XNL is supported by Foundation for Angelman Syndrome Therapeutic (FAST) Postdoctoral Fellowship. YHJ is also support by FAST and NIH Grant R01HD110195 and R01MH117289.

The described procedures and protocols were in compliance with the guidelines outlined in the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Additionally, the procedures received approval from the Animal Care and Use Committee at Yale University School of Medicine. Newborn wild-type (WT) C57BL/6J male and female mice were used for presented study. The animals were obtained from a commercial source (see Table of Materials).
1. Preparation of the workspace
<ol>
	<li>Prepare the following items first: wet ice for cryo-anesthesia, an empty cage to separate the pups from the dam, a dissecting microscope, a light source, a clean surface for placing the animal during the injection, cotton swabs, a heating pad, a 25/10 &#181;L syringe, and a 34 G/ 0.375&#34;/ 12 DEG needle (see Table of Materials). 
	&#8203;NOTE: Cryo-anesthesia for mouse pups using wet ice is an optional step intended to facilitate handling, reduce pup movement, and minimize potential animal discomfort. This cryo-anesthesia step may also provide the benefit of lowering intracranial pressure and reducing volume-related complications9,10.</li>
	<li>Move the pups to a separate cage away from the dam while handling them.</li>
	<li>Weigh each pup and document their weight.</li>
	<li>Wipe the back of the mouse using gauze and ethanol. Confirm the intervertebral space or, at a minimum, the midline of the spinal canal (which should appear red in P1 pups) using the dissecting microscope (Supplementary Video 1).</li>
</ol>
2. Injection procedure
<ol>
	<li>To anesthetize a single pup, gently place it on a water-proof barrier such as a latex sleeve or aluminum foil on an ice bath for 3-5 min. It&#39;s important to avoid leaving the animal on the ice for an extended period, as doing so could pose potential risks of hypothermia-related complications, including ventricular fibrillation, tissue hypoxia, and metabolic acidosis. 
	NOTE: The duration of 3-5 min may vary on a case-by-case basis. Assess the signs of anesthesia, such as unresponsiveness to a toe pinch, to determine the appropriate duration.</li>
	<li>While the animal is on the ice, load the syringe with 10 &#181;L of the drug formulation, virus preparation, or control artificial spinal fluid, etc. 
	NOTE: During the learning phase, consider the option of injecting the same volume of mixed 1% Fast Green dye with the delivery materials (see Table of Materials). This can aid in visualizing the injection process and assist in learning and refining the technique. Pups injected with Fast Green Dye or similar materials should be euthanized shortly after injection per the approved protocol, as these materials may result in inflammatory reactions or other side effects in animals.</li>
	<li>Once the animal is fully anesthetized, as confirmed by reduced or absent body movement, gently position the pups under the microscope.</li>
	<li>With the left index finger and thumb, carefully palpate the intervertebral space along the midline, situated between the bilateral pelvic girdles (Supplementary Video 1). Gently rotate the base of the tail slightly to help identify the midline of the spine.</li>
	<li>Adjust the needle bevel toward the head of the animal before injection.</li>
	<li>Carefully insert the needle, tilting it slightly to an angle of 70&#176;-80&#176; at the point where the indentation intersects, while ensuring the syringe remains aligned with the central sagittal plane. As the needle makes contact with the bone, gradually decrease the angle to approximately 30&#176;, then advance the needle about 2 mm into the intervertebral space. 
	NOTE: The ability of the needle to slightly lift the entire body is a sign of successful entry into the intradural space.</li>
	<li>Slowly inject up to 10 &#181;L volume within 50-60 s. Keep the needle in place for 10-20 s after delivery is completed. Withdraw the needle with a gentle rotation to avoid leaking. 
	&#8203;NOTE: The cerebellum will turn green before withdrawing the needle. Also, the slow push is critical to prevent an increase in intracranial pressure associated with delivery and to minimize potential complications. Based on our experience with injections in more than 500 pups, delivering a 10 &#181;L volume over 50-60 s is optimal.</li>
</ol>
3. Post injection
<ol>
	<li>Apply a cotton swab to the injection site if there is any leaking or blood. 
	NOTE: There should not be any in most cases. From our experience, pups treated with traces of leaking or blood are still usable, but consideration of a reduced dose of drugs or treatments may be necessary during data analysis.</li>
	<li>Place the pup on a heating pad and allow 10-15 min for the pups to fully recover and re-warm. Carefully observe the pups to ensure they are alert and actively moving before returning them to their home cage. Adequate recovery of a mouse is indicated by the restoration of pink skin color, increased spontaneous body movement, and responsive reactions to touch.</li>
	<li>Place the pup back into the home cage and ensure that the pup is properly covered with bedding, nestlet, or both. This ensures that the pup receives the necessary maternal care from the dam.</li>
	<li>Assess the general appearance and activity daily for at least 3 days post-injection. A sick appearance and reduced activity may raise the possibility of infection, treatment-associated side effects, or other complications, etc. If necessary, consult with veterinary care.</li>
</ol>

A Comprehensive Protocol for Delivering Intrathecal Injections in Neonatal Mice

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YHJ is a co-founder of Couragene but no conflict of interest for this project.

Described is a step-by-step procedure for intrathecal injection in neonatal mice (P1), resulting in widespread drug distribution in their brains. In comparison to the common intracerebroventricular injection method for delivering medication to neonatal mice, which involves piercing the cerebral cortex11, intrathecal injection avoids direct injury to the neonatal mouse brain due to needle penetration. Due to minimal invasiveness, intrathecal injection can be performed repeatedly when necessary, sim...

Intrathecal (IT) injection is a common clinical procedure used to administer medications, collect cerebrospinal fluid, and maintain intracranial pressure in both pediatric and adult patients in clinics1,2. The administration of medications via intrathecal injection is an effective approach for increasing medication concentrations in the central nervous system (CNS) while minimizing systemic exposure. Consequently, this method enhances therapeutic efficacy and reduces side effects, especially for temperature-sensitive and short half-life drugs3.
In preclinical studies testing new drugs and treatments using rodent models, it is imperative to employ a reliable method of drug administration that offers greater precision and result reproducibility4,5. For preclinical studies evaluating new treatments for neurogenetic and neurodevelopmental disorders, early treatment is crucial for initial proof-of-concept studies because earlier interventions are typically predicted to yield more favorable outcomes6,7,8.
Compared to conventional intracerebroventricular (ICV) injections, IT injections carry significantly lower risks since they obviate the need for direct penetration through the cerebral cortex. This advantage substantially reduces the potential damage to regional cortical tissue and surrounding nerves. Furthermore, IT injections allow for at least a fivefold increase in the administrable volume of medications through a single injection, greatly enhancing the feasibility of repeated administrations. However, due to the small size and fragile nature of newborn mice, performing intrathecal injections in newborn pups is technically challenging and requires specialized techniques, equipment, and meticulous handling.
This article provides a detailed protocol with step-by-step instructions for performing intrathecal injections in P1 newborn pups. The key considerations and good laboratory practices are emphasized here to ensure the consistency of administration and the safety and well-being of the animals during the procedure. By following this protocol, researchers can confidently conduct experiments with precision and reproducibility while minimizing any potential risks or discomfort to the animals.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Balance</td>
			<td>Ohaus Corporation</td>
			<td>30253017</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J mice</td>
			<td>The Jackson Laboratory</td>
			<td>000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital Microscope</td>
			<td>RWD</td>
			<td>DOM-1001</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>ThermoFisher</td>
			<td>14190144</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast Green</td>
			<td>Sigma</td>
			<td>F7252-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>RWD</td>
			<td>69020</td>
			<td></td>
		</tr>
		<tr>
			<td>Needles</td>
			<td>Hamilton</td>
			<td>6PK (34/0.375&#8221;/4/12DEG)S</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>Hamilton</td>
			<td>1702RN</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Filters</td>
			<td>Sigma</td>
			<td>SLGVM33RS</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Successful intrathecal injection immediately resulted in the widespread distribution of the administered solution, although the actual cellular penetration depended on the nature of the delivered drugs and materials. In this study, we used Fast Green to visualize the immediate results after intrathecal injection (IT) in wild-type neonates (Figure 1A-K) and compared it with conventional intracerebroventricular (ICV) injection (Figure 1L</...

Watch this Scientific Journal Video about Author Spotlight: Intrathecal Injection – An Efficient and Reliable Delivery Method to Test the Efficacy of Gene Editing in Neonatal Mouse Brains at JoVE.co

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 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 ...

intrathecal-injection-newborn-mouse-for-genome-editing-drug

Research

JoVE Journal

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3.4K Views.  Yale University School of Medicine. The present protocol outlines step-by-step instructions for performing intrathecal injections in neonatal mice for gene editing and drug delivery.

Video: Author Spotlight: Intrathecal Injection – An Efficient and Reliable Delivery Method to Test the Efficacy of Gene Editing in Neonatal Mouse Brains

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 ...

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals are not viable in those cases where the modified gene is expressed or deleted in the whole organism. Moreover, a variety of compensatory mechanisms often make it difficult to interpret the results. The compensatory effects can be alleviated by either timing the gene expression or limiting the amount of transfected cells.
The method of postnatal non-ventricular microinjection and in vivo electroporation allows targeted delivery of genes, siRNA or dye molecules directly to a small region of interest in the newborn rodent brain. In contrast to conventional ventricular injection technique, this method allows transfection of non-migratory cell types. Animals transfected by means of the method described here can be used, for example, for two-photon in vivo imaging or in electrophysiological experiments on acute brain slices.

This protocol describes a non-viral method of delivery of genetic constructs to a certain area of living rodent brain. The method consists of plasmid preparation, micropipette fabrication, neonatal rat pup surgery, microinjection of the construct, and in vivo electroporation.

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

The Corneal Micropocket Assay: A Model of Angiogenesis in the Mouse Eye

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets containing specific growth factors that trigger blood vessel growth throughout the naturally avascular cornea, angiogenesis can be measured and quantified. In this assay the angiogenic response is generated over the course of several days, depending on the type and dose of growth factor used. The induction of neovascularization is commonly triggered by either basic fibroblast growth factor (bFGF) or vascular endothelial growth factor (VEGF). By combining these growth factors with sucralfate and hydron (poly-HEMA (poly(2-hydroxyethyl methacrylate))) and casting the mixture into pellets, they can be surgically implanted in the mouse eye. These uniform pellets slowly-release the growth factors over five or six days (bFGF or VEGF respectively) enabling sufficient angiogenic response required for vessel area quantification using a slit lamp. This assay can be used for different applications, including the evaluation of angiogenic modulator drugs or treatments as well as comparison between different genetic backgrounds affecting angiogenesis. A skilled investigator after practicing this assay can implant a pellet in less than 5 min per eye.

The protocol describes the corneal micropocket assay as developed in mice.

The mouse corneal micropocket assay is a robust and quantitative in vivo assay for evaluating angiogenesis. By using standardized slow-release pellets ...

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene expression in the mouse brain. Here we describe a technique first introduced by Passini and Wolfe for direct intracranial delivery of virally-encoded transgenes into the neonatal mouse brain. In its most basic form, the procedure requires only an ice bucket and a microliter syringe. However, the protocol can also be adapted for use with stereotaxic frames to improve consistency for researchers new to the technique. The method relies on the ability of adeno-associated virus (AAV) to move freely from the cerebral ventricles into the brain parenchyma while the ependymal lining is still immature during the first 12-24 hr after birth. Intraventricular injection of AAV at this age results in widespread transduction of neurons throughout the brain. Expression begins within days of injection and persists for the lifetime of the animal. Viral titer can be adjusted to control the density of transduced neurons, while co-expression of a fluorescent protein provides a vital label of transduced cells. With the rising availability of viral core facilities to provide both off-the-shelf, pre-packaged reagents and custom viral preparation, this approach offers a timely method for manipulating gene expression in the mouse brain that is fast, easy, and far less expensive than traditional germline engineering.

Here we demonstrate a technique for widespread neuronal transduction by intraventricular injection of adeno-associated virus into the neonatal mouse brain. This method provides a rapid and easy way to attain lifelong expression of virally-delivered transgenes.

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene ...

Designing, Packaging, and Delivery of High Titer CRISPR Retro and Lentiviruses via Stereotaxic Injection

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique has been widely used to encode fluorescent proteins, opto- or chemo-genetic controllers of cell activity, or heterologous expression of human genes in model organisms. These viruses have also successfully been used to deliver recombinases to relevant target sites in transgenic animals, or even deliver small hairpin or micro RNAs in order to manipulate gene expression. While these techniques have been fruitful, they rely on transgenic animals (recombinases) or frequently lack high efficacy and specificity (shRNA/miRNA). In contrast, the CRISPR/Cas system uses an exogenous Cas nuclease which targets specific sites in an organism's genome via an exogenous guide RNA in order to induce double stranded breaks in DNA. These breaks are then repaired by non-homologous end joining (NHEJ), producing insertion and deletion (indel) mutations that can result in deleterious missense or nonsense mutations. This manuscript provides detailed methods for the design, production, injection, and validation of single lenti/retro virus particles that can stably transduce neurons to express a fluorescent reporter, Cas9, and sgRNAs to knockout genes in a model organism.

The CRISPR/Cas9 system offers the potential to make targeted genome editing accessible and affordable to the scientific community. This protocol is intended to demonstrate how to create viruses that will knockout a gene of interest using the CRISPR/Cas9 system, and then inject them stereotaxically into the adult mouse brain.

Replication defective lentiviruses or retroviruses are capable of stably integrating transgenes into the genome of an infected host cell. This technique ...

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

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, ...

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 ...

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

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Chapters in this video

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

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

The Corneal Micropocket Assay: A Model of Angiogenesis in the Mouse Eye

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

Designing, Packaging, and Delivery of High Titer CRISPR Retro and Lentiviruses via Stereotaxic Injection

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

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

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System