A Viral Vector Injection into the Rat Spinal Cord for Retrograde Neuronal Tracing

Abstract

Source: Keefe, K. M., et al. Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord. J. Vis. Exp. (2019).

This video demonstrates the injection of a highly efficient retrograde transport (HiRet) viral vector into the rat spinal cord to trace neuronal pathways. The virus has glycoproteins for synaptic entry and a fluorescent protein coding gene. The needle is inserted into the spinal cord, and the virus suspension is administered at a controlled rate. The needle is finally withdrawn after allowing sufficient time for viral diffusion.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Pre-surgical preparations

1. Prepare pulled glass needles for viral injection a few days before surgery using 3.5-nanoliter glass capillary pipettes designed for nanoliter injectors. Pull each pipette on a two-step needle puller according to the manufacturer’s instructions to create two-needle templates.
2. Refine the tip of the needle templates by cutting off approximately 1-2 mm of excess glass with microscissors. Measure the approximate aperture size under a microscope with a microscope calibration slide to isolate needles with 30-40 µm apertures.
3. With the needle positioned at 30°, use a micropipette beveler to create a tip with a 30-40 µm aperture and a 45° beveled angle. Verify the aperture width using the Vernier scale on the calibration slide. Pass water and ethanol through the glass needle using a syringe with a flexible needle attachment to wash away debris and mark the needle at regular intervals with a black marker.
4. Place the needles in a covered Petri dish previously cleaned with 70% ethanol and sterilize for 30 min in a Biosafety hood under ultraviolet or UV light.
5. Prepare highly efficient retrograde transport (HiRet) lentivirus by removing a suitable volume from the freezer immediately prior to the procedure.
   NOTE: A suitable volume includes the amount needed for injection (1 µL per injection x number of injections) plus a small amount of extra volume to account for pipetting and loading losses. Transport and store the virus on ice when not in use.
6. Prepare the injector by plugging it into the micropump and placing it into a micromanipulator with a Vernier scale.
7. To prepare the glass needle, carefully load a colored dye such as red oil with a syringe outfitted with a flexible needle. Ensure that no bubbles remain in the needle. Use an aseptic technique when handling the needle, and refrain from touching the tip.
8. Insert the glass needle into the injector, ensuring that the needle is seated correctly into the washers, the injector cap is screwed on tight, and the steel injector needle is extended approximately ¾ the length of the glass needle. Viruses can be loaded into the needle in a later step.

2. Anesthesia and surgical site preparation

1. Weigh the animal on a digital scale. Record the pre-operative weight to determine the volume of anesthetic required and to allow for monitoring of weight post-surgery. Female Sprague-Dawley rats, approximately 200–250 g, were used in this protocol.
2. Anesthetize the rat using either isoflurane inhalation or an injected ketamine/xylazine solution (k/x). Here, ketamine is injected intraperitoneally at a 67 mg/kg and xylazine at a 6.7 mg/kg dosage.
3. Confirm an appropriate anesthetic plane by pinching the foot firmly. If reflexive withdrawal occurs, wait several additional minutes before proceeding.
   NOTE: Also observe the whiskers, eyes, and breathing rate for signs of consciousness. If the whiskers are twitching, the eye blinks when touched gently, or breathing is rapid and shallow, wait until the anesthetic plane is deeper to proceed with the protocol. Also, monitor these signs throughout the laminectomy and injection surgery. If the animal displays a shallow anesthetic plane, administer a booster shot of ketamine-only equal to ½ the original k/x dosage.
4. Shave the rat along the dorsal midline from the hips to the inferior angle of the scapulae. Pull the skin of the animal taut for an easier and more precise shave.
5. Apply ophthalmic ointment to both eyes.
6. Apply an antiseptic to the shaved area to sterilize the site. For the first scrub, soak sterile gauze with a 5% iodine solution and wipe away all hair and debris. Follow this with a unidirectional swipe with sterile gauze soaked in 70% ethanol so that no area is contacted twice. Use this same technique with alternating iodine and ethanol-soaked gauze twice more.

3. Surgical field and instrument preparation

1. Prepare a set of autoclaved surgical tools that include a scalpel, rongeurs, rat tooth forceps, spring scissors, hemostats, medium-point curved forceps, and retractors or weighted hooks by unwrapping the sterile wrap to create a sterile field.
2. Open a package of sterile surgical gloves and place the sterile glove wrap on the table. Use this as an additional sterile field for used tools to prevent contamination of the sterile wrap.
3. Drop a #10 scalpel blade onto the sterile field. Secure the blade to a handle with hemostats. Position sterile saline, 4.0 chromic catgut suture, and materials to control bleeding such as a cauterizer, sterile gauze, sterile cotton-tipped applicators (for muscle bleeds), or gelfoam or bonewax (for bone bleeds) in an accessible place.
4. Retrieve the animal and set it on a sterile cloth. Place gauze underneath the bladder to collect urine. Prop up the target area with a rolled towel under the abdomen. If available, place a surgical heating pad underneath the sterile cloth, especially for longer procedures.
   NOTE: Sterility is important during survival surgery. Keep a spray bottle of 70% ethanol on hand to maintain sterility of gloved hands, and a use bead sterilizer if instrument sterility is compromised, or between individual surgeries.

4. Exposing the vertebral column and identifying the laminectomy site

1. Identify the area where a skin incision will be made by pressing the fingers gently at the last rib to locate the Lumber or L1 vertebra. Using this as a landmark, make a 3-4 cm skin incision with a #10 surgical scalpel ending just inferior to L1 to expose the muscle. Hold the skin taut by gently spreading and pressing firmly with the scalpel blade to ensure a clean incision.
2. Cut and spread superficial fat with forceps and scissors if necessary. (Depending on the target vertebra, there may or may not be a large fat pad superficial to the muscle.)
3. Feel for the spinous processes with the flat of the scalpel blade or a finger. Often, the midline area will be outlined by a “V” of white fascia on either side. Make a small rostral cut to allow room to grab securely onto an upper process with rat-tooth forceps, then make two long, deep cuts as close to the processes as possible. At the deepest point of the cut, the dorsal surface of the vertebrae can be felt with the scalpel blade.
4. Hold the lateral muscles aside with retractors or weighted hooks to improve visibility. Clear muscle around the processes with a scalpel, spring scissors, or rongeurs to determine the shape of their heads.
   NOTE: Remember that the spinal cord does not extend the full length of the vertebral column, as spinal cord tissue stops growing earlier in development than bone. This means that the target spinal level may be underneath a differently named vertebra.
5. Locate the thoracic or T11 and the adjoining T12 and T13 processes.
   NOTE: Assistance in targeting correct vertebral levels can be found in a rat spinal cord atlas and previous studies outlining landmarks in the mouse, which has a very similar vertebral structure6,33. Leave a rostral spinous process, such as T9, undisturbed to give a midline landmark.

5. Performing a laminectomy

1. Once the target area has been correctly identified, perform laminectomies of the dorsal aspects of T11-T13. Gently spread the vertebrae to reveal intervertebral ligaments, which are good sites to insert rongeurs for the initial bite of bone. Hold the rongeurs in a half-closed position to increase fine control.
2. Remove the spinous processes and the dorsal aspect of the vertebrae by taking small bites with the rongeurs. Be careful not to damage the spinal cord or disturb the dura. Lift slightly with the rat tooth forceps to help pull the spinal cord away from the vertebrae and decrease the tendency to hit spinal cord tissue.
3. Clear the bone away from the midline so that the midline blood vessel can be observed. Leave a window that clearly shows the spinal cord tissue and is free of debris.
4. Gently touch the spinal cord with forceps. Some animals may reflexively jump even if their anesthetic plane is deep. Apply a few drops of a numbing agent, such as lidocaine, directly to the spinal cord to prevent jumping during the injection procedure.
5. Secure the animal in a spinal holder by fastening stabilizing forceps to the spinous processes rostral and caudal to the laminectomy window. Raise the abdomen of the animal using the spinal holder to negate the effect of breathing movements. This will increase needle stability and ensure the appropriate depth of injection.

6. Loading the virus and positioning the injector

1. Load the virus into the injector by pipetting approximately 5 µL onto a piece of parafilm and positioning the needle so that the tip is inside the drop.
2. Use the micropump to withdraw up to 4 µL of virus at a rate of 20–100 nL/s.
3. Set the controller to inject and release a small amount of virus from the needle to ensure the tip of the needle is not blocked. Wipe off excess virus with a laboratory wipe.
   NOTE: A Hamilton syringe with a steel needle may be used as an alternative to pulled glass pipettes.
4. Position the micromanipulator so that the Vernier scale is visible, and position the needle at the midline of the spinal cord.
   NOTE: The midline can sometimes be located by a large blood vessel running on the anterior surface of the spinal cord. However, this can vary in individual rats, and midline targeting should be confirmed by comparison with an intact spinous process.
5. Direct the needle laterally by 0.8 mm using the Vernier scale on the micromanipulator.
6. Lower the needle to the spinal cord until it is indenting, but not puncturing, the dura. Using a quick twisting motion, puncture the dura with the needle until it has sunk to a depth of 1.5 mm.

7. Injecting the virus into the spinal cord

1. Once the needle is in place, program the injector to inject at a rate of 400 nL/min. Confirm that virus is entering the spinal cord by observing the progress of the dye front. There should be no obvious leakage or bulging of spinal cord tissue. If leakage is observed, this can sometimes be alleviated by reducing the injection speed to 200 nL/min.
2. Once the injection is finished, allow the needle to rest in the spinal cord for 2–5 min (depending on volume injected) to facilitate diffusion of the virus.
3. Slowly withdraw the needle and move to the next injection site. Inject 1 µL of virus into each of 6 evenly spaced sites approximately 1 mm apart along the length of the L1-L4 spinal tissue. The same needle may be used for each injection as long as it continues to function properly.

8. Wound closure and post-operative care

1. Remove the animal from the spinal holder and take out retractors or hooks used to spread lateral muscle. Ensure that the wound is clear of all debris before closing.
2. Suture the muscle using a 4.0 chromic catgut suture. Cut suture threads close to the knot to reduce the likelihood of internal skin irritation.
3. Staple the skin closed using 9 mm wound clips. To allow for optimal healing, line up the edges of the skin before stapling.
4. Place the animal on a water convection warming pad and monitor until awake.
5. Inject 5–10 mL of sterile saline subcutaneously to replenish fluids and an antibiotic such as cefazolin to prevent infection. When the animal is ambulatory, place it back in its home cage and provide initial analgesics. Monitor the rats for any sign of pain and distress, and treat according to your institutional animal care and use committee (IACUC) approved procedure for alleviation of pain.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
#10 Scalpel Blades	Roboz	RS-9801-10	For use with the scalpel.
1 mL Syringes	Becton, Dickinson and Company	309659	For anesthetic IP injection, potential anesthetic booster shots, and antibiotic injections.
10mL Syringes	Becton, Dickinson and Company	309604	For injecting saline into the animal, post-surgery.
4.0 Chromic Catgut Suture	DemeTECH	NN374-16	To re-bind muscle during closing.
48000 Micropipette Beveler	World Precision Instruments	32416	Used to bevel the tips of the pulled glass capillary tubes to form functional glass needles.
5% Iodine Solution	Purdue Products L.P.	L01020-08	For use in sterilzation of the surgical site.
70% Ethanol	N/A	N/A	For sterilization of newly prepared glass needles, animal models during surgical preparation, and surgeon's hands during surgery, as well as all other minor maintainances of sterility.
Anesthetic (Ketamine/Xylazine Solution)	Zoetis	240048	For keeping the animal in the correct plane of consciousness during surgery.
Antibiotic (Cefazolin)	West-Ward Pharmaceuticals	NPC 0143-9924-90	To be injected subcutaneously to prevent infection post-surgery.
Bead Sterilizer	CellPoint	5-1450	To heat sterilize surgical instruments.
Bonewax	Fine Science Tools	19009-00	To seal up bone in the case of bone bleeding.
Cauterizer	Fine Science Tools	18010-00	To seal any arteries or veins severed during surgery to prevent excessive blood loss.
Digital Scale	Okaus	REV.005	For weighing the animal during surgical preparation.
Flexible Needle Attachment	World Precision Instruments	MF34G-5	For cleaning glass needles and loading red oil into glass needles.
Gelfoam	Pfizer	H68079	To seal up bone in the case of bone bleeding.
Glass Capillary Tubes	World Precision Instruments	4878	For pulled glass needles - should be designed for nanoliter injectors.
Hair Clippers	Oster	111038-060-000	For clearing the surgical site of hair.
Hemostats	Roboz	RS-7231	For general use in surgery.
Kimwipes	Kimtech	34155	For general use in surgery.
Medium Point Curved Forceps	Roboz	RS-5136	For general use in surgery.
Micromanipulator with a Vernier Scale	Kanetec	N/A	For precise targeting during surgery.
Microscissors	Roboz	RS-5621	For cutting glass whisps off of freshly pulled glass capillary tubes.
Microscope with Light and Vernier Scale Ocular	Leitz Wetzlar	N/A	Used to visualize and measure beveling of pulled glass capillary tubes into functional glass needles.
MicroSyringe Pump Controller	World Precision Instruments	62403	To control the rate of injection.
Nanoliter 2000 Pump Head Injector	World Precision Instruments	500150	To load and inject virus in a controlled fashion.
Needle Puller	Narishige	PC-100	To heat and pull apart glass capillary tubes to form glass needles.
Ophthalamic Ointment	Dechra Veterinary Products	RAC 0119	To protect the animal's eyes during surgery.
Parafilm	Bemis	PM-996	To assist with loading virus into the nanoinjector.
PrecisionGlide Needles (25G x 5/8)	Becton, Dickinson and Company	305122	For use with the 1mL and 10 mL syringes to allow injection of the animal model.
Rat Tooth Forceps	Roboz	RS-5152	For griping spinous processes.
Red Oil	N/A	N/A	To provide a front for visualization of virus entering tissue during injection.
Retractors	Roboz	RS-6510	To hold open the surgical wound.
Rimadyl Tablets	Bio Serv	MP275-050	For pain management post-surgery.
Rongeurs	Roboz	RS-8300	To remove muscle from the spinal column during surgery.
Scalpel Blade Handle	Roboz	RS-9843	To slice open skin and fat pad of animal model during surgery.
Scissors	Roboz	RS-5980	For general use in surgery.
Stainless Steal Wound Clips	CellPoint	201-1000	To bind the skin of the surgical wound during closing.
Staple Removing Forceps	Kent Scientific	INS750347	To remove the staples, should they be applied incorrectly.
Sterile Cloth	Phenix Research Products	BP-989	To provide a sterile surface for the operation.
Sterile Cotton-Tipped Applicators	Puritan	806-WC	To soak up blood in the surgical wound while maintaining sterility.
Sterile Gauze	Covidien	2146	To clean the surgical area and surgical tools while maintaining sterility.
Sterile Saline	Baxter Healthcare Corporation	281324	For use in blood clearing, and for replacing fluids post-surgery.
Surgical Gloves	N/A	N/A	For use by the surgeon to maintain sterile field during surgery.
Surgical Heating Pad	N/A	N/A	For maintaining the body temperature of the animal model during surgery.
Surgical Microscope	N/A	N/A	For enhanced visualization of the surgical wound.
Surgical Stapler	Kent Scientific	INS750546	To apply the staples.
T/Pump Heat Therapy Water Pump	Gaymar	TP500C	To pump warm water into the water convection warming pad.
Water Convection Warming Pad	Baxter Healthcare Corporation	L1K018	For use in the post-operational recovery area to maintain the body temperature of the unconscious animal.
Weighted Hooks	N/A	N/A	To hold open the surgical wound.