Name: intrathecal injection of newborn mouse for genome editing and drug delivery
Uploaded: 2024-03-08T13:00:00.000+00:00
Duration: 4 min 33 s
Description: 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

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, simulating repeated administrations in humans in a clinical setting12.
Changes in intracranial pressure are commonly associated with intrathecal injection13, potentially leading to the dam's rejection and refusal to feed due to behavioral changes in the pups. However, acute changes in behavior or reduced survival among injected pups have not been observed. Similarly, unusual or abnormal behaviors associated with intrathecal injections in adults have not been noticed (data not shown).
Several technical tips may have contributed to the success and are worth emphasizing. The slower rate of injection is likely an important factor. Additionally, cryo-anesthesia may reduce intracranial pressure before the injection, minimizing backflow during the intrathecal injection and reducing other complications. Lastly, the precision of the injection site may also impact the success rate.
To achieve the best efficacy via intrathecal administration, performing the procedure as soon as possible after the pups are delivered is critical. Drugs and other substances delivered via intrathecal injection enter the intrathecal space, which is the space between the arachnoid mater and pia mater layers of the meninges that surround the brain and spinal cord. Therefore, the drugs delivered via intrathecal injection pass through these layers of the meninges12. In rodents, as in humans, the meninges consist of three layers: the dura mater, arachnoid mater, and pia mater14. These membranes form during embryonic development and are fully mature by postnatal day 2 (P2)15. Therefore, using a rapid genotyping protocol to assign the pups to experimental groups within a few hours is recommended, especially for experiments involving animal genotypes, such as gene-editing experiments. The earlier the pups are injected, the better the outcome. The injections are typically completed within 3 h after birth. This time window allows the injected drugs to follow the flow of cerebrospinal fluid into the brain parenchyma while the ependymal lining is still immature and less affected by the size of drug particles. It's worth noting the developmental differences between humans and mice. P1 neonatal mice correspond to the late gestational stage of human brain development16. Results from experiments on P1 neonatal mice serve as valuable proof of concept, but caution should be exercised when extrapolating these results to humans in translational study designs.
This technique is challenged by a constrained administration time window and the requirement for highly skilled experimenters. High mortality may be associated with the procedure if the experimenter lacks experience. However, the narrow timeframe demands a heightened level of precision and repeatability within and between studies. Furthermore, with adequate practice, the proficiency and success rate of this method can be significantly enhanced.
If the injections are performed correctly, the survival of injected pups is primarily affected by maternal care. Preparing foster female pairs for your targets is recommended. If the targeted pups do not have a milk spot on their bellies in the afternoon of the injection day, they should be transferred to the foster female immediately. Female mice recognize their babies by odor. Therefore, it is critical to avoid introducing unfamiliar odors from experimenters or unrelated dams to the pups during and after the procedure. However, the necessity of using foster females should be assessed for individual experiments. Performing the procedure in a well-ventilated room, ideally in a biological fume hood, is also recommended. Mixing the pups with the dam's bedding and excrement is also helpful. After the initial post-procedure check, minimizing disturbance to the dam for at least 3 days to reduce stress is recommended. Like any surgical procedure, the risk of post-procedure infection should be considered. Thus, strict adherence to good laboratory practices for sterile procedures should be followed during the injection. It should be noted that intrathecal pressure is higher than the external environment, providing natural protection against infection. Experience indicates that the rate of post-injection infection is rare. However, daily monitoring of the general appearance and activity of pups for at least 3 days post-injection to detect signs and symptoms of infection or other complications is recommended. In special cases, consultation with veterinary services may be warranted instead of euthanizing pups with significant complications.

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><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&amp;rdquo;/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-N). The long-term results (10 days after injection) were also examined using YFP reporter mice activated by the delivery of CRISPR/Cas9-based gene editing7. YFP expression was observed widely across the entire mouse brain compared to non-CRISPR/Cas9-treated mice (Figure 2). The expression of YFP was observed in the majority of cells under higher magnification. Injections were performed in more than 500 newborn pups, and over 98% of the injected pups survived the procedure. No harmful effects on the long-term survival and health of treated pups were observed (Supplementary Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65761/65761fig01.jpg" /> 
Figure 1: Temporal and spatial distribution of Fast Green dye in mouse brains. (A) Gross observation of mice, comparing injected and non-injected mice 5 min after intrathecal injection. (B) Visualization of the distribution of Fast Green dye in the mouse brain before dissection. (C-E) Distribution of the dye in dissected brains 5 min after intrathecal administration. (F-H) Distribution of the dye in dissected brains 30 min after intrathecal administration. (I-K) Distribution of the dye in dissected brains 60 min after intrathecal administration. (L-N) For comparison, the distribution of the dye in dissected brains 40 min after intracerebroventricular administration. Scale bar: 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65761/65761fig01large.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/65761/65761fig02.jpg" /> 
Figure 2: Long-term effects of intrathecally administered CRISPR/Cas9 gene editing. Widespread expression of the YFP reporter in the mouse brain following intrathecal injection of CRISPR/Cas9 gene editing: cerebellum (A-F), posterior cortex (D-F), and prefrontal cortex (G-I). Scale bar: 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65761/65761fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Survival curve of Angelman Syndrome mice after intrathecal CRISPR gene editing. Survival curve depicting the outcomes of Angelman Syndrome mice receiving CRISPR gene editing through intrathecal administration, compared to non-treated and wild-type mice. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65761/Supplementary Figure 1.zip" target="_blank">Please click here to download this File.</a>
Supplementary Video 1: Intrathecal injection procedure in neonatal mice. Video demonstrating the process of delivering intrathecal injections to neonatal mice. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65761/Supplementary Video 1.zip" target="_blank">Please click here to download this Video.</a>

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

3.6K Views.  Yale University School of Medicine. Onze groep hanteert een interdisciplinaire benadering met genomische ontdekking en functionele modellering om genetica en epigenetische basis van neurologische ontwikkelingsstoornissen te bestuderen. We voeren moleculaire, neurofysiologische en neurologische gedragsanalyses uit met behulp van door de patiënt geïnduceerde pluripotente stamcel, of iPSC, in de gemuteerde muizen om de pathofysiologie van neurologische ontwikkelingsstoornissen te ontleden. We streven ernaar om een op genbewerkin...

Video: Auteur in de schijnwerpers: Intrathecale injectie - een efficiënte en betrouwbare toedieningsmethode om de werkzaamheid van genbewerking in neonatale muizenhersenen te testen

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

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 ...

Transposon Mediated Integration of Plasmid DNA into the Subventricular Zone of Neonatal Mice to Generate Novel Models of Glioblastoma

An urgent need exists to test the contribution of new genes to the pathogenesis and progression of human glioblastomas (GBM), the most common primary brain tumor in adults with dismal prognosis. New potential therapies are rapidly emerging from the bench and require systematic testing in experimental models which closely reproduce the salient features of the human disease. Herein we describe in detail a method to induce new models of GBM with transposon-mediated integration of plasmid DNA into cells of the subventricular zone of neonatal mice. We present a simple way to clone new transposons amenable for genomic integration using the Sleeping Beauty transposon system and illustrate how to monitor plasmid uptake and disease progression using bioluminescence, histology and immuno-histochemistry. We also describe a method to create new primary GBM cell lines. Ideally, this report will allow further dissemination of the Sleeping Beauty transposon system among brain tumor researchers, leading to an in depth understanding of GBM pathogenesis and progression and to the timely design and testing of effective therapies for patients.

Here we describe an efficient and versatile protocol to induce, monitor and analyze novel glioblastomas (GBM) using transposon DNA injected into the ventricles of neonatal mice. Cells of the subventricular zone, which take up the plasmid, transform, proliferate and generate tumors with histo-pathological characteristics of human GBM.

An urgent need exists to test the contribution of new genes to the pathogenesis and progression of human glioblastomas (GBM), the most common primary ...

Murine Neural Plate Targeting by In Utero Nano-Injection (NEPTUNE) at Embryonic Day 7.5

Manipulating gene expression in the developing mouse brain in utero holds great potential for functional genetics studies. However, it has previously largely been restricted to the manipulation of embryonic stages post-neurulation. A protocol was developed to inject the amniotic cavity at embryonic day (E)7.5 and deliver lentivirus, encoding cDNA or shRNA, targeting &gt;95% of the neural plate and neural crest cells, contributing to the future brain, spinal cord, and peripheral nervous system. This protocol describes the steps necessary to achieve successful transduction, including grinding of the glass capillary needles, pregnancy verification, developmental staging using ultrasound imaging, and optimal injection volumes matched to embryonic stages. 
Following this protocol, it is possible to achieve transduction of &gt;95% of the developing brain with high-titer lentivirus and thus perform whole-brain genetic manipulation. In contrast, it is possible to achieve mosaic transduction using lower viral titers, allowing for genetic screening or lineage tracing. Injection at E7.5 also targets ectoderm and neural crest contributing to distinct compartments of the eye, tongue, and peripheral nervous system. This technique thus offers the possibility to manipulate gene expression in mouse neural-plate- and ectoderm-derived tissues from preneurulation stages, with the benefit of reducing the number of mice used in experiments.

Murine Neural Plate Targeting by In Utero Nano-Injection NEPTUNE at Embryonic Day 7.5

In this protocol, we describe how to inject the mouse amniotic cavity at E7.5 with lentivirus, leading to uniform transduction of the entire neural plate, with minimal detrimental effects on survival or embryonic development.

Manipulating gene expression in the developing mouse brain in utero holds great potential for functional genetics studies. However, it has previously ...

Developmental Biology

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

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 ...

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

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 ...

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 ...

Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery

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