Name: intrathecal delivery of antisense oligonucleotides in the rat central nervous system
Uploaded: 2019-10-29T14:00:00.000+00:00
Duration: 7 min 47 s
Description: Watch this Scientific Journal Video about Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System at JoVE.com

We would like to thank Ionis Pharmaceuticals for supplying the ASOs described in the article.

All in vivo procedures were performed under Biogen Institutional Animal Use and Care Committee (IACUC) approved protocols which follow the guidelines set forth by the United States National Institutes of Health guide for the care and use of laboratory animals.
1. Material and instrument preparation
<ol>
	<li>Prepare the special guide cannulas.
	<ol>
		<li>Use a rotary tool with cut-off wheel (or a sharp saw) to cut off the two ends of a 19 G needle, resulting in a ~1.5&#8722;2 cm long guide cannula (Figure 1Aiii). Use the grinding wheel of the rotary tool to smooth the two ends. 
		NOTE: Alternatively, premade and sterile guide cannulas can be purchased from a commercial vendor (Table of Materials).</li>
	</ol>
	</li>
	<li>Prepare the catheter/wire assembly.
	<ol>
		<li>Cut an 8 cm long piece of PE-10 tubing (polyethylene tubing, diameter 0.011 inch) to serve as the intrathecal catheter. Make a mark 2 cm from one end with an ethanol resistant marker pen. Cut an 11 cm long stylet wire from polytetrafluoroethylene coated stainless steel wire. Insert the stylet wire (Figure 1Aii) into the lumen of the PE-10 catheter (Figure 1Ai). 
		NOTE: One catheter/wire assembly set (Figure 1Av) is needed for each animal. Alternatively, catheters and stylet wires can be purchased from a commercial vendor (Table of Materials).</li>
	</ol>
	</li>
	<li>Prepare delivery catheter assembly.
	<ol>
		<li>Cut a piece of PE-50 catheter (5&#8722;10 cm, polyethylene tubing, diameter 0.023 inch) (Figure 1Bi). To one end of the PE50 catheter insert a 23 G tubing adaptor (Figure 1Biv). This end will be connected to a 100 &#181;L syringe (Table of Materials) during surgery. Insert a modified 30 G needle with hub cut off (Figure 1Biii) into an approximately 0.5 cm piece of PE-10 tubing (Figure 1Bii) and connect the PE-10 tubing to the other end of the catheter.</li>
		<li>The 30 G needle end of the delivery catheter assembly (Figure 1Bv) will be connected to the distal end of the implanted PE-10 catheter in the rat during surgery. 
		NOTE: Alternatively, the delivery catheter assembly (Figure 1Bv) can be purchased from a commercial vendor (Table of Materials).</li>
	</ol>
	</li>
	<li>Prepare the guide cannula-needle assembly (Figure 1Avi) by placing a guide cannula (Figure 1Aiii) over the end of a 23 G needle. </li>
	<li>Sterilize all surgical instruments including the guide cannula and the wire/catheter sets using an ethylene oxide sterilizer for 12 h. 
	NOTE: All surgical instruments except the catheter can be autoclaved; catheter will melt at high temperature.</li>
</ol>
2. Surgery preparation
NOTE: This procedure is routinely performed on male and female Sprague Dawley rats with body weights between 200 g and 400 g. Two rats are housed per cage under a 12 h light/dark cycle with free access to food and water.
<ol>
	<li>Weight a rat and place it in an isoflurane chamber to induce anesthesia (1&#8722;5% isoflurane in O2, titrated to effect). 
	NOTE: The rat is then continuously anesthetized with isoflurane to maintain deep anesthesia via a nose cone throughout the procedure. An alternative anesthesia method (e.g., administration of ketamine 100 mg/kg and xylazine 10 mg/kg) can be used as approved by the IACUC.</li>
	<li>When the rat fails to respond to a toe pinch, shave its back from the tail to the caudal thoracic spine and place the shaved rat on the a sterile sheet laid on top of a heating pad.</li>
	<li>Position a 50 ml conical centrifuge tube under the abdomen of the rat to flex the spine in the lumbar region (Figure 2A) and apply ophthalmic ointment to the eyes.</li>
	<li>Inject sustained release buprenorphine (1.0 mg/kg; Table of Materials) subcutaneously in the rat. Clean the exposed skin with alternating povidone and alcohol scrubs and repeat this three times. 
	NOTE: An alternative pain relief can be used as approved by the IACUC protocol.</li>
	<li>Drape the animal with a sterile transparent drape that has been fenestrated over the surgical site.</li>
</ol>
3. Surgery
<ol>
	<li>With the rat supported by the 50 mL conical tube, identify the two natural pits between muscles above the pelvis (arrows in Figure 2A). With one hand holding those pits, use the other hand to gently press and feel the spine from caudal to rostral direction and find the first major indentation between vertebrae and this is the intervertebral space between S1 and L6 vertebrae (Figure 2B).</li>
	<li>Move slightly rostrally to identify the next indentation, the intervertebral space between the L5 and L6 vertebrae and the injection site (* in Figure 2A). Use a scalpel to make an incision no more than 2 cm long in the skin along the midline from rostral to caudal so that the injection site is at the center of the incision (dotted line in Figure 2A).</li>
	<li>Use dissection scissors to dissect away the connective tissue in order to visualize the muscle layer. Then make a 1 cm incision in the muscle capsule immediately lateral to the dorsal spinal process of the L6 lumbar vertebra. 
	NOTE: The bones of the L6 lumbar vertebra could be visualized at this point.</li>
	<li>Position the guide cannula-needle assembly near the anterior aspect of the 6th lumbar vertebra and push it into the intervertebral space along the anterior aspect of the 6th vertebra so that the end of the needle penetrates the spinal canal. Push the guide cannula in place along the needle and remove the 23 G needle leaving only the guide cannula in place. 
	NOTE: It is helpful to use blunt forceps to locate the dorsal process of the L6 lumbar vertebra before inserting the needle. Generally, CSF fluid can be seen entering the needle hub (this fluid may be tinged with a hint of blood, but this does not indicate that harm has been done or that the needle is not placed correctly). The authors have not seen large amount of blood or severe bleeding during this procedure. If either occurs, a veterinarian should be contacted to determine the appropriate treatment and if animals should be euthanized.</li>
	<li>Insert the catheter-wire assembly into the guide cannula. Angle down the catheter-wire assembly at approximately a 45&#176; angle to the spinal canal and force the end approximately 0.3 cm into the spinal canal.</li>
	<li>Remove the guide cannula, leaving the catheter with the stylet wire in place. Remove the stylet wire approximately 2.5 cm from the intrathecal tip of the catheter and advance the catheter into the spinal canal until the 2 cm mark is at the entrance of the canal (just visible below the muscle) as shown in Figure 1Bvi. 
	NOTE: The inserted catheter should extend rostrally into the subarachnoid space. Successful placement should allow for free movement of the catheter in that space.</li>
	<li>Completely withdraw the stylet wire, and CSF may be seen entering the implanted catheter.</li>
	<li>Connect the delivery catheter assembly to the distal end of the implanted catheter via the 30 G needle end (Figure 1Bv,vi).</li>
	<li>Load 60 &#181;L of sterile saline into a 100 &#181;L syringe (flushing syringe). Load a bolus of 30 &#181;L of the test compound (e.g., ASO solution) into a second syringe (injection syringe).</li>
	<li>Connect the flushing syringe (loaded with saline) to the tubing adaptor end of the delivery catheter assembly (Figure 1Bv). Inject 20 &#181;L of sterile saline into the intrathecal space (pre-injection flushing).</li>
	<li>Connect the injection syringe (loaded with test compound) to the tubing adaptor end of the delivery catheter assembly (Figure 1Bv). Inject 30 &#181;L of test compound into the intrathecal space over 30 s. 
	NOTE: The routine injection volume of ASO is 30 &#181;L to achieve good knockdown in the spinal cord and in the cortex. It has been reported that the injection volumes may impact compound distribution12, though different injection volumes have not been tested. If other compound or volume is used, the safety and effectiveness need to be empirically determined.</li>
	<li>Repeat step 3.10 and flush the catheter with another 40 &#181;L of sterile saline (post-injection flushing). Then detach the delivery catheter assembly from the implanted catheter. 
	NOTE: The pre and post-injection flush is thought to reduce local sequestration of the compounds and improve their distribution to the rostral structures12.</li>
	<li>Aseptically cut and heat seal the implanted catheter: Place a pair of sterile dissection forceps in a bead sterilizer until they are very hot, then clamp down on the tubing with the hot end of the forceps. 
	NOTE: This action melts the catheter. Thus, the hole in the tube is collapsed and all sides stick to each other, sealing the tubing in an aseptic fashion and then it is placed into the subcutaneous space.</li>
	<li>Use absorbable monofilament sutures to secure the remaining heat-sealed catheter to the connective tissue. Then use non-absorbable monofilament sutures to close the skin over the secured heat-sealed catheter. 
	NOTE: Wound clips can also be used as approved by the IACUC protocol. The technique is compatible with repeated injections though it has only been used for one-time injections in our hands. The feasibility of repeated injections should be empirically evaluated with the approval of the IACUC.</li>
	<li>Use gauze and saline to wash any blood from the skin and allow the animal to recover from the anesthesia in a heated incubator until mobile, at which point it is returned to its home cage (two rats per cage). 
	NOTE: When performing surgeries on multiple rats on the same day, clean tools using water to remove blood and re-sterilize using a heated dry bead sterilizer (for at least 20 s, with time to cool) between animals. A new set of instruments is used every 5 animals.</li>
	<li>Monitor the animals daily for at least 3 days after surgery and continue to monitor the animals weekly after recovering from surgery according to the IACUC protocol. 
	NOTE: If any complications occur (urine retention, incision infections, neurological disorder such as seizure or paralysis), a veterinarian should be contacted to determine the appropriate treatment and if animals should be euthanized. If sustained release buprenorphine is not used, pain relief should be given daily after surgery according to the IACUC protocol.</li>
</ol>
4. Evaluation of tissue specific knockdown after IT injection
<ol>
	<li>Two weeks after IT bolus injection of ASOs, collect different regions of the brain (i.e., cerebral cortex, striatum and cerebellum) as well as different segments of the spinal cord (i.e., cervical, thoracic, and lumbar). Extract total tissue RNA using a commercial RNA extraction kit and perform cDNA synthesis reaction as described previously13. 
	NOTE: Standard reagents were used for qPCR with the following assays: rat Malat1 and rat GAPDH. The relative transcript levels were calculated using the 2-&#916;&#916;CT method (CT = threshold cycle).</li>
</ol>

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The authors are all employees of Biogen, Inc. or Ionis Pharmaceuticals. The authors receive the antisense oligonucleotides described in the article from Ionis Pharmaceuticals.

The present article shows a powerful method to deliver therapeutic agents directly into the rat CNS. In theory, a similar technique can be also performed in mice, though due to the smaller size, the method can be more challenging. Therefore, our group performs intracerebroventricular (ICV) injections in mice for CNS drug delivery, which reach the same goals through a different route of administration. That method has been described in another study16.
The advantage of t...

The vascular system of the central nervous system (CNS) has evolved as a critical regulator of homeostasis, controlling traffic of molecules, supplying nutrients and getting rid of waste. This system is also the first line of defense from attacks of external pathogens, thanks to a dense distribution of tight junctions along the walls of the endothelial cells. These tight junctions make up one aspect of the blood brain barrier (BBB). While the BBB allows the transport of molecules required to fulfill nutrient and energy demands (e.g., ions, glucose), it also selectively limits the passage of pathogens as well as toxic chemicals1,2,3.
Ironically, the same protective function of the BBB that limits passage of pathogens and toxic chemicals also is the major obstacle to our ability to easily access the CNS with therapeutic treatments after systemic administration to the organism2,4,5. This role of the BBB has prompted the development of a plethora of new drug distribution technologies and approaches6.
One way to overcome this obstacle is to inject the drugs directly into the cerebrospinal fluid (CSF) that continuously perfuses both the brain and spinal cord7,8,9,10. In this article, we describe a method to successfully deliver agents into the lumbar intrathecal space by placing the internal end of the catheter completely in the cauda equina space of the rat spine. A description of this procedure was previously published by Mazur et al. elsewhere11.
The protocol is very effective and produces a greater than 90% success rate of antisense oligonucleotide (ASO) delivery to the CNS when assessed by quantitative polymerase chain reaction (qPCR) analysis of target gene knockdown8. The procedure causes minimal discomfort to the animals, as 100% of the rats survive the surgery and show minimal swelling around the surgical wound and no signs of distress (e.g., hyperactivity, dehydration, circling, loss of balance, decreased food intake, and dehydration) during post-op observation. Another advantage of the method described here is that it does not require expensive equipment, nor any special tools.

<table><tbody><tr><td>3M Steri-Drape Small Drape with Adhesive Aperture</td><td>3M</td><td></td><td>1020</td></tr><tr><td>70% ethanol</td><td>Decon Laboratories, Inc</td><td></td><td>8416-160Z</td></tr><tr><td>Alcohol swab sticks</td><td>Dynarex</td><td></td><td>NO 1204</td></tr><tr><td>&lt;a href=&quot;https://punchout.fishersci.com/shop/products/bd-general-use-syringes-5/22124967?crossRef=BD%20302830#?keyword=BD+302830&quot;&gt;BD General Use Syringes 1 mL Luer-Lok tip&lt;/a&gt;</td><td>BD</td><td>1ml TB Luer-Lok tip</td><td>BD 302830</td></tr><tr><td>BD Intramedic PE Tubing</td><td>BD</td><td>Polyethylene tubing PE50 Diameter 0.023 in</td><td>BD 427400 (10ft, Fischer Scientific 22-204008) or 427401 (100ft, Fischer Scientific 14-170-12P)</td></tr><tr><td>BD Intramedic PE Tubing</td><td>BD</td><td>Polyethylene tubing PE10 Diameter 0.011 in</td><td>BD 427410 (10ft, Fischer Scientific 14-170-11B) or 4274011 (100ft, Fischer Scientific 14-170-12B)</td></tr><tr><td>BD Intramedic PE Tubing Adapters</td><td>BD</td><td>23 gauge intramedic luer stub adaper</td><td>BD 427565 or Fisher Scientific 14-826-19E</td></tr><tr><td>BD PrecisionGlide Single-use Needles 30G</td><td>BD</td><td></td><td>BD 305128</td></tr><tr><td>Buprenorphine Sustained Release-lab</td><td>ZooPharm</td><td>Prescription required</td><td></td></tr><tr><td>Ethylene oxide sterilizer</td><td>Andersen Sterilizer INC.</td><td>AN 74i, gas sterilizer</td><td>AN 74i</td></tr><tr><td>Guide cannula</td><td>BD</td><td>19G x 1 WT (1.1 mm x 25mm) needle</td><td>BD 305186</td></tr><tr><td>Hamilton syringe 100ul</td><td>Hamilton company</td><td>Hamilton syringe 100ul</td><td></td></tr><tr><td>Hot bead Sterilizer</td><td>Fine Science Tools</td><td></td><td>STERILIZER MODELNO FST 250</td></tr><tr><td>Ophthalmic ointment</td><td>Dechra veterranery product</td><td></td><td>17033-211-38</td></tr><tr><td>Pocket Pro Pet Trimmer</td><td>Braintree Scientific</td><td></td><td>CLP-9931 B</td></tr><tr><td>Povidone scrub</td><td>PDI</td><td></td><td>S48050</td></tr><tr><td>Saline</td><td>Baxter</td><td>Sodium Chloride 0.9% Intravenous Infusion BP 50ml</td><td>FE1306G</td></tr><tr><td>Scalpel</td><td>Feather</td><td>disposable scalpel</td><td>No. 10</td></tr><tr><td>Small animal heating pad</td><td>K&amp;amp;H Manufacturing</td><td></td><td>Model # 1060</td></tr><tr><td>Stylet Wire</td><td>McMaster-Carr</td><td>1749T14</td><td>LH-36233780</td></tr><tr><td>Surgery Towel drape</td><td>Dynarex</td><td></td><td>4410</td></tr><tr><td>Surgical scissors and forceps</td><td>FST and Fisher Scientific</td><td></td><td></td></tr><tr><td>Sutures</td><td>Ethicon</td><td></td><td>4-0 or 5-0</td></tr><tr><td>Tool to make the Guide cannular</td><td>Grainger</td><td>Rotary tool (Dremel)</td><td>14H446 (Mfr: EZ456)</td></tr><tr><td></td><td></td><td>EZ lock cut off Wheel</td><td>1PKX5 (Mfr: 3000-1/24)</td></tr><tr><td></td><td></td><td>Grinding Wheel, Aluminum Oxide</td><td>38EY44 (Mfr: EZ541GR)</td></tr><tr><td></td><td></td><td>EZ lock Mandrel</td><td>1PKX8 (Mfr: EZ402-01)</td></tr><tr><td></td><td></td><td>Diamond wheel floor Tile</td><td>3DRN4 (Mfr: EZ545)</td></tr><tr><td>&lt;strong&gt;Alternative source for pre-made and sterilized materials for this procedure&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Dosing catheter system</td><td>SAI Infusion Systems</td><td></td><td>RIDC-01</td></tr><tr><td>Guide cannula</td><td>SAI Infusion Systems</td><td></td><td>RIDC-GCA</td></tr><tr><td>Internal Catheters</td><td>SAI Infusion Systems</td><td></td><td>RIDC-INC</td></tr><tr><td>Stylet Wire</td><td>SAI Infusion Systems</td><td></td><td>RIDC-STY</td></tr></tbody></table>

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.

Using the method described here, we injected two groups of adult female rats (250&#8722;300 g; n = 10/group) with either a single bolus of phosphate-buffered saline PBS or 300 &#181;g of ASO targeting the long non-coding (linc) RNA Malat1; in our lab we routinely use the Malat1 ASO as a tool compound, because Malat1 is expressed ubiquitously and at high levels in all tissues14, including brain and spinal cord. The Malat1 ASO works via an RNaseH1-mediated mechanism<...

Watch this Scientific Journal Video about Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System at JoVE.com

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

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 ...

intrathecal-delivery-antisense-oligonucleotides-rat-central-nervous

Research

JoVE Journal

Neuroscience

18.3K Views.  Biogen, Inc.. 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.

Video: Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution in assessing therapeutic efficacy. Between the two major classifications of administration, local vs. systemic, systemic delivery approaches are often preferred due to ease of administration. However, systemic delivery may result in suboptimal drug concentration being achieved in the CNS, and lead to erroneous conclusions regarding agent efficacy. Local drug delivery methods are more invasive, but may be necessary to achieve therapeutic CNS drug levels. Here, we demonstrate proper technique for three routes of systemic drug delivery: intravenous injection, intraperitoneal injection, and oral gavage. In addition, we show a method for local delivery to the brain: convection-enhanced delivery (CED). The use of fluorescently-labeled compounds is included for in vivo imaging and verification of proper drug administration. The methods are presented using murine models, but can easily be adapted for use in rats.

Thorough preclinical testing of drugs that act in the central nervous system often involves assessing and comparing drug biodistribution in association with specific routes of administration. Here, three commonly used methods of systemic delivery (intravenous, intraperitoneal, and oral) as well as a method for local delivery (convection-enhanced delivery) are demonstrated in mice.

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Methods and Tips for Intravenous Administration of Adeno-associated Virus to Rats and Evaluation of Central Nervous System Transduction

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), optogenetics, cre-lox targeting, etc. The purpose of this manuscript is to aid the investigator attempting expansive central nervous system (CNS) gene transfer in the rat via tail vein injection of AAV. Wide-scale expression is relevant for conditions with widespread pathology, and a rat model is significant due to its greater size and physiologic similarities to humans compared to mice. In this example application, a wide-scale neuronal transduction is used to mimic a neurodegenerative disease that affects the entire spinal cord, amyotrophic lateral sclerosis (ALS). The efficient wide-scale CNS transduction can also be used to deliver therapeutic protein factors in pre-clinical studies. After a post-injection expression interval of several weeks, the effects of the transduction are evaluated. For a green fluorescent protein (GFP) control vector, the amount of GFP in the cerebellum is estimated quickly and reliably by a basic imaging program. For motor disease phenotypes that are induced by the ALS related protein transactive response DNA-binding protein of 43 kDa (TDP-43), the deficits are scored by escape reflex and rotarod. Beyond disease modeling and gene therapy, there are diverse potential applications for the wide-scale gene targeting described here. The expanded use of this method will aid in expediting hypothesis testing in the neurosciences and neurogenetics.

Methods for a wide-scale central nervous system gene delivery in the rat are covered. In this example, the purpose is to mimic a disease that affects the entire spinal cord. The widespread transduction can be used to deliver a therapeutic protein to the CNS from a one-time, peripheral administration.

Adeno-associated virus (AAV) vectors are a key reagent in the neurosciences for clustered regularly interspaced short palindromic repeats (CRISPR), ...

A Positioning Device for the Placement of Mice During Intranasal siRNA Delivery to the Central Nervous System

Intranasal (IN) drug delivery to the brain has emerged as a promising method to bypass the blood-brain barrier (BBB) for the delivery of drugs into the central nervous system (CNS). Recent studies demonstrate the use of a peptide, RVG9R, incorporating the minimal receptor-binding domain of the Rabies virus glycoprotein, in eliciting the delivery of siRNA into neurons in the brain. In this protocol, the peptide-siRNA formulation is delivered intranasally with a pipette in the dominant hand, while the anesthetized mouse is restrained by the scruff with the nondominant hand in a "head down-and-forward position" to avoid drainage into the lung and stomach upon inhalation. This precise gripping of mice can be learned but is not easy and requires practice and skill to result in effective CNS uptake. Furthermore, the process is long-drawn, requiring about 45 min for the administration of a total volume of ~20-30 &#181;L of solution in 1-2 &#181;L droplet volumes per inhalation, with 3-4 min rest periods between each inhalation. The objective of this study is to disclose a mouse positioning device that enables the appropriate placement of mice for efficient IN administration of the peptide-siRNA formulation. Multiple features are incorporated into the design of the device, such as four or eight positioning chairs with adjustable height and tilt to restrain anesthetized mice in the head down-and-forward position, enabling easy visualization of the mice's nares and a built-in heating pad to maintain the mice's body temperatures during the procedure. Importantly, the ability to treat four or eight mice simultaneously with RVG9R-siRNA complexes in this manner enables studies on a much quicker time scale, for the testing of an IN therapeutic siRNA approach. In conclusion, this device allows for appropriate and controlled mouse head positioning for IN application of RVG9R-siRNA and other therapeutic molecules, such as nanoparticles or antibodies, for CNS delivery.

Here, we present a protocol using a mouse positioning device that enables the appropriate placement of mice for the intranasal administration of a brain-targeting peptide-siRNA formulation enabling effective gene silencing in the central nervous system.

Intranasal (IN) drug delivery to the brain has emerged as a promising method to bypass the blood-brain barrier (BBB) for the delivery of drugs into the ...

Immunology and Infection

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, ...

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 ...

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 ...

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

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.

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.

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 ...

Lumbar Intrathecal Injection of Gene Therapy Vectors for Central Nervous System Targeting in Mice and Rats

One method to target the central nervous system for treating neurological diseases involves utilizing the lumbar intrathecal route of administration. This approach bypasses the blood-brain barrier to directly access the cerebrospinal fluid and preferentially target cells within the central nervous system. Multiple published preclinical studies employing the lumbar intrathecal injection route have contributed to the development of gene therapy clinical trials; however, the described protocols are variable and dispersed across multiple resources. Here, a comprehensive set of protocols for lumbar intrathecal injections in neonatal, juvenile, and adult mice and rats for preclinical gene therapy studies is presented. With proper training, this injection technique can be performed quickly and reliably. In addition to detailing the injection protocol at each developmental stage, associated parameters, such as injection volume, that can influence study outcomes are discussed. To demonstrate the application of lumbar intrathecal injections for targeting the central nervous system, the expression of adeno-associated virus serotype 9 in the brain, spinal cord, and peripheral tissues is presented following a successful or unsuccessful injection.

A lumbar intrathecal injection represents a translationally relevant route of administration for delivering gene therapy to the central nervous system. This comprehensive standardized protocol for lumbar intrathecal injections in neonatal, juvenile, and adult mice and rats aims to guide researchers in adopting this technique for preclinical gene therapy studies.

One method to target the central nervous system for treating neurological diseases involves utilizing the lumbar intrathecal route of administration. This ...

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

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Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Methods and Tips for Intravenous Administration of Adeno-associated Virus to Rats and Evaluation of Central Nervous System Transduction

A Positioning Device for the Placement of Mice During Intranasal siRNA Delivery to the Central Nervous System

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

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

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Lumbar Intrathecal Injection of Gene Therapy Vectors for Central Nervous System Targeting in Mice and Rats