The authors would like to thank Steven Gray, Matthew Rioux, Nanda Regmi, and Lacey Stearman of UT Southwestern for a productive discussion of the challenge posed by juvenile rats for intrathecal injection. This work was partly supported by funding from Jaguar Gene Therapy (to JLFK).

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
<ol>
	<li>Thaw the AAV9 vector (see Table of Materials) on ice at the beginning of the procedure.</li>
	<li>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.</li>
	<li>Gently flick the microcentrifuge tube to ensure that the solution is well-mixed.</li>
</ol>
2. Preparation of the recovery cage
<ol>
	<li>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.</li>
	<li>Set the temperature of the blanket to ~37 &#176;C.</li>
</ol>
3. Preparation of the surgical platform
<ol>
	<li>Warm an isothermal pad (see Table of Materials) to 39 &#176;C in a microwave or a water bath so that the contents become liquid.</li>
	<li>Place the isothermal pad on the surgical platform and cover it with a clean absorbent bench pad.</li>
</ol>
4. Animal preparation
<ol>
	<li>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.</li>
	<li>Move the box holding the animal to a fume hood and open the box. 
	NOTE: This limits the surgeon&#39;s exposure to the anesthetic.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Apply lubricating eye ointment to each eye to prevent drying of the corneas during the procedure.</li>
	<li>Disinfect the surgical area using three alternating applications of povidone-iodine and isopropanol wipes.</li>
	<li>Inject the analgesic(s) subcutaneously. 
	NOTE: Buprenorphine is generally used at a dose of 0.01-0.05&#160;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&#39;s IACUC for their guidelines regarding pain management.</li>
	<li>Inject 100 &#181;L of 1% lidocaine subcutaneously above the L2 to L6 spinous processes to provide local anesthesia.</li>
	<li>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.</li>
	<li>Place a fenestrated drape (see Table of Materials) on the animal, centering the fenestration over the lumbar spine.</li>
</ol>
5. Exposing the lumbar spine
<ol>
	<li>Confirm the depth of anesthesia by pinching each of the animal&#39;s paws and looking for the absence of a withdrawal response.</li>
	<li>Using a #11 scalpel blade, create an incision of approximately 3 cm long in the skin down the midline from L2 to L6.</li>
	<li>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.</li>
	<li>Remove the fascia covering the L2-L5 spinous processes.</li>
</ol>
6. Loading of the syringe
<ol>
	<li>Pipette 25-35 &#181;L of the vector (to achieve the desired dose) into the cap of a sterile microcentrifuge tube.</li>
	<li>Draw the entire volume into the insulin syringe. 
	NOTE: Take care not to draw up air during this process.</li>
</ol>
7. Performing the injection
<ol>
	<li>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.</li>
	<li>Place one hand so that the thumb rests gently on the animal&#39;s tail and one leg. Use a finger to steady the syringe.</li>
	<li>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&#176; off midline and 30&#176; 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.</li>
	<li>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.</li>
	<li>Depress the plunger slowly for about 5 s. 
	NOTE: There may be a twitch in the leg or tail during the injection.</li>
	<li>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.</li>
	<li>Slowly remove the needle.</li>
</ol>
8. Closing of the incision
<ol>
	<li>Approximate the edges of the incision.</li>
	<li>Beginning at one end of the wound, use a 4-0 suture (see Table of Materials) or surgical staples to close the incision.</li>
</ol>
9. Recovery and monitoring
<ol>
	<li>Place the animal in the prewarmed cage.</li>
	<li>Check the animal at least every 15 min until it is fully ambulatory. 
	NOTE: This should take between 15 min and 45 min.</li>
	<li>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.</li>
	<li>One week after surgery, remove the sutures or staples.</li>
</ol>
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.
<ol>
	<li>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.</li>
	<li>Once respiration and cardiac activity cease, open the chest cavity to ensure death.&#160;Extend the surgical incision up the back to the neck.</li>
	<li>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.</li>
	<li>Using fine forceps or scissors, remove the muscle from in between the spinous processes.</li>
	<li>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.</li>
	<li>Insert the lower tip of the rongeur under the L5 lamina and remove the bone overlying the spinal cord by taking several &#34;bites&#34; 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.</li>
	<li>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.</li>
</ol>

Lumbar Puncture Delivery of AAV9 in Juvenile Rats for CNS Gene Therapy

<ol>
	<li>Wurster, C., Petri, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progress+in+spinal+muscular+atrophy+research.">Progress in spinal muscular atrophy research.</a> Curr Opin Neurol. 35 (5), 693-698 (2022).</li><li>Wu, K. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retinitis+pigmentosa:+Novel+therapeutic+targets+and+drug+development.">Retinitis pigmentosa: Novel therapeutic targets and drug development.</a> Pharmaceutics. 15 (2), 685 (2023).</li><li>Larkin, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=First+FDA-approved+gene+therapy+for+hemophilia.">First FDA-approved gene therapy for hemophilia.</a> JAMA. 329 (1), 14 (2023).</li><li>Lee, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nadofaragene+firadenovec:+First+approval.">Nadofaragene firadenovec: First approval.</a> Drugs. 83 (4), 353-357 (2023).</li><li>Taghian, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+safe+and+reliable+technique+for+CNS+delivery+of+AAV+vectors+in+the+cisterna+magna.">A safe and reliable technique for CNS delivery of AAV vectors in the cisterna magna.</a> Mol Ther. 28 (2), 411-421 (2020).</li><li>Donsante, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracerebroventricular+delivery+of+self-complementary+adeno-associated+virus+serotype+9+to+the+adult+rat+brain.">Intracerebroventricular delivery of self-complementary adeno-associated virus serotype 9 to the adult rat brain.</a> Gene Ther. 23 (5), 401-407 (2016).</li><li>Pellot, J. E., Jesus, O. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Suboccipital+puncture.+[Updated+2022+Jul+25].">Suboccipital puncture. [Updated 2022 Jul 25].</a> StatPearls [Internet]. , (2022).</li><li>Elliger, S. S., Elliger, C. A., Aguilar, C. P., Raju, N. R., Watson, G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elimination+of+lysosomal+storage+in+brains+of+MPS+vii+mice+treated+by+intrathecal+administration+of+an+adeno-associated+virus+vector.">Elimination of lysosomal storage in brains of MPS vii mice treated by intrathecal administration of an adeno-associated virus vector.</a> Gene Ther. 6 (6), 1175-1178 (1999).</li><li>De La Calle, J. L., Paino, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+procedure+for+direct+lumbar+puncture+in+rats.">A procedure for direct lumbar puncture in rats.</a> Brain Res Bull. 59 (3), 245-250 (2002).</li><li>O'connor, D. M., Lutomski, C., Jarrold, M. F., Boulis, N. M., Donsante, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lot-to-lot+variation+in+adeno-associated+virus+serotype+9+(AAV9)+preparations.">Lot-to-lot variation in adeno-associated virus serotype 9 (AAV9) preparations.</a> Hum Gene Ther Methods. 30 (6), 214-225 (2019).</li><li>Manfredsson, F. P., Rising, A. C., Mandel, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AAV9:+A+potential+blood-brain+barrier+buster.">AAV9: A potential blood-brain barrier buster.</a> Mol Ther. 17 (3), 403-405 (2009).</li><li>Hudry, E., Vandenberghe, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+AAV+gene+transfer+to+the+nervous+system:+A+clinical+reality.">Therapeutic AAV gene transfer to the nervous system: A clinical reality.</a> Neuron. 101 (5), 839-862 (2019).</li><li>Georg-Fries, B., Biederlack, S., Wolf, J., Zur Hausen, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+proteins,+helper+dependence,+and+seroepidemiology+of+a+new+human+parvovirus.">Analysis of proteins, helper dependence, and seroepidemiology of a new human parvovirus.</a> Virology. 134 (1), 64-71 (1984).</li><li>Schulz, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Binding+and+neutralizing+anti-aav+antibodies:+Detection+and+implications+for+rAAV-mediated+gene+therapy.">Binding and neutralizing anti-aav antibodies: Detection and implications for rAAV-mediated gene therapy.</a> Mol Ther. 31 (3), 616-630 (2023).</li><li>Gray, S. J., Nagabhushan Kalburgi, S., Mccown, T. J., Samulski, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+CNS+gene+delivery+and+evasion+of+anti-aav-neutralizing+antibodies+by+intrathecal+aav+administration+in+non-human+primates.">Global CNS gene delivery and evasion of anti-aav-neutralizing antibodies by intrathecal aav administration in non-human primates.</a> Gene Ther. 20 (4), 450-459 (2013).</li><li>Meyer, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+single+injection+CSF+delivery+of+AAV9-mediated+gene+therapy+for+sma:+A+dose-response+study+in+mice+and+non-human+primates.">Improving single injection CSF delivery of AAV9-mediated gene therapy for sma: A dose-response study in mice and non-human primates.</a> Mol Ther. 23 (3), 477-487 (2015).</li><li>Hinderer, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+intrathecal+routes+of+administration+for+adeno-associated+viral+vectors+in+large+animals.">Evaluation of intrathecal routes of administration for adeno-associated viral vectors in large animals.</a> Hum Gene Ther. 29 (1), 15-24 (2018).</li><li>Wang, Y. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cerebrospinal+fluid+leakage+and+headache+after+lumbar+puncture:+A+prospective+non-invasive+imaging+study.">Cerebrospinal fluid leakage and headache after lumbar puncture: A prospective non-invasive imaging study.</a> Brain. 138, 1492-1498 (2015).</li><li>Hordeaux, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adeno-associated+virus-induced+dorsal+root+ganglion+pathology).">Adeno-associated virus-induced dorsal root ganglion pathology).</a> Hum Gene Ther. 31 (15-16), 808-818 (2020).</li><li>Semple, B. D., Blomgren, K., Gimlin, K., Ferriero, D. M., Noble-Haeusslein, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+development+in+rodents+and+humans:+Identifying+benchmarks+of+maturation+and+vulnerability+to+injury+across+species.">Brain development in rodents and humans: Identifying benchmarks of maturation and vulnerability to injury across species.</a> Prog Neurobiol. 106-107, 1-16 (2013).</li><li>Fang, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+adeno-associated+virus+serotypes+and+delivery+methods+for+cardiac+gene+transfer.">Comparison of adeno-associated virus serotypes and delivery methods for cardiac gene transfer.</a> Hum Gene Ther Methods. 23 (4), 234-241 (2012).</li></ol>

Dr. Donsante is an inventor on a pending patent regarding CSF administration of AAV9 vectors.

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 also a barrier. Environmental exposure to the various AAV serotypes increases with age13 and often leads to the production of antibodies that can neutralize therapeutic vectors14. Therefore, intravenous delivery of gene therapy vectors for CNS disorders is generally limited to infants and is not used in patients diagnosed later in life. 
For older patients, direct vector injection into the CSF can yield broad transduction in the CNS, bypassing both the BBB and preexisting anti-AAV antibodies15. Since this approach is targeted, lower vector doses can also be used. There are two primary approaches in the clinical setting: (1) lumbar puncture and (2) injection into the lateral ventricles. The latter carries more risk, but generally provides greater brain transduction. Lumbar puncture is safer, but transduction is skewed towards the spinal cord. Brain transduction might be enhanced by placing the patient into the Trendelenburg position, but data on this are mixed16,17. The use of a catheter to reach the cisterna magna via a lumbar puncture may provide a better option in the clinic, but it is in an early stage of use5. There may be other challenges to translating approaches worked out in animal models to the clinic, such as vector loss due to CSF leakage18 and toxicity in the dorsal root ganglia19. 
Most studies of CNS-directed therapies performed in rodents use neonates or adult animals (&gt;60 days of age). Neonates have the benefit of a small body size, allowing for higher effective doses, and an immature immune system, avoiding the complications of an immune response against the therapeutic. However, in terms of brain development, a newborn mouse or rat better represents a fetal stage in humans. For therapies intended for children in the 5-10 year age range, the juvenile rat (25-35 days old) is a better model in terms of neurological development20. Since a method for intrathecal injection had not been previously described for juvenile rats, and methods established for adult mice and rats proved to be ineffective in rats at this age, the approach described above was developed. To be clear, juvenile rats are not only smaller than adults but may also differ in the elasticity of the dura that protects the spinal cord, making a procedure that works to puncture this layer in an adult rat ineffective in a juvenile.
When learning how to perform intrathecal injection in juvenile rats, using a dye (such as trypan blue) as a surrogate for the therapeutic is necessary, and the user should be highly confident in their ability to successfully and reproducibly perform the procedure prior to starting a study with a therapeutic. Becoming proficient in the technique will require practice to get experience with how the syringe feels when the trajectory is on-target versus off-target. There are two common errors. If the angle of approach is too shallow, the needle will strike the top of one of the laminae or the back of the rostral lamina. There will be no twitch, and the distance that the needle advances will be a few millimeters short of 8 mm. If the angle of approach is too great, there is a risk that the needle will pass between the two laminae and penetrate the abdominal cavity. When this happens, the needle will advance much farther than 8 mm. If this happens, remove the needle, reposition, and try again. On the few occasions that this has happened, transiently entering the abdominal cavity by a few millimeters before withdrawal and repositioning has not caused any apparent lasting harm to the animals. 
It has been found that observing a physical response to the placement of the needle is critical to achieving reproducibility with a high rate of success with this procedure. When there was no response, the success rate for the injection was low. However, in some cases, an animal required attempts at multiple sites to achieve a response, and trace amounts of dye were observed in one or more of the previous needle tracks. No dye was observed in the epidural space, suggesting that some of the previous needle sticks had penetrated the dura without producing a tail or leg twitch. Since the reflux was minimal (similar to what is observed in the needle track from the injection), it is thought that the effect of previous needle sticks on delivery efficacy in these instances was negligible. 
Once one achieves proficiency in the delivery technique, a second, non-surgical challenge may be encountered. Specifically, in adult rats (~70 days of age), the potency of AAV9 vectors for intrathecal delivery to the spinal cord and brain can vary substantially from lot to lot, even when they are generated by the same vector core. Some batches will perform as expected, yielding transduction in the spinal cord gray matter along its length. Others, though, will fail to penetrate the gray matter, primarily transducing dorsal root ganglia10. The cause for this variability is unclear, as the vectors are potent in vitro and when injected directly into the spinal cord. It is recommended that a pilot study of 3-4 animals be performed with any new batch of virus to confirm that the new lot performs as expected before beginning a large study. Potency can be assessed using either immunohistochemical or immunofluorescence staining of the protein transgene product or quantifying the amount of transgene mRNA or vector genomes using quantitative PCR or ddPCR21. In addition to the unknown variables that distinguish viral lots, small differences in animal age, injection volume, speed of delivery, and vector concentration may cause variability in results. Before beginning a large study, they may need to be optimized for each virus or other candidate therapeutic agent.
Once trained, an experienced surgeon can complete the intrathecal injection procedure of a juvenile rat within about 30 min, from anesthesia induction to the beginning of the recovery period. This allows for large cohorts to be treated in a short amount of time. Recovery from the surgery is also rapid. Most animals ambulate normally within 20-30 min. After performing more than 200 of these surgeries, no adverse effects from this procedure have been encountered. 
Finally, minimizing animal distress and ensuring animal welfare during surgical procedures are paramount considerations. Thus, the proper use of anesthetics and analgesics is required, and body temperature must be maintained during the procedure and until the animal fully recovers from anesthesia. The relevant regulatory bodies and veterinary staff at different institutions may have different requirements and recommendations regarding these topics. The use of anesthetics and analgesics described in this procedure was developed in consultation with the Emory University veterinarians and IACUC staff. Researchers should work closely with their local veterinarians and IACUC to meet the needed goals.
There are certain limitations to this procedure. The method described here was developed for use in juvenile rats, and myriad structural and other differences between humans and rats may limit the translation of these procedures to humans. The point of enabling lumbar intrathecal injection of a therapeutic in juvenile rats is to facilitate the use of the juvenile rat model for testing the efficacy of the candidate therapeutic treatment - even if the precise mode of delivery would need to be altered for application in patients.

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 more1,2,3,4. Countless other therapeutics are currently in development. Gene therapy aims to deliver a therapeutic gene to a patient&#39;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 mice8,9. 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.

<table><tbody><tr><td>200 &amp;micro;L filtered pipette tips</td><td>MidSci</td><td>PR-200RK-FL</td><td>Pipetting virus</td></tr><tr><td>AAV9-GFP</td><td>Vector Builder</td><td>P200624-1005ynr</td><td>AAV9 vector expressing GFP</td></tr><tr><td>Absorbable Suture with Needle Coated Vicryl Polyglactin 910 FS-2 3/8 Circle Reverse Cutting Needle Size 4 - 0 Braided</td><td>McKesson</td><td>J422H</td><td>Suture</td></tr><tr><td>Bench pad</td><td>VWR</td><td>56616-031</td><td>Surgery</td></tr><tr><td>Braintree Scientific Isothermal Pads, 8&#039;&#039; x 8&#039;&#039;</td><td>Fisher Scientific</td><td>50-195-4664</td><td>Maintains body temperature</td></tr><tr><td>Buprenorphine</td><td>McKesson</td><td>1013922</td><td>Analgesic</td></tr><tr><td>Buprenorphine-ER (1 mg/mL)</td><td>Zoopharma</td><td></td><td>Extended-release analgesic</td></tr><tr><td>Cotton swabs</td><td>Fisher Scientific</td><td>19-365-409</td><td>Blood removal</td></tr><tr><td>Drape, Mouse, Clear Plastic, 12&quot; x 12&quot;, with Adhesive Fenestration</td><td>Steris</td><td>1212CPSTF</td><td>Surgical drape</td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11251-20</td><td>Forceps</td></tr><tr><td>Electric Blanket</td><td>CVS Health</td><td></td><td>CVS Health Series 500 Extra Long Heating Pad</td></tr><tr><td>Eppendorf Research plus, 1-channel pipette, variable, 20&amp;ndash;200 &amp;micro;L</td><td>Eppendorf</td><td>3123000055</td><td>Pipetting virus</td></tr><tr><td>Fine Scissors</td><td>Fine Science Tools</td><td>14059-11</td><td>Curved surgical scissors</td></tr><tr><td>Friedman-Pearson Rongeurs</td><td>Fine Science Tools</td><td>16121-14</td><td>Laminectomy</td></tr><tr><td>Halsey Needle Holders</td><td>Fine Science Tools</td><td>12001-13</td><td>Needle driver</td></tr><tr><td>Insulin Syringes with Ultra-Fine Needle 12.7 mm x 30 G 3/10 mL/cc</td><td>BD</td><td>328431</td><td>Syringe</td></tr><tr><td>Isoflurane</td><td>McKesson</td><td>803250</td><td>Anesthetic</td></tr><tr><td>Isopropanol wipes</td><td>Fisher Scientific</td><td>22-031-350</td><td>Skin disinfection</td></tr><tr><td>Lidocaine, 1%</td><td>McKesson</td><td>239935</td><td>Local anesthesia</td></tr><tr><td>Microcentrifuge Tubes: 1.5mL</td><td>Fisher Scientific</td><td>05-408-137</td><td>Loading the syringe</td></tr><tr><td>Povidone-iodine</td><td>Fisher Scientific</td><td>50-118-0481</td><td>Skin disinfection</td></tr><tr><td>Scalpel Handle - #4</td><td>Fine Science Tools</td><td>10004-13</td><td>Scalpel blade holder</td></tr><tr><td>Sure-Seal Induction Chamber</td><td>Braintree Scientific</td><td>EZ-17</td><td>Anesthesia box</td></tr><tr><td>Surgical Blade Miltex Carbon Steel No. 11 Sterile Disposable Individually Wrapped</td><td>McKesson</td><td>4-111</td><td>#11 Scalpel blade</td></tr><tr><td>SYSTANE NIGHTTIME Eye Ointment</td><td>Alcon</td><td></td><td>Eye ointment</td></tr><tr><td>Trypan Blue</td><td>VWR</td><td>97063-702</td><td>Injection</td></tr></tbody></table>

intrathecal vector delivery in juvenile rats via lumbar cistern injection

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 &#181;L 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.

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. However, when the needle fails to penetrate the dura mater, the dye ends up in the epidural space. Both the dura mater and the surrounding tissues (the surface of the bone and the ligaments and muscles connecting the laminae) will be stained blue. These patterns are easily visible to the naked eye.
The difference between correctly and incorrectly administered injection is difficult to assess if one simply makes a transverse cut through the spinal cord and spinal column. Instead, performing a laminectomy using a pair of rongeurs beginning at the L5 lamina and moving rostrally is recommended. Care must be taken not to damage the dura mater in the process. Using the dissecting microscope makes this process easier. Figure 1 provides comparative examples of successful injections and only partially successful injections. With a successful injection, little to no reflux along the needle track is observed when the needle is withdrawn. Following a successful injection, removal of the lamina to expose the spinal cord reveals trypan blue within the spinal cord but not on the surface of the bone (Figure 1A). The dye is also visible on the brainstem and cerebellum following a successful injection (Figure 1B). In contrast, a partially successful injection is evidenced by significant reflux of dye back up the needle tract during the injection process and/or visible evidence of dye on the bone (Figure 1C).
Using the above procedure, 28 &#181;L of an AAV9 vector expressing enhanced green fluorescent protein (GFP) was injected at a concentration of 3 x 1013 vector genomes/mL, for a total dose of 8.4 x 1011 vector genomes/animal. Four weeks later, the animals were euthanized and perfused with 4% paraformaldehyde10. The brain and spinal cord were then harvested and prepared for frozen sectioning. 40 &#181;m thick sections were obtained and stained for GFP. Figure 2 provides examples of the transduction pattern obtained with this vector. The transduction was generally highest in the spinal cord, particularly in the lumbar region (Figure 2A-C), likely due to its proximity to the injection site. The transduction of the brain was achieved (Figure 2D-F), but, as expected, it tended to be more limited than what was seen in the spinal cord.
To illustrate the reproducibility of results achieved by a single, experienced surgeon using a single lot of virus, stained sections of the cerebellum and cortex from each of the 15 rats injected for this study are presented in Figure 3 and Figure 4, respectively. The stained sections of the cervical spinal cord are also presented for 7 of these 15 rats (Figure 5). Of course, the amount of brain transduction may show even greater variability with different doses, vector lots, and surgeons10.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66463/66463fig01.jpg" /> 
Figure 1: Exposure of the spinal cord following an injection of the dye. Laminectomies were performed following the injection of trypan blue. (A) The spinal cord is stained blue in a correctly administered injection, and no dye is seen on the bone removed during the laminectomy (arrows). (B) The dye can also be observed around the brainstem. (C) In an injection where there was significant reflux during the injection, there is less dye within the cord, and dye is present on the surface of the bone (arrows). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66463/66463fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66463/66463fig02.jpg" /> 
Figure 2: Examples of transduction patterns achieved following intrathecal delivery of AAV9-GFP. 40 &#181;m thick sections of (A) cervical, (B) thoracic, and (C) lumbar spinal cord were immunohistochemically stained for GFP (black stain). High levels of gray matter transduction were observed at all levels. (D) In contrast, the brain exhibits sparser overall transduction. The left and right boxes are magnified in (E) and (F), respectively. (E) The majority of the staining is observed in the cerebellum, primarily in Purkinje neurons (arrows). (F) Neurons (arrows) and astrocytes (arrowheads) are transduced within the cerebral cortex. Scale bars: (A-C) 325 &#181;m; (D) 5 mm; and (E,F) 200 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66463/66463fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66463/66463fig03v2.jpg" /> 
Figure 3: Reproducibility of transduction in the cerebellum. Reproducibility of transduction in the cerebellum of 15 rats injected by the same surgeon using the same dose and lot of the virus. Scale bar: 1 mm. The numbers in the images indicate the rat ID numbers (&#39;litter&#39;.&#39;individual&#39;). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66463/66463fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66463/66463fig04.jpg" /> 
Figure 4: Reproducibility of transduction in the cortex. Reproducibility of transduction in the cortex of 15 rats injected by the same surgeon using the same dose and lot of virus. Scale bar: 500 &#181;m. The numbers in the images indicate the rat ID numbers (&#39;litter&#39;.&#39;individual&#39;). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66463/66463fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66463/66463fig05.jpg" /> 
Figure 5: Reproducibility of transduction in the cervical spinal cord. Reproducibility of transduction in the cervical spinal cord of 7 rats injected by the same surgeon using the same dose and lot of virus. Scale bar: 500 &#181;m. The numbers in the images indicate the rat ID numbers (&#39;litter&#39;.&#39;individual&#39;). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66463/66463fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats at JoVE.com

Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats

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.

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

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 ...

intrathecal-vector-delivery-juvenile-rats-via-lumbar-cistern

Nanyang Technological University

Research

JoVE Journal

Neuroscience

919 Views.  Emory University. 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. 

Video: Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats

Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most often results in failure of potential therapeutics designed for neurodegenerative disorders 1,2. Neurodegenerative diseases such as Spinal Muscular Atrophy (SMA), in which the lower motor neurons are affected, can benefit greatly from introducing the therapeutic agents into the CNS. The purpose of this video is to demonstrate two different injection paradigms to deliver therapeutic materials into neonatal mice soon after birth. One of these methods is injecting directly into cerebral lateral ventricles (Intracerebroventricular) which results in delivery of materials into the CNS through the cerebrospinal fluid 3,4. The second method is a temporal vein injection (intravenous) that can introduce different therapeutics into the circulatory system, leading to systemic delivery including the CNS 5. Widespread transduction of the CNS is achievable if an appropriate viral vector and viral serotype is utilized. Visualization and utilization of the temporal vein for injection is feasible up to postnatal day 6. However, if the delivered material is intended to reach the CNS, these injections should take place while the blood brain barrier is more permeable due to its immature status, preferably prior to postnatal day 2. The fully developed blood brain barrier greatly limits the effectiveness of intravenous delivery. Both delivery systems are simple and effective once the surgical aptitude is achieved. They do not require any extensive surgical devices and can be performed by a single person. However, these techniques are not without challenges. The small size of postnatal day 2 pups and the subsequent small target areas can make the injections difficult to perform and initially challenging to replicate.

Delivery of Therapeutic Agents Through Intracerebroventricular ICV and Intravenous IV Injection in Mice

This article demonstrates two very different methods of injection: 1) into the brain (intracerebroventricular) and 2) systemic (intravenous) to introduce the therapeutic agents into the central nervous system of neonatal mice.

Despite the protective role that blood brain barrier plays in shielding the brain, it limits the access to the central nervous system (CNS) which most ...

Medicine

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. Using subpially-placed polyethylene catheters (PE-10 or PE-5) for AAV9 delivery, potent transgene expression through the spinal parenchyma (white and gray matter) in subpially-injected spinal segments has been demonstrated. Because of the wide range of transgenic mouse models of neurodegenerative diseases, there is a strong desire for the development of a potent central nervous system (CNS)-targeted vector delivery technique in adult mice. Accordingly, the present study describes the development of a spinal subpial vector delivery device and technique to permit safe and effective spinal AAV9 delivery in adult C57BL/6J mice. In spinally immobilized and anesthetized mice, the pia mater (cervical 1 and lumbar 1-2 spinal segmental level) was incised with a sharp 34 G needle using an XYZ manipulator. A second XYZ manipulator was then used to advance a blunt 36G needle into the lumbar and/or cervical subpial space. The AAV9 vector (3-5 &#181;L; 1.2 x 1013 genome copies (gc)) encoding green fluorescent protein (GFP) was then injected subpially. After injections, neurological function (motor and sensory) was assessed periodically, and animals were perfusion-fixed 14 days after AAV9 delivery with 4% paraformaldehyde. Analysis of horizontal or transverse spinal cord sections showed transgene expression throughout the entire spinal cord, in both gray and white matter. In addition, intense retrogradely-mediated GFP expression was seen in the descending motor axons and neurons in the motor cortex, nucleus ruber, and formatio reticularis. No neurological dysfunction was noted in any animals. These data show that the subpial vector delivery technique can successfully be used in adult mice, without causing procedure-related spinal cord injury, and is associated with highly potent transgene expression throughout the spinal neuraxis.

Subpial Adeno-associated Virus 9 AAV9 Vector Delivery in Adult Mice

The goal of the present study was to develop and validate the potency and safety of spinal adeno-associated virus 9 (AAV9)-mediated gene delivery by using a novel subpial gene delivery technique in adult mice.

The successful development of a subpial adeno-associated virus 9 (AAV9) vector delivery technique in adult rats and pigs has been reported on previously. ...

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, and excellent transduction efficacy in the CNS.&#160;Previous studies have demonstrated the therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration. However, high rates of unpredictable failure due to the technical limitation of IT administration in small animals have been reported. Here, we established a scoring system to indicate the success extent of lumbar puncture in small animals by adding 1% lidocaine hydrochloride into the injection solution. We further show that the extent of transient weakness following injection can predict the transduction efficiency of AAV. Thus, this IT injection method can be used to optimize therapeutic trials in mouse models of CNS diseases that afflict wide regions of the CNS.

Here we present a direct intrathecal injection technique using 1% lidocaine hydrochloride in a viral solution to ensure efficient adeno-associated virus delivery to small animals and establish a scoring system to predict transduction efficiency in the central nervous system according to the degree of transient weakness induced by lidocaine.

Intrathecal (IT) injection of adeno-associated virus (AAV) has drawn considerable interest in CNS gene therapy by virtue of its safety, noninvasiveness, ...

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. Injection directly into spinal cord tissue bypasses these barriers, providing access to cell bodies or synapses where molecules can be incorporated. Combining viral vector technology with this method allows for introduction of target genes into nervous tissue for the purpose of gene therapy or tract tracing. Here a virus engineered for highly efficient retrograde transport (HiRet) is introduced at the synapses of propriospinal interneurons (PNs) to encourage specific transport to neurons in the spinal cord and brainstem nuclei. Targeting PNs takes advantage of the numerous connections they receive from motor pathways such as the rubrospinal and reticulospinal tracts, as well as their interconnection with each other throughout spinal cord segments. Representative tracing using the HiRet vector with constitutively active green fluorescent protein (GFP) shows high fidelity details of cell bodies, axons and dendritic arbors in thoracic PNs and in reticulospinal neurons in the pontine reticular formation. HiRet incorporates well into brainstem pathways and PNs but shows age dependent integration into corticospinal tract neurons. In summary, spinal cord injection using viral vectors is a suitable method for introduction of proteins of interest into neurons of targeted tracts.

This protocol demonstrates injection of a retrogradely transportable viral vector into rat spinal cord tissue. The vector is taken up at the synapse and transported to the cell body of target neurons. This model is suitable for retrograde tracing of important spinal pathways or targeting cells for gene therapy applications.

Introducing proteins of interest into cells in the nervous system is challenging due to innate biological barriers that limit access to most molecules. ...

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

The blood brain barrier (BBB) is an important defense against the entrance of potentially toxic or pathogenic agents from the blood into the central nervous system (CNS). However, its existence also dramatically lowers the accessibility of systemically administered therapeutic agents to the CNS. One method to overcome this, is to inject those agents directly into the cerebrospinal fluid (CSF), thus bypassing the BBB. This can be done via implantation of a catheter for either continuous infusion using an osmotic pump, or for single bolus delivery. In this article, we describe a surgical protocol for delivery of CNS-targeting antisense oligonucleotides (ASOs) via a catheter implanted directly into the cauda equina space of the adult rat spine. As representative results, we show the efficacy of a single bolus ASO intrathecal (IT) injection via this catheterization system in knocking down the target RNA in different regions of the rat CNS. The procedure is safe, effective and does not require expensive equipment or surgical tools. The technique described here can be adapted to deliver drugs in other modalities as well.

Here, we describe a method for delivering drugs to the rat central nervous system by implanting a catheter into the lumbar intrathecal space of the spine. We focus on the delivery of antisense oligonucleotides, though this method is suitable for delivery of other therapeutic modalities as well.

The blood brain barrier (BBB) is an important defense against the entrance of potentially toxic or pathogenic agents from the blood into the central ...

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the underlying parenchyma. Previously published protocols recommend using straight needles that should be lowered to a maximum of 1-2 mm from the dural surface. The sudden drop in resistance once the dural membrane has been punctured makes it difficult to maintain the needle in a steady position. Our method, instead, employs a needle bent at the tip that can be stabilized against the occipital bone of the skull, thus preventing the syringe from penetrating into the tissue after perforation of the dural membrane. The procedure is straightforward, reproducible, and does not cause long-lasting discomfort in the operated animals. We describe the intracisternal injection strategy in the context of genetic fate mapping of vascular leptomeningeal cells. The same technique can, furthermore, be utilized to address a wide range of research questions, such as probing the role of leptomeninges in neurodevelopment and the spreading of bacterial meningitis, through genetic ablation of genes putatively implicated in these phenomena. Additionally, the procedure can be combined with an automatized infusion system for a constant delivery and used for tracking cerebrospinal fluid movement via injection of fluorescently labelled molecules.

We describe an intracisternal injection that employs a needle bent at the tip that can be stabilized to the skull, thus eliminating the risk of damage to the underlying parenchyma. The approach can be used for genetic fate mapping and manipulations of leptomeningeal cells and for tracking cerebrospinal fluid movement.

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the ...

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Biology

Targeting the Corticospinal Tract in Neonatal Rats with a Double-Viral Vector using Combined Brain and Spine Surgery

Successfully tackling the obstacles that constrain research on neonatal rats is important for studying the differences in outcomes seen in pediatric spinal cord injuries (SCIs) compared to adult SCIs. In addition, reliably introducing therapies into the target cells of the central nervous system (CNS) can be challenging, and inaccuracies can compromise the efficacy of the study or therapy. This protocol combines viral vector technology with a novel surgical technique&#160;to accurately introduce gene therapies into neonatal rats at postnatal day 5. Here, a virus engineered for retrograde transport (retroAAV2) of Cre is introduced at the axon terminals of corticospinal neurons in the spinal cord, where it is subsequently transported to the cell bodies. A double-floxed&#160;inverted orientation (DIO) designer receptor exclusively activated by designer drug(s) (DREADD) virus is then injected into the somatomotor cortex of the brain. This double-infection technique promotes the expression of the DREADDs only in the co-infected corticospinal tract (CST) neurons.&#160;Thus, the simultaneous co-injection of the somatomotor cortex and cervical CST terminals is a valid method for studying the chemogenetic modulation of recovery following cervical SCI models in neonatal rats.

This protocol demonstrates a novel method for applying gene therapies to subpopulations of cells in neonatal rats at postnatal ages 5-10 days by injecting an anterograde chemogenetic modifier into the somatomotor cortex and a retrogradely transportable Cre recombinase into the cervical spinal cord.

Successfully tackling the obstacles that constrain research on neonatal rats is important for studying the differences in outcomes seen in pediatric ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

A Comprehensive Protocol for Delivering Intrathecal Injections in Neonatal Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and treatments. By directly delivering medications and treatments into the cerebrospinal fluid of the central nervous system, this method achieves higher localized drug concentrations while reducing systemic side-effects compared to other routes such as intravenous, subcutaneous, or intramuscular injections. Its importance extends beyond clinical settings, as intrathecal injection plays a vital role in preclinical studies focused on treating neurogenetic disorders in rodents and other large animals, including non-human primates. However, despite its widespread application, intrathecal injection in young, particularly neonatal pups, poses significant technical challenges due to their small size and fragile nature. Successful and reliable administration of intrathecal injections in newborn mice requires meticulous attention to detail and careful consideration of various factors. Thus, there is a crucial need for a standardized protocol that not only provides instructions but also highlights key technical considerations and good laboratory practices to ensure procedural consistency, as well as the safety and welfare of the animals.
To address this unmet need, we present a detailed and comprehensive protocol for performing intrathecal injections specifically in newborn pups on postnatal day 1 (P1). By following the step-by-step instructions, researchers can confidently perform intrathecal injections in neonatal pups, enabling the accurate delivery of drugs, antisense oligos, and viruses for gene replacement or genome editing-based treatments. Furthermore, the importance of adhering to good laboratory practices is emphasized to maintain the well-being of animals and ensure reliable experimental outcomes. This protocol aims to address the technical challenges associated with intrathecal injections in neonatal mice, ultimately facilitating advances in the field of neurogenetic research that aims to develop potential therapeutic interventions.

Author Spotlight: Intrathecal Injection &#8211; An Efficient and Reliable Delivery Method to Test the Efficacy of Gene Editing in Neonatal Mouse Brains

The present protocol outlines step-by-step instructions for performing intrathecal injections in neonatal mice for gene editing and drug delivery.

Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and ...

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice

Subpial Adeno-associated Virus 9 (AAV9) Vector Delivery in Adult Mice

Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice

Direct Injection of a Lentiviral Vector Highlights Multiple Motor Pathways in the Rat Spinal Cord

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

Targeting the Corticospinal Tract in Neonatal Rats with a Double-Viral Vector using Combined Brain and Spine Surgery

Intravenous Injections in Neonatal Mice

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery