Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Summary

A surgical procedure is described to perform injections into the lumbar cistern of the juvenile rat. This approach has been used for the intrathecal delivery of gene therapy vectors, but it is anticipated that this approach can be used for a variety of therapeutics, including cells and drugs.

Abstract

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a toxic gene, or provide a gene whose product can modulate the biology of the disease. The delivery method for the therapeutic vector can take many forms, ranging from intravenous infusion for systemic delivery to direct injection into the target tissue. For neurodegenerative disorders, it is often desirable to skew transduction towards the brain and/or spinal cord. The least invasive approach to target the entire central nervous system involves injection into the cerebrospinal fluid (CSF), allowing the therapeutic to reach a large fraction of the central nervous system. The safest approach to deliver a vector into the CSF is the lumbar intrathecal injection, where a needle is introduced into the lumbar cistern of the spinal cord. This technique, also known as a lumbar puncture, has been widely used in neonatal and adult rodents and in large animal models. While the technique is similar across species and developmental stages, subtle differences in size, structure, and elasticity of tissues surrounding the intrathecal space require accommodations in the approach. This article describes a method for performing lumbar puncture in juvenile rats to deliver an adeno-associated serotype 9 vector. Here, 25-35 µL of vector were injected into the lumbar cistern, and a green fluorescent protein (GFP) reporter was used to evaluate the transduction profile resulting from each injection. The benefits and challenges of this approach are discussed.

Introduction

The promise of viral-mediated gene therapies has finally been realized in recent years with the FDA approval of treatments for spinal muscular atrophy, retinal dystrophy, factor IX hemophilia, cancer, and more^1,2,3,4. Countless other therapeutics are currently in development. Gene therapy aims to deliver a therapeutic gene to a patient's cells. The products of this new gene can replace the missing activity from a deficient endogenous gene, inhibit a toxic gene, kill cancerous cells, or provide some other beneficial function.

For diseases affecting the central nervous system (CNS), delivering the gene therapy vector directly to the target tissue is often desirable. Non-systemic approaches provide two benefits: they minimize off-target side effects that may be caused by peripheral transduction, and they greatly reduce the amount of vector needed to achieve adequate levels of transduction in the target tissue⁵.

There are a variety of approaches to delivering gene therapy vectors to the CNS. Intraparenchymal injection, the injection of a vector directly into the spinal cord or brain tissue, can be used for delivery to a defined region. However, for many diseases, broad transduction of the CNS is desired. This can be accomplished by delivering a vector to the cerebrospinal fluid (CSF)⁵, the fluid that flows in and around the brain and spinal cord. There are three primary ways to deliver vectors to the CSF. The most invasive approach is intracerebroventricular delivery, which involves drilling a burr hole through the skull and advancing a needle through the brain into the lateral ventricles. This yields transduction throughout the brain. However, the procedure may cause intracranial hemorrhage, and the approach generally produces only limited transduction of the spinal cord⁶. Injection into the cisterna magna at the base of the skull is less invasive, but carries the risk of damage to the brainstem. While often used in animal research⁵, injection into the cisterna magna is no longer used routinely in the clinic⁷. Lumbar puncture is the least invasive approach to access the CSF. This involves placing a needle between two lumbar vertebrae and into the lumbar cistern.

Lumbar puncture for vector delivery is routinely performed in adult rats and mice and in neonatal mice^8,9. The authors of this study recently performed lumbar punctures in juvenile rats (28-30 days of age) to deliver adeno-associated virus serotype 9 (AAV9) vectors. In adult rats, a neonatal lumbar puncture needle was placed vertically between the L3 and L4 vertebrae⁹. Proper placement results in a tail flick and CSF flowing up into the needle reservoir. In juvenile rats, though, neither of these read-outs could be achieved. The authors then attempted to adapt an adult mouse procedure using a 27 G insulin syringe inserted at an angle between L5 and L6¹⁰. In adult mice, which are typically smaller than P28 rats, this does not produce a tail flick, but incorrect needle placement is evident by the backflow of the injectate. In juvenile rats, however, this approach uniformly led to the injectate being delivered epidurally, likely resulting from different elasticity between adult mice and juvenile rats of the tissue layers surrounding the spinal cord. Catheter approaches were evaluated next. Specifically, a catheter was introduced through an incision in the dura of the lumbar cistern and up to the mid-thoracic spinal cord; however, this approach led to substantial reflux of the injectate back out of the incision site during delivery. Attempts to place the catheter into the intrathecal space percutaneously using a guide needle were also unsuccessful. Due to the narrowness of the interlaminar width, the catheter would likely hit the rostral lamina and fail to advance.

Here, a method is described to achieve successful and reproducible solution delivery via a lumbar puncture in the juvenile rat. This approach can be used for viral vectors, and likely also for cells, pharmaceuticals, and other therapeutics.

Protocol

This study was approved by the Emory University Institutional Animal Care and Use Committee (IACUC). Sprague-Dawley rats (28-30 days of age, mass in the range of about 90-135 g, males and females) were used in the present study.

1. Preparation of the vector

1. Thaw the AAV9 vector (see Table of Materials) on ice at the beginning of the procedure.
2. Centrifuge the microcentrifuge tube containing the vector briefly in a table-top centrifuge to ensure that all of the liquid is in the bottom of the tube.
3. Gently flick the microcentrifuge tube to ensure that the solution is well-mixed.

2. Preparation of the recovery cage

1. Place a clean cage onto an electric blanket (see Table of Materials) so that only half of the cage is in contact with the blanket.
2. Set the temperature of the blanket to ~37 °C.

3. Preparation of the surgical platform

1. Warm an isothermal pad (see Table of Materials) to 39 °C in a microwave or a water bath so that the contents become liquid.
2. Place the isothermal pad on the surgical platform and cover it with a clean absorbent bench pad.

4. Animal preparation

1. Anesthetize the rats with isoflurane in a clear box (following institutionally approved protocols). Begin the anesthesia induction using 5% isoflurane, and step down 1% per minute until reaching 2%. Hold the animal at 2% for an additional 3 min.
2. Move the box holding the animal to a fume hood and open the box.
   NOTE: This limits the surgeon's exposure to the anesthetic.
3. Remove the hair from the back of the animal using electric hair clippers.
   NOTE: Alternatively, a depilatory cream or manual razor and shaving cream can be used.
4. Place the animal on the surgical platform with its snout in the anesthesia nose cone.
   NOTE: The animal may begin to regain consciousness while the fur is removed from the surgical site. If this happens, anesthetize it again as described above.
5. Apply lubricating eye ointment to each eye to prevent drying of the corneas during the procedure.
6. Disinfect the surgical area using three alternating applications of povidone-iodine and isopropanol wipes.
7. Inject the analgesic(s) subcutaneously.
   NOTE: Buprenorphine is generally used at a dose of 0.01-0.05 mg/kg, given every 12 h. Alternatively, a slow-release form of this drug can be given once at 1 mg/kg to provide adequate pain control for 72 h. Consult with the institution's IACUC for their guidelines regarding pain management.
8. Inject 100 µL of 1% lidocaine subcutaneously above the L2 to L6 spinous processes to provide local anesthesia.
9. Place a roll of paper towel or a tube of 1.5 cm in diameter under the animal, just rostral to the hips. This helps flex the spine, making it easier to insert the needle between the two laminae.
10. Place a fenestrated drape (see Table of Materials) on the animal, centering the fenestration over the lumbar spine.

5. Exposing the lumbar spine

1. Confirm the depth of anesthesia by pinching each of the animal's paws and looking for the absence of a withdrawal response.
2. Using a #11 scalpel blade, create an incision of approximately 3 cm long in the skin down the midline from L2 to L6.
3. Loosen the skin from the muscle by inserting a sterile curved pair of surgical scissors between the muscle and the skin and then opening the tips.
4. Remove the fascia covering the L2-L5 spinous processes.

6. Loading of the syringe

1. Pipette 25-35 µL of the vector (to achieve the desired dose) into the cap of a sterile microcentrifuge tube.
2. Draw the entire volume into the insulin syringe.
   NOTE: Take care not to draw up air during this process.

7. Performing the injection

1. Identify the L5 and L4 spinous processes.
   NOTE: L6 sits directly between the two iliac crests, and its spinous process should be easy to identify by probing with a blunt instrument. The instrument can then be gently run up the back to find the boundaries of the L5 and L4 processes.
2. Place one hand so that the thumb rests gently on the animal's tail and one leg. Use a finger to steady the syringe.
3. Position the needle of the syringe so that it is to the left of the L5 spinous process and lined up with its caudal end. Position the syringe so that it is about 30° off midline and 30° up from the plane of the table.
   NOTE: It may be helpful to use a surgical microscope to better identify landmarks and position the tip of the needle.
4. Advance the syringe needle forward about 8 mm, over the top of the L5 lamina and then under the L4 lamina into the lumbar cistern until the bone is hit. Correct placement will result in a twitch of the leg and/or tail that can be seen or felt by the thumb resting on the leg/tail. If there is no twitch, remove the needle and attempt the procedure from the left side. If there is still no twitch, repeat the procedure between L4/L3 and L3/L2 as necessary.
5. Depress the plunger slowly for about 5 s.
   NOTE: There may be a twitch in the leg or tail during the injection.
6. Hold the syringe in place for about 30 s after fully depressing the plunger to allow the pressure to equilibrate and minimize reflux of the injectate when the needle is withdrawn.
7. Slowly remove the needle.

8. Closing of the incision

1. Approximate the edges of the incision.
2. Beginning at one end of the wound, use a 4-0 suture (see Table of Materials) or surgical staples to close the incision.

9. Recovery and monitoring

1. Place the animal in the prewarmed cage.
2. Check the animal at least every 15 min until it is fully ambulatory.
   NOTE: This should take between 15 min and 45 min.
3. For the next three days, perform wellness checks at least daily. Provide analgesics for the first 2 days following surgery or as required by the IACUC.
4. One week after surgery, remove the sutures or staples.

10. Follow-up procedure

NOTE: To determine the accuracy of the injection technique, inject trypan blue dye as described above and then immediately euthanize the animal (following institutionally approved protocols) and perform a laminectomy to visualize the result.

1. While the animal remains under anesthesia, euthanize it by administering a lethal dose of pentobarbital via intraperitoneal injection at a dose of 150 mg/kg.
2. Once respiration and cardiac activity cease, open the chest cavity to ensure death. Extend the surgical incision up the back to the neck.
3. Make an incision 4 cm long into the muscle parallel to the spine on both sides of the spinous processes, keeping as close as possible to the processes.
4. Using fine forceps or scissors, remove the muscle from in between the spinous processes.
5. Remove the spinous processes from L6 up to the lower-thoracic spine using a rongeur (see Table of Materials). Avoid twisting motions, as this may damage the rongeurs.
6. Insert the lower tip of the rongeur under the L5 lamina and remove the bone overlying the spinal cord by taking several "bites" out of it.
   NOTE: Pulling back on the L6 spinous process can make it easier to insert the tip of the rongeur. Care must be taken to prevent damage to the spinal cord.
7. Continue expanding the laminectomy at least four laminae rostrally. Inspect the interior surface of the laminae for signs of dye, which can indicate a failed injection.

Results

To determine the accuracy of the injection technique, a dye, trypan blue, was used as a surrogate for the therapeutic. This dye readily binds to proteins, so it generally stays within the structure into which it was injected. This means the dye may not accurately predict the post-injection distribution of the therapeutic; it is simply used to reveal the accuracy of the injection. When successfully introduced into the lumbar cistern, trypan blue binds to the dura mater, staining the perimeter of the spinal cord blue. Howe...

Discussion

A wide variety of diseases affect the CNS. Providing a functional copy of the relevant gene via a viral vector is an attractive treatment strategy for those that are recessive and monogenic in nature, such as spinal muscular atrophy. However, the blood-brain barrier (BBB) excludes most gene therapy vectors given intravenously¹¹. Those that can cross the BBB, such as AAV9, must be given in high doses to overcome the vector loss due to peripheral transduction¹². The age is al...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
200 µL filtered pipette tips	MidSci	PR-200RK-FL	Pipetting virus
AAV9-GFP	Vector Builder	P200624-1005ynr	AAV9 vector expressing GFP
Absorbable Suture with Needle Coated Vicryl Polyglactin 910 FS-2 3/8 Circle Reverse Cutting Needle Size 4 - 0 Braided	McKesson	J422H	Suture
Bench pad	VWR	56616-031	Surgery
Braintree Scientific Isothermal Pads, 8'' x 8''	Fisher Scientific	50-195-4664	Maintains body temperature
Buprenorphine	McKesson	1013922	Analgesic
Buprenorphine-ER (1 mg/mL)	Zoopharma		Extended-release analgesic
Cotton swabs	Fisher Scientific	19-365-409	Blood removal
Drape, Mouse, Clear Plastic, 12" x 12", with Adhesive Fenestration	Steris	1212CPSTF	Surgical drape
Dumont #5 Forceps	Fine Science Tools	11251-20	Forceps
Electric Blanket	CVS Health		CVS Health Series 500 Extra Long Heating Pad
Eppendorf Research plus, 1-channel pipette, variable, 20–200 µL	Eppendorf	3123000055	Pipetting virus
Fine Scissors	Fine Science Tools	14059-11	Curved surgical scissors
Friedman-Pearson Rongeurs	Fine Science Tools	16121-14	Laminectomy
Halsey Needle Holders	Fine Science Tools	12001-13	Needle driver
Insulin Syringes with Ultra-Fine Needle 12.7 mm x 30 G 3/10 mL/cc	BD	328431	Syringe
Isoflurane	McKesson	803250	Anesthetic
Isopropanol wipes	Fisher Scientific	22-031-350	Skin disinfection
Lidocaine, 1%	McKesson	239935	Local anesthesia
Microcentrifuge Tubes: 1.5mL	Fisher Scientific	05-408-137	Loading the syringe
Povidone-iodine	Fisher Scientific	50-118-0481	Skin disinfection
Scalpel Handle - #4	Fine Science Tools	10004-13	Scalpel blade holder
Sure-Seal Induction Chamber	Braintree Scientific	EZ-17	Anesthesia box
Surgical Blade Miltex Carbon Steel No. 11 Sterile Disposable Individually Wrapped	McKesson	4-111	#11 Scalpel blade
SYSTANE NIGHTTIME Eye Ointment	Alcon		Eye ointment
Trypan Blue	VWR	97063-702	Injection