This research was supported by the Robert A. and Rene E. Belfer Family Foundation, NIA R21 AG038856 (JLJ), BrightFocus Foundation Alzheimer&#8217;s Disease research grant A2010097 (JLJ), and NIA Biology of Aging Training grant T32 AG000183 (support for SDG).

Perform all procedures and protocols involving animals in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. The procedures described here were reviewed and approved by the Baylor College of Medicine Institutional Animal Care and Use Committee.
Replication-incompetent adeno-associated virus (AAV) vectors for transgene delivery in the rodent brain are approved for Biosafety Level 1 use. Refer to the CDC website for the US Government publication &#8220;Biosafety in Microbiology and Biomedical Laboratories (BMBL)&#8221; which details specific requirements regarding proper protection and virus handling procedures. Check with local veterinary and environmental safety staff to learn institutional specific requirements for procedures using viruses. Regulations regarding procedure rooms, quarantine, and identification of biohazard cages vary across institutions.
1. Prepare P0 Pups and Foster Mothers
<ol>
	<li>Provide high fat chow for all pregnant and nursing females to support the energy demands of breeding and lactation.</li>
	<li>In the evening, set up a breeding cage by adding one or two healthy females to the cage of a single resident male. The following morning, check the females for a mating plug and separate the females from the males if a plug is present.
	<ol>
		<li>Use foster mothers to raise pups from a genetic background with poor maternal characteristics (for example, C57BL/6 or C3HeJ). Foster mothers accept pups from another litter if the transferred pups are born within four days of the foster mother&#8217;s own litter.</li>
		<li>Set up a separate mating cage using ICR or FVB females alongside the experimental breeders so that the two sets of females deliver at approximately the same time.</li>
	</ol>
	</li>
	<li>Females will deliver in 19 &#177;1 days from the plug date, depending on the background strain. Three days prior to delivery (16 days after the plug date), place the pregnant female into a clean cage with fresh nesting material and a covered shelter.</li>
	<li>Check for newborn pups twice daily starting 2 days prior to the expected delivery date (17 days after the plug date). Examine the cage with minimum stress to the mothers by peeking through the bottom for the presence of pink newborns. Mice generally deliver pups in the morning but on rare occasions will deliver in afternoon.</li>
	<li>Plan to perform injections as soon as the pups are nursing (which will be evident by visible milk spots), or within 6 hr of birth, whichever comes first.</li>
</ol>
2. Prepare Equipment for Injection
<ol>
	<li>Prepare a 10 &#181;l injection syringe with a 32 G&#160;needle for general P0 neonates. If performing stereotaxic injection, mount the syringe onto the manipulator arm.</li>
	<li>If using a stereotaxic frame for injections, cool the neonatal stage to 4-8 &#176;C by adding 100% ethanol and dry ice to the reservoir at the front end of the block. Maintain the temperature above 1 &#176;C to avoid frostbite of the pups.</li>
</ol>
3. Prepare Viral Dilutions for Injection
<ol>
	<li>Prepare a 1% trypan blue dye solution (20x stock).</li>
	<li>Prepare 5-25 &#181;l aliquots of adeno-associated virus (AAV) stock inside a Class 2 biosafety cabinet. Place these aliquots at -80 &#176;C for future use. Avoid additional freeze-thaw cycles as this causes the virus to lose transduction efficacy.</li>
	<li>Remove an aliquot of AAV from the -80 &#176;C freezer and place on ice to thaw.</li>
	<li>Prepare the viral solution for injection by diluting the virus into ice-cold 1x phosphate buffered saline (PBS). Start with a ten-fold serial dilution (1010 to 108 viral particles/hemisphere) for initial optimization of the transduction pattern.</li>
	<li>Add 20x trypan blue to the AAV solution to a final concentration of 0.05%.</li>
</ol>
4. Staging the Newborn Pups for Injection
<ol>
	<li>Place a small aluminum plate on ice to cool. Place a dry task wipe on top of the plate to protect the pup&#8217;s skin from the cold metal. This serves as a flat cold surface for anesthetizing and injecting the pups.</li>
	<li>Prepare a warming pad suitable for keeping neonatal mice warm before and after injection.</li>
	<li>Once the newborn pups have begun to nurse, remove approximately half of them from the cage and leave the other half with their mother. Place the collected pups on the warming pad while awaiting injection. 
	NOTE: If the biological mother does not care for the pups, remove the entire litter from the cage immediately and place the pups onto a warming pad to await injection. This situation may arise with the first litter born to C57BL/6 females or with other inbred strains that have poor breeding characteristics. In this situation, milk spots will not be visible on newborn pups.</li>
	<li>Transfer one pup from the warming pad onto the cold metal plate to induce hypothermia anesthesia. Wait 2-3 min for the pup to become fully anesthetized. Confirm anesthesia by very gently squeezing a paw and monitor for lack of movement or respiration.</li>
</ol>
5. Injection of AAV into Neonatal Mice
<ol>
	<li>Free-hand intracranial injection of AAV into neonatal mice:
	<ol>
		<li>Load 5 &#181;l of diluted AAV with 0.05% trypan blue into the injection syringe.</li>
		<li>Gently wipe the head of the anesthetized pup with a cotton swab soaked in 70% ethanol.</li>
		<li>Identify the injection sites at 2/5 of the distance from the lambda suture to each eye. An alternative injection site is located approximately 0.8-1 mm lateral from the sagittal suture, halfway between lambda and bregma. These landmarks are visible through the skin at P0.</li>
		<li>Mark the injection site with a non-toxic laboratory pen.</li>
		<li>Hold the syringe with the scale visible for monitoring the volume of solution dispensed.</li>
		<li>Ensure that the thumb can reach the top of the plunger. Then remove the thumb from the plunger while positioning the needle to avoid accidentally dispensing virus.</li>
		<li>Find a comfortable position to brace the arm for injection by putting the elbow on the bench and leaning the arm on the ice bucket.</li>
		<li>Lay the pup on its side with its head directly under the syringe. Turn the pup&#8217;s head so that the marked injection site is facing up, and gently but firmly hold this position with an open hand.</li>
		<li>Hold the syringe perpendicular to the surface of the skull and insert the needle at the marked injection site to a depth of approximately 3 mm. Inject the needle until the resistance decreases slightly, indicating that the needle has penetrated into the lateral ventricle. If a reference is needed, mark 3 mm from the tip of needle with non-toxic maker. Adjust the injection depth for pup size and strain based on dye targeting of the ventricles.</li>
		<li>Hold the syringe rigidly so that the plunger can be depressed without moving the needle farther into the brain. If the injection is displaced into the thalamus, viral spread will be severely limited.</li>
		<li>Begin slowly injecting virus while monitoring the volume dispensed from the syringe. Administer a maximum volume of 2 &#181;l into each ventricle. If the needle is in the correct position, the dye will spread to fill the ventricle.</li>
		<li>Slowly withdraw the needle.</li>
		<li>Allow the first injection site to close before injecting the other hemisphere. 
		NOTE: Do not inject the same site more than once. Reinserting the needle forces virus that has already been injected to leak out and causes injury or death of the pup.</li>
		<li>Inject the contralateral ventricle using the same procedure.</li>
	</ol>
	</li>
	<li>Stereotaxic injection of AAV into neonatal mice:
	<ol>
		<li>Load 5 &#181;l of diluted AAV with 0.05% trypan blue into the injection syringe held by the stereotaxic manipulator.</li>
		<li>Gently place the pup&#8217;s head between the ear bars of the neonatal frame. Make sure the head is level in the Y-axis (front to back) by checking that the line between lambda and bregma is parallel to the stage. Make sure the head is level in the X-axis (side to side) by checking that an imagined line between the ears, or a line between the eyes, is parallel to the stage.</li>
		<li>Gently wipe the anesthetized pup&#8217;s head with a cotton swab soaked in 70% ethanol.</li>
		<li>Use the stereotaxic manipulator to position the syringe above lambda and then zero the X and Y coordinates. Move the stereotaxic arms to (X, Y) = (0.8, 1.5) mm for standard P0 pups. 
		NOTE: Pup size and stereotaxic coordinates may vary by strain and age. Adjust coordinates based on dye injections as needed to target the lateral ventricles.</li>
		<li>Slowly lower the needle into the injection site. The surface of the skull will indent and then release once the needle has penetrated the skin. Retract the needle until the skull recovers its normal concave shape, but keep the bevel of the needle under the skin. Zero the Z coordinate at this point.</li>
		<li>Insert the needle until Z = -1.7 mm and then retract to -1.5 mm.</li>
		<li>Slowly inject the virus. Each hemisphere can accept up to 2 &#181;l of solution.</li>
		<li>Keep the syringe in place for 30-60 sec after completing the injection and then slowly retract the needle over 1-2 min.</li>
		<li>Repeat for the contralateral hemisphere, using negative coordinates in the X-axis for the injection site. 
		NOTE: Alternative coordinates for injection are (X, Y, Z) = (0.8, 2.0, -1.5) mm and (1.2, 1.0, -1.5) mm. Perform the first injection at (0.8, 2.0), then move to (0.8, -2.0), (1.2, 1.0) and finally (1.2, -1.0). Inject 1 &#956;l of solution at each site, for a total of 2 &#956;l per hemisphere. Move quickly between injections to complete the procedure in less than 10 min.</li>
	</ol>
	</li>
</ol>
6. Post-injection Care
<ol>
	<li>After completing injections into both hemispheres, place the pup back on the warming pad until its body temperature and skin color return to normal and the pup begins to move.</li>
	<li>If using the biological mother to nurse the injected neonates, return the injected pups to the biological mother after they recover normal movement. Place the remaining uninjected pups on the warming pad. Repeat the procedure until all mice have been injected.</li>
	<li>If using a foster mom to nurse the injected neonates, transfer all injected pups to the foster mother for care after they recover normal movement.
	<ol>
		<li>Remove most or all of the pups belonging to the foster mother from the cage to ensure the success of the injected animals.</li>
		<li>If foster mother&#8217;s pups and injected pups have the same eye and skin color, mark pups from the foster mother with a laboratory pen so they can be distinguished later.</li>
		<li>Place the foster mother&#8217;s pups and some of their bedding together with the injected pups. Ensure that the injected pups acquire the scent of her biological offspring to improve acceptance of the new pups.</li>
		<li>Place only the injected pups back into the foster mother&#8217;s cage. Cull any remaining pups from the foster mother.</li>
	</ol>
	</li>
	<li>Ensure that the mother is attending to and nursing the injected pups within 10 min&#160;of placing them into the cage. If the mother does not collect the pups within this time, transfer pups to another foster mother.</li>
	<li>Label the cage with a biohazard card to show that it contains virally-injected animals. House the cage in an approved area for 72 hr.</li>
	<li>After the 72 hr quarantine period, transfer the mother and pups (but no bedding or other cage items) to a clean cage. Perform this transfer inside a Level 2 biosafety cabinet to avoid exposure to any virally-contaminated bedding. Return the clean cage with the injected animals to the regular mouse facility.</li>
	<li>Place the dirty cage into a biohazard bag, sterilize by autoclaving, and then return the cage to the vivarium.</li>
</ol>
7. Cleanup
<ol>
	<li>Place the remaining virus solution at 4 &#176;C for future use. The virus retains transduction efficiency for at least 2 months when stored at 4 &#176;C8.</li>
	<li>Disinfect the injection needle and syringe with 2% bleach followed by repeated rinses in deionized water.</li>
	<li>Clean the working area with 2% bleach followed by 70% ethanol.</li>
	<li>Collect all disposable items including pipette tips, tubes, mask, and gloves in a plastic biohazard bag. Dispose of waste materials as required by local law and institutional policies (i.e. autoclave or incinerate).</li>
</ol>
8. Prepare Mouse Brains for Imaging
<ol>
	<li>Place an injected mouse into a carbon dioxide euthanasia chamber at 3-4 weeks post-injection. Follow institutional guidelines for proper euthanasia procedures. Do not perfuse animals with paraformaldehyde as this will quench the fluorescence label.</li>
	<li>Remove the whole brains from the skull and fix for 3-12 hr in 4% paraformaldehyde in PBS at 4 &#176;C.</li>
	<li>Transfer the fixed brain to 30% sucrose for cryoprotection. Incubate at 4 &#176;C until the brain sinks to the bottom.</li>
	<li>Collect a bucket of finely crushed dry ice. Cut the brain in half down the midline. Bury the brain halves in the dry ice to freeze.</li>
	<li>Cool the microtome stage by adding dry ice and 100% ethanol.</li>
	<li>Secure the brain to the stage with the midline down using PBS, and allow to freeze.</li>
	<li>Section through the brain at 30-45 &#181;m thickness using a freezing-sliding microtome.</li>
	<li>Place sections into 24-well plates containing antifreeze solution (250 ml glycerol, 300 ml ethylene glycol, 500 ml 0.1 M phosphate buffer, pH 7.4). Cover plates with tin foil and store at -20 &#176;C for future use.</li>
	<li>Collect sections from 1-4 wells and wash 3x in PBS.</li>
	<li>Mount sections onto microscope slides and coverslip. Allow slides to dry in a dark location.</li>
	<li>Image slides using appropriate filters for native YFP or tdTomato to check the virus transduction pattern.</li>
</ol>

<ol>
	<li>De Vry, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+electroporation+of+the+central+nervous+system:+a+non-viral+approach+for+targeted+gene+delivery.">In vivo electroporation of the central nervous system: a non-viral approach for targeted gene delivery.</a> Progress in neurobiology. 92, 227-244 (2010).</li><li>Sun, B., Gan, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manipulation+of+gene+expression+in+the+central+nervous+system+with+lentiviral+vectors.">Manipulation of gene expression in the central nervous system with lentiviral vectors.</a> Methods Mol Biol. 670, 155-168 (2011).</li><li>Puntel, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+transfer+into+rat+brain+using+adenoviral+vectors.">Gene transfer into rat brain using adenoviral vectors.</a> Curr Protoc Neurosci. 4, (2010).</li><li>Passini, M. A., Wolfe, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Widespread+gene+delivery+and+structure-specific+patterns+of+expression+in+the+brain+after+intraventricular+injections+of+neonatal+mice+with+an+adeno-associated+virus+vector.">Widespread gene delivery and structure-specific patterns of expression in the brain after intraventricular injections of neonatal mice with an adeno-associated virus vector.</a> J Virol. 75, 12382-12392 (2001).</li><li>Passini, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraventricular+brain+injection+of+adeno-associated+virus+type+1+(AAV1)+in+neonatal+mice+results+in+complementary+patterns+of+neuronal+transduction+to+AAV2+and+total+long-term+correction+of+storage+lesions+in+the+brains+of+beta-glucuronidase-deficient+mice.">Intraventricular brain injection of adeno-associated virus type 1 (AAV1) in neonatal mice results in complementary patterns of neuronal transduction to AAV2 and total long-term correction of storage lesions in the brains of beta-glucuronidase-deficient mice.</a> J Virol. 77, 7034-7040 (2003).</li><li>Kim, J. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Viral+transduction+of+the+neonatal+brain+delivers+controllable+genetic+mosaicism+for+visualising+and+manipulating+neuronal+circuits+in+vivo.">Viral transduction of the neonatal brain delivers controllable genetic mosaicism for visualising and manipulating neuronal circuits in vivo.</a> Eur J Neurosci. 37, 1203-1220 (2013).</li><li>Chakrabarty, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Capsid+serotype+and+timing+of+injection+determines+AAV+transduction+in+the+neonatal+mice+brain.">Capsid serotype and timing of injection determines AAV transduction in the neonatal mice brain.</a> PloS one. 8, (2013).</li><li>Croyle, M. A., Cheng, X., Wilson, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+formulations+that+enhance+physical+stability+of+viral+vectors+for+gene+therapy.">Development of formulations that enhance physical stability of viral vectors for gene therapy.</a> Gene therapy. 8, 1281-1290 (2001).</li><li>Bauer, H. C., Bauer, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+induction+of+the+blood-brain+barrier:+still+an+enigma.">Neural induction of the blood-brain barrier: still an enigma.</a> Cellular and molecular neurobiology. 20, 13-28 (2000).</li><li>Cearley, C. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expanded+repertoire+of+AAV+vector+serotypes+mediate+unique+patterns+of+transduction+in+mouse+brain.">Expanded repertoire of AAV vector serotypes mediate unique patterns of transduction in mouse brain.</a> Molecular therapy : the journal of the American Society of Gene Therapy. 16, 1710-1718 (2008).</li><li>Grimm, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+and+in+vivo+gene+therapy+vector+evolution+via+multispecies+interbreeding+and+retargeting+of+adeno-associated+viruses.">In vitro and in vivo gene therapy vector evolution via multispecies interbreeding and retargeting of adeno-associated viruses.</a> J Virol. 82, 5887-5911 (2008).</li></ol>

Authors declare that they have no competing financial interests.

We have described a versatile procedure for manipulating neuronal gene expression using AAV as a vehicle for widespread delivery into the neonatal mouse brain. Compared with other methods of neuronal transgenesis such as in utero electroporation1 or stereotaxic intracranial injection2,3, neonatal viral injection is relatively easy and simple. The basic procedure can be performed in minutes with only an ice bucket and a microliter syringe. Optimal survival and transgene expression can be attained by ...

Traditional methods for modifying neural gene expression require time-consuming and expensive germline manipulations. Alternative de novo approaches such as in utero electroporation or stereotaxic lentiviral injection yield faster results and are less costly but have the disadvantage of requiring complex surgical intervention1-3. Furthermore, transgene expression has a limited spatial range with these methods. Herein, we describe a fast, easy, and economical method for widespread neuronal manipulation via intraventricular injection of adeno-associated virus (AAV) into the neonatal mouse brain. The method was first described by John Wolfe and Marco Passini in 2001, where they suggested small particle size of AAV allowed it to diffuse within the cerebral spinal fluid as it passes from the lateral ventricles through the immature ependymal barrier and into the brain parenchyma4,5. Intraventricular injection of AAV within the first 24 hr after birth yields widespread viral transduction of neural subsets spanning every region of the brain, from the olfactory bulbs to the brain stem6,7. Virally-delivered transgenes are expressed and active within days of injection and persist for up to a year after transduction. Thus, this versatile manipulation enables studies ranging from early postnatal brain development to aging and degeneration in the adult.
In adapting the technique to our specific experimental needs, we have focused primarily on AAV8 serotype because it is the most efficient at transducing neurons6. We show that viral titer can be diluted to control the density of transduced neurons for experiments testing cell-intrinsic consequences of genetic manipulation. In addition, we demonstrate that two viruses could be co-injected to produce expression patterns that are biased towards distinct or overlapping sets of neurons, depending on the serotypes chosen for viral packaging. Our work expands the versatility of this technique for use in a broad range of experimental neuroscience settings.

<table border="0" cellpadding="0" cellspacing="0" width="564">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">Name of Material/ Equipment</td>
			<td style="width:89px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">ICR outbred mice</td>
			<td style="width:89px;">Harlan</td>
			<td style="width:119px;">Hsd:ICR (CD-1)</td>
			<td style="width:167px;">This strain is also known as CD-1</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">FVB inbred mice</td>
			<td style="width:89px;">The Jackson Laboratory</td>
			<td style="width:119px;">1800</td>
			<td style="width:167px;">5-6 weeks of age</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:189px;">Nestlets</td>
			<td style="width:89px;">Lab Supply</td>
			<td style="width:119px;">NESTLETS</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:189px;">Shepherd shacks</td>
			<td style="width:89px;">Lab Supply</td>
			<td style="width:119px;">SS-mouse</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">High fat rodent chow</td>
			<td style="width:89px;">Purina Mills</td>
			<td style="width:119px;">PicoLab Mouse diet 20, #5058</td>
			<td style="width:167px;">This is our standard breeder chow</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:189px;">High fat rodent chow (alternative)</td>
			<td style="width:89px;">Harlan Laboratories</td>
			<td style="width:119px;">Teklad Global 19% protein rodent diet #2019S</td>
			<td style="width:167px;">If low phytoestrogen, autoclavable diet is needed</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:189px;">Injection syringe</td>
			<td style="width:89px;">Hamilton</td>
			<td style="width:119px;">7653-01</td>
			<td style="width:167px;">10 ml syringe</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">Injection needles</td>
			<td style="width:89px;">Hamilton</td>
			<td style="width:119px;">7803-04, RN 6PK PT4</td>
			<td style="width:167px;">32 gauge, for standard P0 injections</td>
		</tr>
		<tr height="126">
			<td height="126" style="height:126px;width:189px;">Metal plate for cryoanesthesia</td>
			<td style="width:89px;">McMaster Carr</td>
			<td style="width:119px;">8975K439</td>
			<td style="width:167px;">Raw aluminum plate, 6&#8221; x 12&#8221;, 0.25&#8221; thick, will need to be cut into 3 equal pieces and edges sanded by local machine shop</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">Small animal stereotaxic device with digital readout</td>
			<td style="width:89px;">David Kopf Instruments</td>
			<td style="width:119px;">Model 940</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">Universal syringe holder with needle support foot</td>
			<td style="width:89px;">David Kopf Instruments</td>
			<td style="width:119px;">Model 1772-F1</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:189px;">Neonatal frame</td>
			<td style="width:89px;">Stoelting</td>
			<td style="width:119px;">51625</td>
			<td style="width:167px;">Officially called a mouse and neonatal rat adaptor</td>
		</tr>
		<tr height="126">
			<td height="126" style="height:126px;width:189px;">Biohazard disposal bags with sterile indicator</td>
			<td style="width:89px;">VWR</td>
			<td style="width:119px;">14220-030</td>
			<td style="width:167px;">Important! &#8211; Check with local veterinary and environmental safety staff to learn your institute&#8217;s protocol for biohazard waste disposal</td>
		</tr>
	</tbody>
</table>

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.

Successful intraventricular viral injection yields widespread and robust neuronal expression. Here we evaluated viral transduction using YFP or tdTomato fluorescent genes under the control of chicken beta actin promoter (CBA promoter). These constructs were packaged into AAV8 and injected into the lateral ventricles of neonatal (P0) ICR mice. High viral titers (1010 particles per hemisphere) resulted in dense labeling of the olfactory bulb, striatum, cerebral cortex, hippocampus and cerebellum (Figure ...

Watch this Scientific Journal Video about Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction at JoVE.com

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

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 ...

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JoVE Journal

Neuroscience

58.5K 조회수.  Baylor College of Medicine. 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.

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

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

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 ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the hematogenous route, which involves infection of the endothelial cells and pericytes of the brain. The second is the intracerebroventricular (ICV) route. Once within the central nervous system (CNS), viruses may spread to the subarachnoid space, meninges, and choroid plexus via the cerebrospinal fluid. In experimental models, the earliest stages of CNS viral distribution are not well characterized, and it is unclear whether only certain cells are initially infected. Here, we have analyzed the distribution of cytomegalovirus (CMV) particles during the acute phase of infection, termed primary viremia, following ICV or intravascular (IV) injection into the neonatal mouse brain. In the ICV injection model, 5 &#181;l of murine CMV (MCMV) or fluorescent microbeads were injected into the lateral ventricle at the midpoint between the ear and eye using a 10-&#181;l syringe with a 27 G needle. In the IV injection model, a 1-ml syringe with a 35 G needle was used. A transilluminator was used to visualize the superficial temporal (facial) vein of the neonatal mouse. We infused 50 &#181;l of MCMV or fluorescent microbeads into the superficial temporal vein. Brains were harvested at different time points post-injection. MCMV genomes were detected using the in situ hybridization method. Fluorescent microbeads or green fluorescent protein expressing recombinant MCMV particles were observed by fluorescent microscopy. These techniques can be applied to many other pathogens to investigate the pathogenesis of encephalitis.

Here, we describe a simple method of intracerebroventricular and intravascular injection of viral particles or fluorescent microbeads into the neonatal mouse brain. The localization pattern of the virus and nanoparticles could be detected by microscopic evaluation or by in situ hybridization.

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the ...

Neurobehavioral Assessments in a Mouse Model of Neonatal Hypoxic-ischemic Brain Injury

We performed unilateral carotid artery occlusion on CD-1 mice to create a neonatal hypoxic-ischemic (HI) model and investigated the effects of neonatal HI brain injury by studying neurobehavioral functions in these mice compared to non-operated (i.e., normal) mice. During the study, Rice-Vannucci&#39;s method was used to induce neonatal HI brain damage in postnatal day 7-10 (P7-10) mice. The HI operation was performed on the pups by unilateral carotid artery ligation and exposure to hypoxia (8% O2 and 92% N2 for 90 min). One week after the operation, the damaged brains were evaluated with the naked eye through the semi-transparent skull and were categorized into subgroups based on the absence (&#34;no cortical injury&#34; group) or presence (&#34;cortical injury&#34; group) of cortical injury, such as a lesion in the right hemisphere. On week 6, the following neurobehavioral tests were performed to evaluate the cognitive and motor functions: passive avoidance task (PAT), ladder walking test, and grip strength test. These behavioral tests are helpful in determining the effects of neonatal HI brain injury and are used in other mouse models of neurodegenerative diseases. In this study, neonatal HI brain injury mice showed motor deficits that corresponded to right hemisphere damage. The behavioral test results are relevant to the deficits observed in human neonatal HI patients, such as cerebral palsy or neonatal stroke patients. In this study, a mouse model of neonatal HI brain injury was established and showed different degrees of motor deficits and cognitive impairment compared to non-operated mice. This work provides basic information on the HI mouse model. MRI images demonstrate the different phenotypes, separated according to the severity of brain damage by motor and cognitive tests.

We performed unilateral carotid artery occlusion on postnatal day 7-10 CD-1 mouse pups to create a neonatal hypoxic-ischemic (HI) model and investigated the effects of HI brain injury. We studied neurobehavioral functions in these mice compared to non-operated normal mice.

We performed unilateral carotid artery occlusion on CD-1 mice to create a neonatal hypoxic-ischemic (HI) model and investigated the effects of neonatal HI ...

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, ...

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

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

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

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

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages over conventional multi-photon calcium imaging systems: (1) compact; (2) light-weighted; (3) affordable; and (4) allows recording from freely behaving animals. This protocol describes brain surgeries for deep brain in vivo calcium imaging using a custom-developed miniscope recording system. The preparation procedure consists of three steps, including (1) stereotaxically injecting the virus at the desired brain region of a mouse brain to label a specific subgroup of neurons with genetically encoded calcium sensor; (2) implantation of gradient-index (GRIN) lens that can relay calcium image from deep brain region to the miniscope system; and (3) affixing the miniscope holder over the mouse skull where miniscope can be attached later. To perform in vivo calcium imaging, the miniscope is fastened onto the holder, and neuronal calcium images are collected along with simultaneous behavior recordings. The present surgery protocol is compatible with any commercial or custom-built single-photon and two-photon imaging systems for deep brain in vivo calcium imaging.

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

Miniscope in vivo calcium imaging is a powerful technique to study neuronal dynamics and microcircuits in freely behaving mice. This protocol describes performing brain surgeries to achieve good in vivo calcium imaging using a miniscope.

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages ...

Robust and Highly Reproducible Generation of Cortical Brain Organoids for Modelling Brain Neuronal Senescence In Vitro

Brain organoids are three-dimensional models of the developing human brain and provide a compelling, cutting-edge platform for disease modeling and large-scale genomic and drug screening. Due to the self-organizing nature of cells in brain organoids and the growing range of available protocols for their generation, issues with heterogeneity and variability between organoids have been identified. In this protocol paper, we describe a robust and replicable protocol that largely overcomes these issues and generates cortical organoids from neuroectodermal progenitors within 1 month, and that can be maintained for more than 1 year. This highly reproducible protocol can be easily carried out in a standard tissue culture room and results in organoids with a rich diversity of cell types typically found in the developing human cortex. Despite their early developmental make-up, neurons and other human brain cell types will start to exhibit the typical signs of senescence in neuronal cells after prolonged in vitro culture, making them a valuable and useful platform for studying aging-related neuronal processes. This protocol also outlines a method for detecting such senescent cells in cortical brain organoids using senescence-associated beta-galactosidase staining.

Robust and Highly Reproducible Generation of Cortical Brain Organoids for Modelling Brain Neuronal Senescence In Vitro

In this study, we provide a detailed technique for a simple yet robust cortical organoid culture system using standard feeder-free hPSC cultures. This is a rapid, efficient, and reproducible protocol for generating organoids that model aspects of brain senescence in vitro.

Brain organoids are three-dimensional models of the developing human brain and provide a compelling, cutting-edge platform for disease modeling and ...

Whole-Brain Single-Cell Imaging and Analysis of Intact Neonatal Mouse Brains Using MRI, Tissue Clearing, and Light-Sheet Microscopy

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of structural changes caused by genetic or environmental perturbations. Whole-brain imaging results in more accurate quantification of cells and the study of region-specific differences that may be missed with commonly used microscopy of physically sectioned tissue. Using light-sheet microscopy to image cleared brains greatly increases acquisition speed as compared to confocal microscopy. Although these images produce very large amounts of brain structural data, most computational tools that perform feature quantification in images of cleared tissue are limited to counting sparse cell populations, rather than all nuclei.
Here, we demonstrate NuMorph (Nuclear-Based Morphometry), a group of analysis tools, to quantify all nuclei and nuclear markers within annotated regions of a postnatal day 4 (P4) mouse brain after clearing and imaging on a light-sheet microscope. We describe magnetic resonance imaging (MRI) to measure brain volume prior to shrinkage caused by tissue clearing dehydration steps, tissue clearing using the iDISCO+ method, including immunolabeling, followed by light-sheet microscopy using a commercially available platform to image mouse brains at cellular resolution. We then demonstrate this image analysis pipeline using NuMorph, which is used to correct intensity differences, stitch image tiles, align multiple channels, count nuclei, and annotate brain regions through registration to publicly available atlases.
We designed this approach using publicly available protocols and software, allowing any researcher with the necessary microscope and computational resources to perform these techniques. These tissue clearing, imaging, and computational tools allow measurement and quantification of the three-dimensional (3D) organization of cell-types in the cortex and should be widely applicable to any wild-type/knockout mouse study design.

This protocol describes methods for conducting magnetic resonance imaging, clearing, and immunolabeling of intact mouse brains using iDISCO+, followed by a detailed description of imaging using light-sheet microscopy, and downstream analyses using NuMorph.

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of ...