This work was supported by American Heart Association postdoctoral fellowship (AY), Defense Advanced Research Projects Agency (DARPA) Reorganization and Plasticity to Accelerate Injury Recovery (REPAIR; N66001-10-C-2010), R01.NS073940, and by the UCSF Neuroscience Imaging Center. This work was also supported by the Eunice Kennedy Shiver National Institute of Child Health &#38; Human Development of the National Institutes of Health under Award Number K12HD073945, the Washington National Primate Research Center (WaNPCR, P51 OD010425), and the Center for Neurotechnology (CNT, a National Science Foundation Engineering Research Center under Grant EEC-1028725). We thank Camilo Diaz-Botia, Tim Hanson, Viktor Kharazia, Daniel Silversmith, Karen J. MacLeod, Juliana Milani, and Blakely Andrews for their help with the experiments and Nan Tian, Jiwei He, Peter Ledochowitsch, Michel Maharbiz, and Toni Haun for technical help.

All procedures have been approved by the University of California, San Francisco Institutional Animal Care and Use Committee (IACUC) and are compliant with the Guide for the Care and Use of Laboratory Animals. The following procedure was performed using two adult male rhesus macaques of 8 and 7 years of age, weighing 17.5 kg and 16.5 kg (monkey G and monkey J), respectively.
NOTE: Use standard aseptic techniques for all surgical procedures.
1. Baseline Imaging
<ol>
	<li>Sedate and intubate the animal and maintain general anesthesia under isoflurane (concentration changed between 0 to 5% as needed, depending on vital signs such as respiratory rate and heart rate). Monitor the animal&#8217;s temperature, heart rate, oxygen saturation, electrocardiographic responses, and end-tidal partial pressure of CO2 throughout the procedure.</li>
	<li>Place the animal in an MR-compatible stereotaxic frame (see the Table of Materials) in which it will remain throughout the procedure. Connect the animal to a portable MR-compatible isoflurane machine and transport it to the MRI scanner (3 T).</li>
	<li>Acquire standard T1 (flip angle = 9&#176;, repetition time/echo time = 668.6, matrix size = 192 x 192 x 80 , slice thickness = 1 mm) and T2 (flip angle = 130&#176;, repetition time/echo time = 52.5, matrix size = 256 x 256 x 45, slice thickness = 1 mm) anatomical MR images as baseline reference and for surgical planning.</li>
	<li>Recover the animal from anesthesia and transport it to the animal housing area.</li>
	<li>Use the acquired T1 and T2 images to determine the placement of the craniotomy. The location of the craniotomy can be precisely determined by matching the cortical area of interest from MR with the macaque brain atlas (Paxinos, Huang, Petride, Toga 2nd Edition). In addition, these baseline images can provide an estimate for the locations of viral infusion. The cortical regions of interest demonstrated here are M1 and S1.</li>
</ol>
2. MR-Compatible Chamber Implantation
<ol>
	<li>Sedate and intubate the animal and maintain general anesthesia with standard anesthetic monitoring as in 1.1.</li>
	<li>Place the animal in the MR-compatible stereotaxic frame in which it will remain throughout chamber implantation and viral vector delivery. Connect the animal to a portable MR-compatible isoflurane machine.</li>
	<li>Create a sagittal incision approximately 2 cm from the midline with a length of about 5 cm with a scalpel.</li>
	<li>Remove the underlying soft tissue from the skull using elevators (see Table of Materials).</li>
	<li>Create a circular craniotomy (2.5 cm diameter) to cover the pre-planned trajectories for injections using a trephine (see Table of Materials).
	<ol>
		<li>Lower the centering point of the trephine past the edge of the trephine. Create an indentation at the center of the planned craniotomy sufficiently deep in the skull to anchor the trephine using the adjustable centering point of the trephine. Exercise caution to avoid completely penetrating through the depth of the skull as this could cause damage to the underlying neural tissue.</li>
		<li>Periodically flush the area with saline as needed to maintain tissue moisture throughout the craniotomy.</li>
		<li>Once the center has been made, lower the trephine onto the skull and rotate the trephine clockwise and counterclockwise while applying downward pressure until the bone cap can be removed with forceps. Exercise caution to avoid damaging the underlying tissue with the trephine.</li>
	</ol>
	</li>
	<li>Thread a fine suture (size 6-0) through&#160;the dura in the center of the craniotomy and lift the dura by gently pulling the suture from the surface of the brain creating a tent in the center of the craniotomy.</li>
	<li>Next, puncture the dura close to the center of the tent using fine ophthalmic scissors to avoid damaging the brain. Then cut the dura from the center to the edge of the craniotomy and continue along the edge with fine opthalmic scissors.</li>
	<li>Mount the cylindrical custom-made CED MR-compatible chamber (Figure 1; see section 4 for production instructions) to the skull on top of the craniotomy to provide cannula support during CED infusion such that the curvature of the chamber flange aligns well with the curvature of the skull.</li>
	<li>Secure the implants to the skull using either plastic screws and dental acrylic or a few titanium screws.</li>
</ol>
3. Viral Vector Delivery
<ol>
	<li>Soon after implanting the MR-compatible chamber and inserting the cannula injection grid (Figure 1A-B, Supplementary Figure 1) into the chamber, fill the grid with saline (0.9% NaCl) for visualizing the injection locations via MR anatomical images and fill the chamber cavities with wet sterile absorbable gelatin to maintain moisture of the brain (Figure 1F).</li>
	<li>Cover the skin and the cylinder with a sterile antimicrobial incise drape to maintain the sterility of the cylinders during transport and MR infusions. Place a vitamin E capsule to mark the top of the injection grid for positive identification.</li>
	<li>While the animal remains intubated, detach the endotracheal tube from the anesthesia circuit and reattach it to a portable MR-compatible isoflurane machine and transport the animal to the MRI scanner.</li>
	<li>Acquire T1 images (flip angle = 9&#176;, repetition time/echo time = 668.6, matrix size = 192 x 192, slice thickness= 1 mm, 80 slices) to calculate the distance from the top of the injection grid and the cortical surface. The vitamin E capsule is clearly visible in T1 images and should be used as a marker for the top of the injection grid (Figure 2A). Use the MR imaging software&#8217;s ruler tool to measure the distance between the top of the injection grid and the cortical surface.</li>
	<li>Acquire T2 images (flip angle = 130&#176;, repetition time/echo time = 52.5, matrix size = 256 x 256, slice thickness = 1 mm, 45 slices) to determine the optimal cannula guides for each site based on the targeted site of infusion (Figure 2B-C). As mentioned earlier, the cannula grid is filled with saline which is best visible in T2 images. Using the MR imaging software, scroll through the coronal and sagittal planes to find the target location of infusion.</li>
	<li>After thawing the viral vector for a few hours at room temperature, mix the viral vector with the MR contrast agent gadoteridol (Ratio of 250:1; 2mM Gd-DTPA, see Table of Materials) by pipette or vortex mixing. 
	NOTE: The presented technique was tested for AAV vectors with a CamKIIa promoter driving expression of C1V1 fused to EYFP (AAV2.5-CamKII-C1V1-EYFP, titer: 2.5 x 1012 virus molecules/mL; see Table of Materials).</li>
	<li>Load the mixed virus into a 0.2 mL high pressure IV tubing (see Table of Materials).</li>
	<li>Establish a sterile field outside the MR scanner for preparing the viral infusion line.</li>
	<li>Using high pressure IV tubing, connect a long extension line (approximately 3-5 meters, depending on the location of the infusion pump with respect to the MR bore) to an MR-compatible 3 mL syringe and prime with saline.</li>
	<li>Connect the distal end of the long extension line to the proximal end of the 0.2 mL IV tubing loaded with the virus and attach the reflux-resistant cannula with a 1 mm stepped tip (Figure 2D; see section 5 for cannula production instructions) to the distal end of this assembly with a clamp style catheter connector (see Table of Materials) (Figure 2E).</li>
	<li>Lastly, set the syringe in an MR-compatible pump (see Table of Materials). Place the pump controller in the scanner control room as it is not MR-compatible.</li>
	<li>Using the baseline anatomical MR images obtained before, select the cannula injection grid location and insertion depth needed to reach the target infusion site. Mark the insertion depth on the cannula using sterile tape.</li>
	<li>Begin the infusion at a rate of 1 &#956;L/min and visually validate the flow of the fluid in the infusion line by observing the ejection of fluid from the cannula tip.</li>
	<li>Insert the cannula manually through the injection grid to the target depth while maintaining flow in the infusion line, as this will prevent the penetrated tissue from clogging the cannula during insertion.</li>
	<li>Acquire fast (2 minute) flash T1 weighted images (flip angle = 30&#176;, repetition time/echo time = 3.05, matrix size = 128 x 128, slice thickness = 1 mm, 64 slices) for online monitoring of the viral vector infusion.</li>
	<li>After infusing ~10 &#956;L of the vector such that enough virus is infused to detect in the MR scanner, obtain MR images to verify correct cannula placement as evident by the observed spread of the virus. If the depth of the inserted cannula is incorrect, adjust the depth accordingly or slowly remove the cannula and re-attempt the insertion as in 3.12.</li>
	<li>Monitor the infusion via guidance of online MR images (flash T1 weighted) and increase the infusion rate to 5 &#181;L/min by 1 &#181;L/min steps every few minutes (Figure 3A).</li>
	<li>After infusing approximately 40 &#956;L of the viral vector, begin reducing the infusion rate by 1 &#956;L/min steps and stop the infusion after ~50 &#956;L is injected and leave the cannula in place for 10 minutes.</li>
	<li>Slowly remove the cannula from the brain and move to the next location. Repeat steps 3.9 to 3.19&#160;for each location.</li>
	<li>Cover the cylinder with a sterile drape at the end of injections, before animal transport. Then transport the animal back to the operating room.</li>
	<li>Either explant the MR-compatible chamber and close the surgical incision by replacing the bone flap and overlying muscle and suturing the skin using standard aseptic techniques or replace the chamber with a titanium cylinder and artificial dura (see Yazdan-Shahmorad et al. 20169) for neural recording and stimulation.</li>
</ol>
4. MR-compatible Chamber Production
<ol>
	<li>Fabricate the nylon cannula injection grid (Figure 1A-B) by machining according to the specified dimensions (Supplementary Figure 1).</li>
	<li>3D print the MR-compatible cylinder out of acrylonitrile butadiene styrene (ABS0 plastic (Figure 1C-E) (see Supplementary File 1&#8211;3; see Table of Materials for 3D printer and materials). 
	NOTE: One design maintains a fixed cannula injection grid within the MR-compatible cylinder (Figure 1C), while another allows for rotation of the grid, extending the injection region (Figure 1D-E).</li>
	<li>Thread the cannula injection grid and tap the corresponding cavity of the MR-compatible chamber such that both pieces exhibit the same threading. 
	NOTE: Any type of threading can be used provided that both components have the same threading.</li>
	<li>Insert the nylon cannula injection grid by screwing into the tapped hole of the MR-compatible cylinder.</li>
</ol>
5. Reflux-resistant Cannula Production13
<ol>
	<li>Cut a 30 cm-long silica tubing with 0.32 mm ID and 0.43 mm OD for the inner silica tubing (see Table of Materials) using a razor blade.</li>
	<li>Cut a 7.5 cm-long and a 5 cm-long silica tubing with 0.45 mm ID and 0.76 mm OD for the outer silica tubing (see Table of Materials).</li>
	<li>Adhere the 7.5 cm outer tubing to the 30 cm inner tubing with cyanoacrylate such that the inner tube extends 1 mm beyond the outer tube, making the reflux-resistant step (Figure 2D). To ensure that the cyanoacrylate does not enter the inside of the inner tubing, place the cyanoacrylate on the outside of the inner tubing far from the cannula tip.</li>
	<li>Paste the 5 cm-long silica tubing to the other end of the inner tubing for attachment to the clamp style catheter connector (see step 3.10).</li>
</ol>

<ol>
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Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Saccade+modulation+by+optical+and+electrical+stimulation+in+the+macaque+frontal+eye+field.">Saccade modulation by optical and electrical stimulation in the macaque frontal eye field.</a> Journal of Neuroscience. 33 (42), 16684-16697 (2013).</li><li>Gerits, 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=Optogenetically+induced+behavioral+and+functional+network+changes+in+primates.">Optogenetically induced behavioral and functional network changes in primates.</a> Current Biology. 22 (18), 1722-1726 (2012).</li><li>Jazayeri, M., Lindbloom-Brown, Z., Horwitz, G. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Saccadic+eye+movements+evoked+by+optogenetic+activation+of+primate+V1.">Saccadic eye movements evoked by optogenetic activation of primate V1.</a> Nature Neuroscience. 15 (10), 1368-1370 (2012).</li><li>Dai, J., Brooks, D. I., Sheinberg, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optogenetic+and+electrical+microstimulation+systematically+bias+visuospatial+choice+in+primates.">Optogenetic and electrical microstimulation systematically bias visuospatial choice in primates.</a> Current Biology. 24 (1), 63-69 (2014).</li><li>Afraz, A., Boyden, E. S., DiCarlo, J. 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PNS has financial interest in Neuralink Corp., a company that is developing clinical therapies using brain stimulation.

Here, we outline a feasible and efficient technique for achieving large-scale optogenetic expression in NHP primary somatosensory and motor cortex by MR-guided CED. The use of MR-guided CED presents significant advantages over traditional methods of viral infusion in the NHP brain. One such advantage is the ability to attain expression over large areas with fewer required infusions. For instance, with conventional methods, multiple injections of 1-2 &#181;L of the vector yield expression in a 2-3 mm diameter region1,2,5,17. While previous attempts at achieving large-scale vector spread have yielded expression volumes of approximately 10 mm3 with multiple injections18, here we present a method to achieve expression over 200 mm3 with only a few infusions. &#160;A single infusion of 50 &#181;L by CED can achieve vector spread up to 10 mm from the injection site for cortical CED9 and full coverage of frontal cortical areas with thalamic CED14.
Furthermore, it has been shown that using CED can achieve more homogeneous distribution of the injected agent12,19,20,21,22, leading to uniform expression levels around the site of viral vector infusion, compared to traditional microinjections. In our experiments employing CED, we observed uniformly high expression levels with approximately 70-80% of neurons expressing the opsin8,9 (Figure 5). In contrast, conventional diffusion-based injections exhibit regions of high expression concentrated near injection sites enveloped by weakening expression levels4,5,17. The reduced need for multiple injections renders CED infusion a time-efficient method of obtaining large-scale, uniform optogenetic expression. Due to the smaller number of injections and faster infusion rates, we were able to plan and infuse the viral vector under the guidance of MRI. The use of online MR images enables precise targeting of infusions and the monitoring of the spread of the viral vector during infusion. However, if the initial depth of the inserted cannula is incorrect, this may yield expression in undesired locations as a result of the initial infused ~10 &#181;L needed to visually detect the infusion via MR. Alternatively, commercial neurosurgical navigation systems utilizing fiducial markers can also be employed in place of MR guidance. Additionally, the costs of online imaging may be circumvented by marking the desired depth of insertion on the cannula prior to insertion and visually confirming complete infusion through the infusion line, although the accuracy of the cannula placement would be reduced. However, as seen by the unsuccessful injection in M1 of monkey G (Figure 5D), MR guidance can be critical for ensuring successful infusion.
Previous studies have demonstrated the safety of CED infusions of both non-viral and viral particles with no observed tissue damage outside the cannula tract and no behavioral deficits9,11,12,13,14,15,20. Here and in our previous publications9,14 we report similar results following CED infusion of optogenetic viral vectors. In addition to no observed behavioral deficits following infusion, NeuN and Nissl9,14,15 staining revealed neuronal cell death and gliosis was limited only to the cannula tract, as has been reported with diffusion-based methods5. To minimize the damage from the cannula track, we can potentially reduce the diameter of the cannula. However, further investigation is needed to assess the effect of cannula size reduction on achieving large areas of expression. Moreover, because high infusion rates can also lead to tissue damage13, it is recommended to maintain an infusion rate at or below 5 &#181;L/min. For more detailed information about the CED technique please see our previous publication9.
The use of MR-guided CED for attaining broad regions of targeted optogenetic expression in the primate cortex can lead to further investigations of large-scale circuit dynamics, neural plasticity, and network connectivity.

Optogenetics is a powerful tool that allows for the manipulation of neural activity and the study of network connections in the brain. Implementing this technique in non-human primates (NHPs) has the potential to enhance our understanding of large-scale neural computation, cognition, and behavior in the primate brain. Although optogenetics has been successfully implemented in NHPs in recent years1,2,3,4,5,6,7, a challenge that researchers face is achieving high levels of expression across large brain areas in these animals. Here, we are providing an efficient and simplified approach to achieve high levels of optogenetic expression across large areas of the cortex in macaques. This technique has great potential to improve current optogenetic studies in these animals in combination with state-of-the-art recording8,9 and optical stimulation10 technologies.
Convection enhanced delivery (CED) is an established method of delivery of pharmacological agents and other large molecules, including viral vectors, to the central nervous system11,12,13. Whereas conventional delivery methods involve multiple low volume infusions distributed across small regions of the brain, CED can achieve broader and more even agent distribution with fewer infusions. Pressure-driven bulk fluid flow (convection) during infusion allows for more widely and uniformly distributed transduction of the target tissue when delivering viral vectors with CED. In recent studies, we demonstrated the transduction and subsequent optogenetic expression of large areas of primary motor (M1) and somatosensory (S1) cortices9 and thalamus14 using magnetic resonance (MR)-guided CED.
Here, we outline the use of CED to achieve optogenetic expression across large cortical areas with only a few cortical injections.

<table>
	<tbody>
		<tr>
			<td>0.2 mL High Pressure IV Tubing</td>
			<td>Smiths Medical Inc., Dublin, OH, USA</td>
			<td>533640</td>
			<td></td>
		</tr>
		<tr>
			<td>0.32 mm ID, 0.43 mm OD Silica Tubing</td>
			<td>Polymicro Technologies</td>
			<td>1068150027</td>
			<td></td>
		</tr>
		<tr>
			<td>0.45 mm ID, 0.76 mm OD Silica Tubing</td>
			<td>Polymicro Technologies</td>
			<td>1068150625</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV2.5-CamKII-C1V1-EYFP</td>
			<td>Penn Vector Core, University of Pennsylvania</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ABS plastic</td>
			<td>Stratasys, MN, USA</td>
			<td>ABSplus-P430</td>
			<td></td>
		</tr>
		<tr>
			<td>Antimicrobial incise drape</td>
			<td>3M</td>
			<td>6650EZ</td>
			<td>Ioban Drape</td>
		</tr>
		<tr>
			<td>Dental Acrylic</td>
			<td>Henry Schein, Inc.</td>
			<td>1013117</td>
			<td>Acrylic Bonding Agent</td>
		</tr>
		<tr>
			<td>Elevators</td>
			<td>VWR International, LLC.</td>
			<td>10196-564</td>
			<td>Langenbeck Elevator, Wide Tip</td>
		</tr>
		<tr>
			<td>Fine suture</td>
			<td>McKesson Medical-Surgical Inc.</td>
			<td>1034505</td>
			<td></td>
		</tr>
		<tr>
			<td>Gadoteridol</td>
			<td>Prohance, Bracco Diagnostics, Princeton, NJ</td>
			<td>0270-1111-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser for light stimulation</td>
			<td>Omicron-Laserage, Germany</td>
			<td>PhoxX 488-60</td>
			<td></td>
		</tr>
		<tr>
			<td>MR compatible 3cc syringe</td>
			<td>Harvard apparatus, Holliston, MA, USA</td>
			<td>59-8377</td>
			<td></td>
		</tr>
		<tr>
			<td>MR Imaging Software</td>
			<td>Pixmeo</td>
			<td>OsiriX MD 10.0</td>
			<td></td>
		</tr>
		<tr>
			<td>MR-Compatible Pump</td>
			<td>Harvard apparatus, Holliston, MA, USA</td>
			<td>Harvard PHD 2000</td>
			<td></td>
		</tr>
		<tr>
			<td>MR-compatible stereotaxic frame</td>
			<td>KOPF</td>
			<td>1430M MRI</td>
			<td></td>
		</tr>
		<tr>
			<td>Perifix Clamp Style Catheter Connector</td>
			<td>B-Braun, Bethlehem, PA, USA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic Screws</td>
			<td>Plastics 1</td>
			<td>0-80 x 1/8N</td>
			<td>Nylon screws</td>
		</tr>
		<tr>
			<td>Titanium screws</td>
			<td>Crist Instrument Co., Inc.</td>
			<td>6-YCX-0312</td>
			<td>Self-tapping bone screws</td>
		</tr>
		<tr>
			<td>Trephine</td>
			<td>GerMedUSA Inc,</td>
			<td>SKU:GV70-42</td>
			<td></td>
		</tr>
		<tr>
			<td>uPrinter SE 3D printer</td>
			<td>Stratasys, MN, USA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Vitamin E Capsule</td>
			<td>Pure Encapsulations, LLC.</td>
			<td>DE1</td>
			<td></td>
		</tr>
		<tr>
			<td>Wet sterile absorbable gelatin</td>
			<td>Pfizer Inc.</td>
			<td>AZL0009034201</td>
			<td>Gelfoam</td>
		</tr>
	</tbody>
</table>

convection enhanced delivery of optogenetic adeno-associated viral vector to the cortex of rhesus macaque under guidance of online mri images

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we demonstrate the use of magnetic resonance (MR)-guided convection enhanced delivery (CED) of optogenetic viral vectors into primary somatosensory (S1) and motor (M1) cortices of macaques to obtain efficient, widespread cortical expression of light-sensitive ion channels. Adeno-associated viral (AAV) vectors encoding the red-shifted opsin C1V1 fused to yellow fluorescent protein (EYFP) were injected into the cortex of rhesus macaques under MR-guided CED. Three months post-infusion, epifluorescent imaging confirmed large regions of optogenetic expression (&#62;130 mm2) in M1 and S1 in two macaques. Furthermore, we were able to record reliable light-evoked electrophysiology responses from the expressing areas using micro-electrocorticographic arrays. Later histological analysis and immunostaining against the reporter revealed widespread and dense optogenetic expression in M1 and S1 corresponding to the distribution indicated by epifluorescent imaging. This technique enables us to obtain expression across large areas of the cortex within a shorter period of time with minimal damage compared to the traditional techniques and can be an optimal approach for optogenetic viral delivery in large animals such as NHPs. This approach demonstrates great potential for network-level manipulation of neural circuits with cell-type specificity in animal models evolutionarily close to humans.&#160;

Convection Enhanced Delivery (CED) under MRI Guidance
The spread of the viral vector was monitored during CED infusion under the guidance of online MR images (Figure 3A). In this study, S1 and M1 of two monkeys were targeted (Figure 3B). The three-dimensional distribution volumes were estimated in a post-hoc analysis of the MR images (Figure 3C) as described by Yazdan-Shahmorad et al. 20169, confirming coverage of large areas of cortex. The M1 infusions for monkey G were done without MR guidance due to time constraints.
Validation of Viral Expression
Large cortical optogenetic expression was confirmed by epifluorescent imaging, electrophysiological recordings, and histological analysis as described by Yazdan-Shahmorad et al. 20169. We monitored optogenetic expression by surface epifluorescence imaging of the fluorescent reporter2,9 and estimated more than 130 mm2 of expression along the cortical surface in M1 and S1 of two macaques from only three cortical injections9,15. Light stimulation (488 nm, 20 mW power at the tip; see Table of Materials) yielded reliable electrophysiological responses in the expressing areas as measured by semi-transparent micro-electrocorticographic arrays8,9,16 (Figure 4).
Analysis of the MR images yielded an estimated 233 mm3 and 433 mm3 of vector spread in M1 and S1 of monkey J, respectively, and 317 mm3 in S1 of monkey G9. We performed histological analyses on serial coronal sections and observed large-scale optogenetic expression around the infusion sites in M1 and S1 (Figure 5C-I), with an estimated 70-80% of cells expressing the opsin. The expression observed from histology aligned with the estimated expression distribution from epifluorescent imaging (Figure 5A-D). One of the two infusions performed outside of the MR scanner in monkey G was unsuccessful, yielding no expression in the corresponding region (Figure 5D). The spread estimated from MR imaging was completely within the bounds of both the epifluorescent and the histological measures of viral spread (Figure 5E-G). The expansion of the epifluorescent imaging beyond the MR estimate can be attributed to the diffusion of viral particles beyond the advection region. Moreover, the histological estimate extended beyond the epifluorescence estimate due to a lack of optical access beyond the craniotomy. The details of these techniques are included in our previous paper9.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59232/59232fig1.jpg" /> 
Figure 1: MR-compatible cylinder and cannula injection grid. (A,B) custom-designed nylon injection grid. (C) MR-compatible fixed cylinder. (D,E) Rotating MR-compatible cylinder. (F) MR-compatible infusion cylinder with fixed grid position. The arrows point to the cavities that are designed to be filled with wet&#160;sterile absorbable gelatin&#160;keeping the surface of the brain moist for the duration of injection. (G) Cannula inserted in the grid. This figure has been modified from Yazdan-Shahmorad et al. 20169. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/59232fig1large.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/59232/59232fig2.jpg" /> 
Figure 2: Baseline MR imaging and reflux resistant cannula. (A) T1-weighted image of vitamin E that was attached to the top of the injection grid that enables us to measure the distance to the surface of the brain (white arrow). (B) T2-weighted image of the brain helps to plan the location of injection from the cannula grid filled with saline. (C) MR image of the infusion chamber and the saline-filled cannula grid. The orthogonal lines represent the sagittal (yellow) and coronal (purple) planes. (D) Photo of the reflux resistant injection cannula tip with the reflux resistant step (black arrow). (E) Infusion lines. This figure has been modified from Yazdan-Shahmorad et al. 20169. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/59232fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59232/59232fig3.jpg" /> 
Figure 3: MR imaging during infusion and estimated distribution. (A) Spread of 50 &#181;L of the viral vector in coronal sections of Monkey G for one injection site in S1 (shown with an arrow in B).&#160;(B) MRI surface rendering of the cortical surface below the cylinder for Monkeys G and J, respectively. The S1 infusion locations are shown in blue, M1 locations in red. (C) MR Volume reconstruction of the spread of viral vector after CED infusion. Brain is shown in light gray; S1 and M1 infusion volumes are shown in blue and red, respectively. No MR volume reconstruction is available for the M1 infusions for monkey G since they were not done in the MR scanner. This figure has been modified from Yazdan-Shahmorad et al. 20169. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/59232fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59232/59232fig4.jpg" /> 
Figure 4: Electrophysiological recordings of light-evoked activity. Micro-electrocorticography (&#956;ECoG) recordings occurred during pulsed optical stimulation. Recording traces were from the closest electrode to the site of stimulation for examples of M1 (red) and S1 (green) stimulation locations. Shaded areas around the traces represent standard error across 25 trials. The blue squares on the traces show the timing of stimulation (1 ms). The full set of stimulus-triggered waveforms for both sample pairs of stimulation and recording sites are superimposed on the mean waveform as shown on the left side of the panel. This figure has been modified from Yazdan-Shahmorad et al. 20169. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/59232fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59232/59232fig5_v2.jpg" /> 
Figure 5: Histological analysis. (A) Baseline coronal MR image in monkey G. (B) Spread of contrast agent after the infusion for the same MR coronal slice as in A. (C) A coronal tissue section from approximately the same site as in A and B; peroxidase staining reflects expression of the EYFP-reporter. (D) Good alignment is observed between the area of EYFP expression measured with surface epiflourescence (dark green areas) and with histological staining (light green lines). These include the region of vector spread estimated from MR images (white line); white dots indicate injection sites, and the entire black region represents the area exposed by the craniotomy. The two left most injection sites are located in M1, and the two right most sites are located in S1. (E) Low magnification image of the coronal section stained with anti-GFP antibody showing the medio-lateral aspect of YFP expression in the somatosensory cortex of monkey J (areas 1, 2, 3). The black arrowhead indicates the location of the cannula track; the adjacent tissue (black frame) is shown in F, at greater magnification to show laminar distribution of the YFP-positive cells. (F) Densely populated regions of YFP-positive cells are located predominantly in layers II-III and V-VI, and also show typical pyramidal morphology (cells in white frames are further enlarged in panels (G-I). White arrowheads on bottom panels G-I point to typical pyramidal cells in layers II-III (G); layer V (H); and layer VI (I). Scale bars =&#160;2 mm (E); &#160;200 &#956;m (F); 100 &#956;m (G-I). This figure has been modified from Yazdan-Shahmorad et al. 20169 and Yazdan-Shahmorad et al. 201815. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/59232fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: The cannula injection grid prior to threading. The length and diameter of the injection grid are 15.00 mm and 12.00 mm, respectively. The injection grid pattern consists of 0.8 mm diameter holes forming three concentric circles (diameter: 1.6, 3.2 and 4.8 mm) around the center of the grid. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/FigureS1.pdf">Please click here to download this file.</a>
Supplementary Files.&#160;&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59232/Supplementary_Files.zip">Please click here to download these files.</a>

Watch this Scientific Journal Video about Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images at JoVE.com

Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images

Here, we demonstrate magnetic resonance (MR)-guided convection enhanced delivery (CED) of viral vectors into the cortex as an efficient and simplified approach for achieving optogenetic expression across large cortical areas in the macaque brain.

In non-human primate (NHP) optogenetics, infecting large cortical areas with viral vectors is often a difficult and time-consuming task. Here, we ...

convection-enhanced-delivery-optogenetic-adeno-associated-viral

Nanyang Technological University

Research

JoVE Journal

Neuroscience

6.8K Views.  University of Washington. Here, we demonstrate magnetic resonance (MR)-guided convection enhanced delivery (CED) of viral vectors into the cortex as an efficient and simplified approach for achieving optogenetic expression across large cortical areas in the macaque brain.

Video: Convection Enhanced Delivery of Optogenetic Adeno-associated Viral Vector to the Cortex of Rhesus Macaque Under Guidance of Online MRI Images

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 ...

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Optogenetic Functional MRI

The investigation of the functional connectivity of precise neural circuits across the entire intact brain can be achieved through optogenetic functional magnetic resonance imaging (ofMRI), which is a novel technique that combines the relatively high spatial resolution of high-field fMRI with the precision of optogenetic stimulation. Fiber optics that enable delivery of specific wavelengths of light deep into the brain in vivo are implanted into regions of interest in order to specifically stimulate targeted cell types that have been genetically induced to express light-sensitive trans-membrane conductance channels, called opsins. fMRI is used to provide a non-invasive method of determining the brain's global dynamic response to optogenetic stimulation of specific neural circuits through measurement of the blood-oxygen-level-dependent (BOLD) signal, which provides an indirect measurement of neuronal activity. This protocol describes the construction of fiber optic implants, the implantation surgeries, the imaging with photostimulation and the data analysis required to successfully perform ofMRI. In summary, the precise stimulation and whole-brain monitoring ability of ofMRI are crucial factors in making ofMRI a powerful tool for the study of the connectomics of the brain in both healthy and diseased states.

This protocol describes the steps and data analysis required to successfully perform optogenetic functional magnetic resonance imaging (ofMRI). ofMRI is a novel technique that combines high-field fMRI readout with optogenetic stimulation, allowing for cell type-specific mapping of functional neural circuits and their dynamics across the whole living brain.

The investigation of the functional connectivity of precise neural circuits across the entire intact brain can be achieved through optogenetic functional ...

Utilizing 3D Printing Technology to Merge MRI with Histology: A Protocol for Brain Sectioning

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume three-dimensionally. While MRI is an extremely sensitive tool for detecting tissue abnormalities, association of signal changes with an underlying pathological process is usually not straightforward. In the central nervous system, for example, inflammation, demyelination, axonal damage, gliosis, and neuronal death may all induce similar findings on MRI. As such, interpretation of MRI scans depends on the context, and radiological-histopathological correlation is therefore of the utmost importance. Unfortunately, traditional pathological sectioning of brain tissue is often imprecise and inconsistent, thus complicating the comparison between histology sections and MRI. This article presents novel methodology for accurately sectioning primate brain tissues and thus allowing precise matching between histology and MRI. The detailed protocol described in this article will assist investigators in applying this method, which relies on the creation of 3D printed brain slicers. Slightly modified, it can be easily implemented for brains of other species, including humans.

The overall goal of this protocol is to accurately align magnetic resonance imaging (MRI) image volumes with histology sections via the creation of customized 3D-printed brain holders and slicer boxes.

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume ...

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 ...

Automated Segmentation of Cortical Grey Matter from T1-Weighted MRI Images

Within neuroimaging research, a number of recent studies have discussed the impact of between-study differences in volumetric findings that are thought to result from the use of different segmentation tools to generate brain volumes. Here, processing pipelines for seven automated tools that can be used to segment grey matter within the brain are presented. The protocol provides an initial step for researchers aiming to find the most accurate method for generating grey matter volumes from T1-weighted MRI scans. Steps to undertake detailed visual quality control are also included in the manuscript. This protocol covers a range of potential segmentation tools and encourages users to compare the performance of these tools within a subset of their data before selecting one to apply to a full cohort. Furthermore, the protocol may be further generalized to the segmentation of other brain regions.

This protocol describes the process of applying seven different automated segmentation tools to structural T1-weighted MRI scans to delineate grey matter regions that can be used for the quantification of grey matter volume.

Within neuroimaging research, a number of recent studies have discussed the impact of between-study differences in volumetric findings that are thought to ...

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, ...