This work was funded by a grant from HEBEI Provincial Department of Human Resources and Social Security (CY201605) and a grant from Natural Science Foundation of Hebei Province (H2017206101), and we are very grateful to Dr. for Guangping Gao, who provided the AAV for this study.

FVB/NJ mice were bred in the animal facility of Key Laboratory of Hebei Neurology. All mouse experiments were approved by the Second Hospital of Hebei Medical University Ethics Committee and carried out according to the regulations of laboratory animal management promulgated by the Ministry of Science and Technology of the People&#39;s Republic of China.
1. Preparation of 20% Lidocaine Hydrochloride Stock Solution
<ol>
	<li>Weigh 2 g of lidocaine hydrochloride. Add 5&#8211;6 mL of sterile water and vortex gently. Increase the volume to a total of 10 mL with sterile water.</li>
	<li>1.2 Filter the stock solution through 0.2-micron filters. Aliquot the stock solution, 1 mL per 1.5 mL microcentrifuge tube,and seal it with a sealing film. Store at 4 &#176;C.</li>
</ol>
2. Direct Intrathecal AAV Delivery in Awake Mice
<ol>
	<li>Wipe the work area using sterile gauze with 70% ethanol and prepare the required supplies as mentioned in Table 1.</li>
	<li>Prepare 100 &#181;L of AAVrh.10/1% lidocaine hydrochloride complex by adding 95 &#181;L of rAAVrh10 stock solution (5 x 1012 genome copies/mL) into a sterile 200 &#181;L microcentrifuge tube, and add 5 &#181;L of 20% lidocaine hydrochloride stock solution. Mix well by pipetting up and down. Then, store the virus solution on ice (4 &#176;C). 
	NOTE: 8 &#181;L per mouse4 will be used.</li>
	<li>Preparing the syringe with AAV solution for injection
	<ol>
		<li>Assemble a 25 &#181;L Hamilton syringe with a 27 G needle and align the beveled tip of the needle with the volumetric scale on the syringe.</li>
		<li>Draw 8 &#181;L with 4 x 1010 genome copies of the virus solution into the syringe gently.Make sure to remove air bubbles.</li>
	</ol>
	</li>
	<li>Preparing the mouse 
	NOTE: Male or female FVB/NJ mice (30&#8211;70 days old) were used in this study. IT injection was operated in the hood.
	<ol>
		<li>Sterilize the work area in a hood with 70% ethanol. Put the awake mouse (male or female, 30&#8211;70 days old, 13&#8211;20 g weight) on a bedpiece in a prone position in the hood. Cover the upper body with sterile gauze to calm the mouse and avoid being bitten.</li>
		<li>Fix the animal by gripping appropriately and firmly on its pelvic girdle with a thumb on one side and forefinger/middle finger on the other side. Keep the skin between bilateral pelvic girdles taut with the thumb and forefinger. Hold gently on the upper body of the animal with the palm.</li>
		<li>Shave the fur on its back between the bilateral pelvic girdles, then sterilize the skin surface with an iodide-based scrub and 70% ethanol.</li>
	</ol>
	</li>
	<li>Intrathecal Injection12
	<ol>
		<li>Feel the intervertebral space along the middle line between the bilateral pelvic girdles with a thumb or forefinger of the other hand and press an indentation with a fingernail to indicate the L5-L6 intervertebral space (locate the injection site).</li>
		<li>Rotate the base of the tail slightly and gently to indicate the midline of the spine. Adjust the needle bevel towards the head of the animal before injection (mentioned in step 2.3.1).</li>
		<li>Make sure that the animals are fixed firmly and align the needle along the midline of spine.</li>
		<li>Insert the needle gently and vertically (or tilt slightly 70&#8211;80&#176;) in the intersection of indentation and keep the syringe in a central sagittal plane. Reduce the angle to approximately 30&#176; slowly when it connects the bone, then slip the needle into the intervertebral space. 
		NOTE: An evident sudden tail flick is a sign of successful entry into the intradural space. Once the needle enters the intervertebral space, the needle tip will feel firmly clamped. The 27 G needle used in this study is suited for IT delivery in mice but not rats.</li>
		<li>Inject the vector solution(mentioned in step 2.2). Start the timer and inject 8 &#181;L of the vector solution at a speed of 1 &#181;L/4 s. Retain the needle approximately 1 min after finishing delivery. Withdraw the needle with gentle rotation to avoid leaking.</li>
		<li>Score the transient weakness of the mouse limbs immediately after delivery to evaluate the injection quality4. 
		NOTE: The standard is as follows4. Score 0: no weakness; score 1: minor weakness of the hind limbs without gait abnormality; score 2: moderate weakness of the hind limbs with obvious gait abnormality; score 3: complete paralysis of the hind limbs; score 4: complete paralysis of the hind limbs, shortness of breath, and moderate weakness of the fore limbs; and score 5: complete paralysis of all four limbs and evident shortness of breath.</li>
		<li>Move the mouse back to the cage for recovery from paralysis.</li>
	</ol>
	</li>
	<li>Clean-up
	<ol>
		<li>Flush the syringe with 1 mL sterile water. Sort laboratory supplies and collect all non-disposable materials for autoclaved sterilization. Clean the bench with 70% ethanol.</li>
	</ol>
	</li>
</ol>
3. Tissue Preparation for Immunohistochemical Staining
<ol>
	<li>Tissue collection
	<ol>
		<li>Anesthetize mice at 21 days post-injection with 3% chloral hydrate (0.1 mL/10 g) deeply by intraperitoneal injection.</li>
		<li>Perfuse transcardially with 20 mL of ice-cold 0.01 M PBS (NaCl 147 mM; NaH2PO4 1.9 mM; K2HPO4 8.1 mM, pH 7.4) firstly, then 4% ice-cold paraformaldehyde (in 0.01 M PBS) with pump (10 mL/min for 1 min, then 5 mL/min for 9 min). 
		CAUTION: Paraformaldehyde is carcinogenic and toxic. Handle it only in the fume hood while wearing gloves.</li>
	</ol>
	</li>
	<li>Dissection of the spinal cord and brain
	<ol>
		<li>Fix the limbs and head of each animal in a prone position on a foam box cover with syringe needles, then strip and remove the skin from the head to sacrum with scissors.</li>
		<li>Clip the skull between eyes, cut alongside the middle route of the skull and horizontal line upon the cerebellum, then open the skull to each side.</li>
		<li>Lift the occipital bone with tweezers and open the spinal canal bilaterally with ophthalmic scissors. Cut off the ribs on both sides and remove the upper half of vertebrae carefully.</li>
		<li>Lift the brain with curved tweezers and sever the nerves of the skull base, then dissect out the whole brain and spinal cord carefully. Post-fix the tissues in 4% paraformaldehyde for 24 h.</li>
	</ol>
	</li>
	<li>Preparation of tissue slices
	<ol>
		<li>Cryoprotect the brain and cervical and lumbar spinal cord in 30% sucrose solution overnight at 4 &#176;C. Embed the tissue in optimum cutting temperature (OCT) compound and freeze fast with liquid nitrogen.</li>
		<li>Cut the tissue at 25 &#181;m using a cryostat and store the frozen sections in 0.01 M PBS at 4 &#176;C for use .</li>
	</ol>
	</li>
</ol>
4. Immunohistochemistry
<ol>
	<li>Pretreat free-floating sections in 1% H2O2 for 10 min, then wash in PBS for 10 min. Incubate in a blocking solution containing 5% serum and 0.3% non-ionic detergent in PBS for 1 h.</li>
	<li>Incubate the slices with corresponding primary antibodies overnight at 4 &#176;C. Wash the sections in PBST (0.2% Tween 20 in PBS) for 30 min (3 times for 10 min each).</li>
	<li>Incubate the slices with corresponding biotin-secondary antibody at room temperature for 1 h. Wash in PBST for 30 min (3 times for 10 min each).</li>
	<li>Incubate the sections with affinity biotin peroxidase complex for 40 min, and stain with achromogenic agent. Mount the sections onto slides and dry properly.</li>
	<li>Soak the slides in anhydrous ethanol for 5 min and xylene for 10 min, then seal the slides with a mounting medium. Finally, image the slides with a microscope equipped with a charge-coupled device (CCD) at 100x, 200x, and 400x magnifications.</li>
</ol>

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The authors have nothing to disclose.

Technically, there are several critical steps during the IT injection in awake mice. First, proper gesture and firm control of the mice throughout the entire operation is a prerequisite for successful delivery. Second, the most difficult point is feeling the intervertebral space with the needle tip, as it is necessary not to insert too deeply without resistance or insert forcibly under strong resistance in the case of injuring the animals or bending the needle tip. Third, although the transient paralysis due to lidocaine...

AAV can mediate long-term and widespread gene expression in the CNS transduction with few side effects, and therefore has become one of the most promising vehicles for gene therapy to treat CNS diseases including amyotrophic lateral sclerosis (ALS), Huntington&#39;s disease (HD), Alzheimer&#39;s disease (AD), lysosomal storage diseases (LSD), Gaucher disease (GD), and neuronal ceroid lipofuscinosis (NCL)1. Presently, more than 100 AAV serotypes have been isolated from humans and animals. Among these, at least 12 have been used in preclinical and clinical trials, including the most commonly used gene vectors such as AAV1, 2, 4, 5, 6, 8, 9, rAAVrh.8, and rAAVrh.101,2,3,4,5,6.
Different CNS diseases require different AAV delivery strategies due to the various affected CNS regions and cell types. The CNS regions and cell types that AAV can transduce varies depending on the serotype as well as delivery method. For example, rAAVrh10 has been shown to transduce predominantly astrocytes when delivered by systemic intravenous injection (IV), whereas it transduced both neurons and glia when delivered by intrathecal injection4,7. Additionally, parenchyma injection resulted in local transduction to the vicinity of the injection site, whereas injection into the cerebrospinal fluid (CSF) through intraventricular or intrathecal injection resulted in widespread CNS transduction8. Studies have also demonstrated therapeutic potency of AAV-delivered gene therapy in neurodegenerative disorders by IT administration9,10,11. In diseases that affect broad areas of the CNS such as ALS, intrathecal injection into the CSF has been shown to cover most areas that are afflicted by the disease with a lower dose, compared to a systemic delivery method4,10. Recent studies have also shown that lumbar puncture can be used to inject AAV in mouse models for ALS, which avoids potential injuries associated with laminectomy and intrathecal catheterization4.
Experimental direct lumbar puncture was first used to deliver agents, especially anesthetics, to the spinal cord for analgesia and anesthesia in 188512,13. In this report, we illustrate the lumbar puncture IT injection method in adult mice with the aid of 1% lidocaine hydrochloride, a local amide-derived anesthetic, in the injection solution to evaluate and monitor injection quality. Successful injections were marked by lidocaine-induced transient paralysis, whereas failed injections did not show this behavior. We classified the level of transient weakness as one of five grades to help predict the injection efficiency. Finally, we show that the rAAVrh10 transduction level may be predicted by the grade of paralysis. Therefore, this intrathecal AAV delivery method can be used to enhance AAV-mediated gene-delivery for experimental therapy of CNS diseases.

<table><tbody><tr><td>FVB/NJ mice</td><td>Charles River Laboratories China</td><td></td><td></td></tr><tr><td>Lidocaine hydrochloride monohydrate</td><td>HEOWNS</td><td>73-78-9</td><td></td></tr><tr><td>AAV</td><td>Viral Vector Core of the Gene Therapy Center at University of Massachusetts Medical School</td><td></td><td></td></tr><tr><td>25 &amp;micro;L&amp;nbsp; Hamilton syringe/27-30 G needle</td><td>GASTIGHT</td><td>1702</td><td></td></tr><tr><td>O.C.T compond</td><td>SAKURA</td><td>4583</td><td></td></tr><tr><td>H&lt;sub&gt; 2&lt;/sub&gt;O&lt;sub&gt; 2&lt;/sub&gt;</td><td>SHUI HUAN PAI</td><td>170401</td><td></td></tr><tr><td>Goat serum</td><td>Solarbio</td><td>S9070</td><td></td></tr><tr><td>Triton X-100</td><td>LIFE SCIENCES</td><td>T8200</td><td></td></tr><tr><td>Rabbit anti-GFP</td><td>Life tech</td><td>G10362</td><td>1:333 dilution</td></tr><tr><td>The second antibody (goat-anti rabbit)</td><td>Jackson Immuno Research</td><td>111-005-144</td><td>1:1,000 dilution</td></tr><tr><td>VECTASTAIN ABC REAGENT</td><td>Vector Lab</td><td>PK-6100</td><td></td></tr><tr><td>ImmPACT DAB Peroxidase Substrate Kit</td><td>Vector Lab</td><td>SK-4105</td><td></td></tr><tr><td>Mounting medium for fluorescence with DAPI</td><td>Vectorshield</td><td>H-1200</td><td></td></tr><tr><td>NaCl</td><td>Yong Da Chemical</td><td></td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Yong Da Chemical</td><td></td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;12H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Yong Da Chemical</td><td></td><td></td></tr><tr><td>Paraformaldehyde</td><td>Yong Da Chemical</td><td>307699</td><td></td></tr><tr><td>Adhesion Microscope Slides</td><td>CITOGLAS</td><td>17083</td><td>25 mm x 75 mm</td></tr><tr><td>SUPER-SLIP MICRO-GLAS</td><td>Electro Microscopy Siences</td><td>72236-60</td><td>24 mm x 60 mm</td></tr><tr><td>15 mL Centrifuge tube</td><td>CORNING</td><td>430790</td><td></td></tr><tr><td>96 well cell culture cluster</td><td>Coster</td><td>3599</td><td></td></tr><tr><td>24 well cell culture cluster</td><td>Coster</td><td>3524</td><td></td></tr><tr><td>70% Ethanol</td><td>WEN ZHI</td><td></td><td></td></tr><tr><td>Gauze</td><td>Wei AN</td><td>05171112</td><td>8 cm x 10 cm x 12 cm</td></tr><tr><td>1 mL syringe</td><td>Hong Da</td><td></td><td></td></tr><tr><td>Microtubes</td><td>Plasmed</td><td></td><td></td></tr><tr><td>Micropipet&amp;nbsp;</td><td>eppendorf</td><td></td><td></td></tr><tr><td>Peppet tips</td><td>Rainin</td><td></td><td></td></tr><tr><td>Centirifuge</td><td>eppendorf</td><td>5427R</td><td></td></tr><tr><td>Regerator</td><td>Haier</td><td>BCD-539WT</td><td></td></tr><tr><td>Filter</td><td>MILLEX GP</td><td>R4PA42342</td><td></td></tr><tr><td>Pump</td><td>LongerPump</td><td>BT-100-2J/YZ1515X</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td>BX53</td><td></td></tr><tr><td>Freezing-microtome</td><td>Leica</td><td>CM1520</td><td></td></tr></tbody></table>

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.

Mice showed different degrees of transient weakness right after IT injection of AAV solution in 1% lidocaine hydrochloride due to various quality of intrathecal injection. According to the semi-quantitative 5-grade scoring system we have established, we tested the transduction patterns of AAV in mice with different degrees of lidocaine-induced limb weakness (score 0, n = 2; score 1, n = 1; score 4, n = 4; score 5, n = 3). EGFP immunostaining of spinal cords showed either no or little tran...

Watch this Scientific Journal Video about Direct Intrathecal Injection of Recombinant Adeno-associated Viruses in Adult Mice at JoVE.com

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

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

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

direct-intrathecal-injection-recombinant-adeno-associated-viruses

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36.6K Views.  Second Hospital of Hebei Medical University. 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.

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

Intracranial Injection of Adeno-associated Viral Vectors

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific subsets of cells in different brain regions both in vivo and in brain sections. Unlike the injection of fluorescent dyes, viral labeling offers targeting of individual cell types and is less expensive and time consuming than establishing transgenic mouse lines. In this technique, an adeno-associated viral (AAV) vector is injected intracranially using stereotaxic coordinates, a micropipette and an automated pump for precise delivery of AAV to the desired area with minimal damage to the surrounding tissue. Injection parameters can be tailored to individual experiments by adjusting the animal age at injection, injection location, volume of injection, rate of injection, AAV serotype and the promoter driving gene expression. Depending on the conditions chosen, virally-induced transgene expression can allow visualization of groups of cells, individual cells or fine cellular processes, down to the level of dendritic spines. The experiment shown here depicts an injection of double-stranded AAV expressing green fluorescent protein for the labeling of neurons and glia in the mouse primary visual cortex.

Here we present the intracranial injection of AAV vectors for fluorescent labeling of neurons and glia in the visual cortex.

Intracranial injection of viral vectors engineered to express a fluorescent protein is a versatile labeling technique for visualization of specific ...

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 ...

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 ...

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

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

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

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

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

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), ...

Adeno-associated Virus-mediated Transgene Expression in Genetically Defined Neurons of the Spinal Cord

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or triple transgenic mice are generated in order to achieve selective expression of a reporter or effector gene (e.g., from the Rosa26 locus) in the desired spinal population. 2) Intraspinal injection of Cre-dependent recombinant adeno-associated virus (rAAV); here Cre-dependent AAV vectors coding for the reporter or effector gene of choice are injected into the spinal cord of mice expressing Cre recombinase in the desired neuronal subpopulation. This protocol describes how to generate Cre-dependent rAAV vectors and how to transduce neurons in the dorsal horn of the lumbar spinal cord segments L3-L5 with rAAVs. As the lumbar spinal segments L3-L5 are innervated by those peripheral sensory neurons that transmit sensory information from the hindlimbs, spontaneous behavior and responses to sensory tests applied to the hindlimb ipsilateral to the injection side can be analyzed in order to interrogate the function of the manipulated neurons in sensory processing. We provide examples of how this technique can be used to analyze genetically defined subsets of spinal neurons. The main advantages of virus-mediated transgene expression in Cre transgenic mice compared to classical reporter mouse-induced transgene expression are the following: 1) Different Cre-dependent rAAVs encoding various reporter or effector proteins can be injected into a single Cre transgenic line, thus overcoming the need to create several multiple transgenic mouse lines. 2) Intraspinal injection limits manipulation of Cre-expressing cells to the injection site and to the time after injection. The main disadvantages are: 1) Reporter gene expression from rAAVs is more variable. 2) Surgery is required to transduce the spinal neurons of interest. Which of the two methods is more appropriate depends on the neuron population and research question to be addressed.

Intraspinal injection of recombinase dependent recombinant adeno-associated virus (rAAV) can be used to manipulate any genetically labelled cell type in the spinal cord. Here we describe how to transduce neurons in the dorsal horn of the lumbar spinal cord. This technique enables functional interrogation of the manipulated neuron subtype.

Selective manipulation of spinal neuronal subpopulations has mainly been achieved by two different methods: 1) Intersectional genetics, whereby double or ...

Purification of AAV Vectors by Single-Step Heparin Affinity Chromatography

Isolation of Adeno-Associated Viral Vectors Through a Single-Step and Semi-Automated Heparin Affinity Chromatography Protocol

Adeno-associated virus (AAV) has become an increasingly valuable vector for in vivo gene delivery and is currently undergoing human clinical trials. However, the commonly used methods to purify AAVs make use of cesium chloride or iodixanol density gradient ultracentrifugation. Despite their advantages, these methods are time-consuming, have limited scalability, and often result in vectors with low purity. To overcome these constraints, researchers are turning their attention to chromatography techniques. Here, we present an optimized heparin-based affinity chromatography protocol that serves as a universal capture step for the purification of AAVs.
This method relies on the intrinsic affinity of AAV serotype 2 (AAV2) for heparan sulfate proteoglycans. Specifically, the protocol entails the co-transfection of plasmids encoding the desired AAV capsid proteins with those of AAV2, yielding mosaic AAV vectors that combine the properties of both parental serotypes. Briefly, after the lysis of producer cells, a mixture containing AAV particles is directly purified following an optimized single-step heparin affinity chromatography protocol using a standard fast protein liquid chromatography (FPLC) system. Purified AAV particles are subsequently concentrated and subjected to comprehensive characterization in terms of purity and biological activity. This protocol offers a simplified and scalable approach that can be performed without the need for ultracentrifugation and gradients, yielding clean and high viral titers.

Author Spotlight: Improved Method for Production and Purification of Adeno-Associated Viral Vectors

This manuscript describes a detailed protocol for the generation and purification of adeno-associated viral vectors using an optimized heparin-based affinity chromatography method. It presents a simple, scalable, and cost-effective approach, eliminating the need for ultracentrifugation. The resulting vectors exhibit high purity and biological activity, proving their value in preclinical studies.

Adeno-associated virus (AAV) has become an increasingly valuable vector for in vivo gene delivery and is currently undergoing human clinical trials. ...

Bioengineering

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 ...