it is used at the time of surgery by a surgeon.
Artificial Tears Ointment 3.5g	Pivetal	NDC 46066-753-55	Used in eyses to prevent excessive dryness of eyes.
Baby-Myxter Hemostat	Fine Science Tools	13013-14	Used to stop bleeding in case of emergency. Also used to bend the 25 G x 5/8" in needle.
BD Intramedic PE Tubing	Becton, Dickinson and Company	14-170-12A	Used in the injection set up system to connect with Hamilton needle and NanoFil Needles. It also holds the injection mixture.
BD Precison Glide Needle, 25 G x 5/8"	Becton, Dickinson and Company	REF#305122	Used to inject saline in rat, and to make a curved needle.
BD Precison Glide Needle, 27 G x ½"	Becton, Dickinson and Company	REF#301629	Used to fill sterile saline into the BD Intradermic tubing.
Benchmark Accuris ”NextPette” Variable Volume Pipette Micro Starter Setincludes 4 pipettes: 10/20/200/1000 μL, plus stand	MilliporeSigma	BMSP7700S1	Used to pippette sterile solution.
Betadine, Povidine Iodine 10%	Honestmed	67618015017	Used to disinfect the surgical area.
Carprofen Injectable solution 50 mg/mL	Supplied by Covtrus (6451506845)	SKU 591149	In our case, we used diluted carprofen at the dose rate of 5 mg/kg provided by the Animal Resource Center of University of Texas Southwestern Medical Center.
Curved needle (custom made)	Becton, Dickinson and Company	REF#305122	BD PrecisionGlide 25 G x 5/8" in needle is curved to 90 degrees with the help of a hemostat. The tip of the needle is made blunt. It needs to be sterilized before use. It is used to hook the vagus nerve and hold it at the time of separation and injection.
Dissecting microscope	Motic	SMZ-171-BLED (Binocular with Lights)	Used to magnify the crifical anatomical area at the time of vagus nerve separation, injeciton, and to check injection leakage.
Drape sheet	Dynarex	Reorder#8122	Used as drape after sterilization.
Dukal Cotton Tip Applicators, Non-Sterile	Dukal	Item 9003	Used to blunt separation of tissue, needs to sterilize before use.
Dumont #7 - Fine Forceps	Fine Science Tools	11274-20	Used to separate the left vagus nerve from common carotid artery. It is curved so easy to use.
Ethicon PDS II Undyed Monofilament Suture - SUTURE, 4/0 18 PDS II CLR MONO PS	Ethicon	VA - Z682G	Used in suturing the wound.
Ethilon Nylon Suture Black Monofilament	Ethicon	1856G	Used in suturing the wound if non-absorbale suture is used. Also used to hook the rat tooth to fix nose inside the nose cone.
Fine Forceps - Mirror Finish	Fine Science Tools	11412-11	Used at the time of vagus nerve separation from the common carotid artery. This is straight.
Fine Scissors - Sharp	Fine Science Tools	14060-09	Ued to cut tissue.
Hamilton cleaning solution	Hamilton	HT18311	Used to clean the Hamilton after use.
Hamilton Needle, 27G, Small Hub RN Needle, 2”, PT3, 6/PK	Hamilton	7762-01	Used to connect BD Intramedic™ PE Tubing.
Hamilton Syringe , 710RN	Hamilton	7638-01	Used to hold drug at the time of vagus nerve injection.
Insulin Syringe	EXEL INT, Comfort point	REF 26027	Used to inject carprofen and local anesthetics.
Lidocaine 2% Injection	Covetrus	Reorder#002468	Used to mix with Bupivacaine and inject at the site of incision.
Luxol Fast Blue MBSN	Acros Organics	212170250	Dye visible within the nerve, used to mix with drug so that injection mixture is visible.
Micro Bead Sterilizer with Glass Beads	Fine Science Tools	Item No. 18090-46	Used to sterilize surgical tools in between the rat surgery.
NanoFil Needles-NF35BV-2	World Precision Instrument	NC9708956	Used to inject drug - dye mixture inside the vagus nerve.
Olsen-Hegar Needle Holders with Suture Cutters	Fine Science Tools	12002-12	Used in wound suturing.
Parafilm M Laboratory Wrapping Film, 4 Inches x 125 Feet, 1 Roll per Box, 12 Count	Honestmed	PM#996	Used to hold the aliquoted 5 uL of drug-dye mixture so that loading of drug-dye mixture into the BDTM intradermic tubing is accurate.
PDI Alcohol Prep Pads	Honestmed	NDC 10819-3914-2	Used to disinfect the surgical area.
Premium Care Sterile Type VII Gauze Sponges, 8-Ply, 2" x 2"	Dukal	Item C5119	Used as cushon under the neck of rat at the time of surgery.
Press’n Seal Cling Film	Glad		Used to cover a rat at the time of surgery like a drape.
Rat Retractor Set	Kent Scientific Corporation	Surgi 5002	Used to keep the incision open so that it is easy to separate the vagus nerve from the carotid artery.
RightTemp Jr.	Kent Scientific Corporation		20.3 cm W x 25.4 cm L (8 in W x 10 in L), used to keep rat warm.
S&T Forceps - SuperGrip Tips	Fine Science Tools	00632-11	Used at the time of suturing to hold tissue without damage.
S&T Suture Tying Forceps	Fine Science Tools	00272-13	Used to tight the suture.
Scalpel blade #15	Fine Science Tools	10015-00	Used to make an incision in the skin at the ventral side of neck.
Scalpel Handle-#7	Fine Science Tools	10007-12	Used to hold the scalpel blade.
Syringe Pump	KD Scientific	78-81-8052GL	Serial #D107034, Model#LEGATO-180, is a programmable pump that can pump small volume of mixture under a program.
TipOne Filter Tip Refill Starter Systems	USA Scientific	Item #1120-3510	Used to pipette the drug and dye mixture.
Vaporizer for Isoflurane, Funnel Filled	Kent Scientific Corporation	Vetflow 1231	Used to anesthetize rats.
Vetbond Tissue Adhesives	3M Science Applied to Life	ID B00016067	Used to seal tissue at the site of cut wound if suturing is not perfect.
Wahl BravMini+ Professional Cordless Clipper Kit	Kent Scientific Corporation	CL7300-Kit	Used to cut hair of rat.