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>

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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 region<sup c...

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 3 cc 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 (<strong class="xfi...

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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 ...

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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 ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

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 ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Automated 3D Brain and Skull Modeling from MRI for NHP Neurosurgical Planning

A Neural Implant Design Toolbox for Nonhuman Primates

This paper describes an in-house method of 3D brain and skull modeling from magnetic resonance imaging (MRI) tailored for nonhuman primate (NHP) neurosurgical planning. This automated, computational software-based technique provides an efficient way of extracting brain and skull features from MRI files as opposed to traditional manual extraction techniques using imaging software. Furthermore, the procedure provides a method for visualizing the brain and craniotomized skull together for intuitive, virtual surgical planning. This generates a drastic reduction in time and resources from those required by past work, which relied on iterative 3D printing. The skull modeling process creates a footprint that is exported into modeling software to design custom-fit cranial chambers and headposts for surgical implantation. Custom-fit surgical implants minimize gaps between the implant and the skull that could introduce complications, including infection or decreased stability. By implementing these pre-surgical steps, surgical and experimental complications are reduced. These techniques can be adapted for other surgical processes, facilitating more efficient and effective experimental planning for researchers and, potentially, neurosurgeons.

Author Spotlight: Streamlined Brain and Skull Modeling for Enhanced Neurosurgical Planning in NHP Research

This paper outlines automated processes for nonhuman primate neurosurgical planning based on magnetic resonance imaging (MRI) scans. These techniques use procedural steps in programming and design platforms to support customized implant design for NHPs. The validity of each component can then be confirmed using three-dimensional (3D) printed life-size anatomical models.

This paper describes an in-house method of 3D brain and skull modeling from magnetic resonance imaging (MRI) tailored for nonhuman primate (NHP) ...

Bioengineering

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of optogenetics, however, requires that safety and efficacy be demonstrated in animal models, ideally in non-human primates (NHPs), because of their neurological similarity to humans. The number of candidate vectors that are potentially useful for neuroscience and medicine is vast, and no high-throughput means to test these vectors yet exists. Thus, there is a need for techniques to make multiple spatially and volumetrically accurate injections of viral vectors into NHP brain that can be identified unambiguously through postmortem histology. Described herein is such a method. Injection cannulas are constructed from coupled polytetrafluoroethylene and stainless-steel tubes. These cannulas are autoclavable, disposable, and have low minimal-loading volumes, making them ideal for the injection of expensive, highly concentrated viral vector solutions. An inert, red-dyed mineral oil fills the dead space and forms a visible meniscus with the vector solution, allowing instantaneous and accurate measurement of injection rates and volumes. The oil is loaded into the rear of the cannula, reducing the risk of co-injection with the vector. Cannulas can be loaded in 10 min, and injections can be made in 20 min. This procedure is well suited for injections into awake or anesthetized animals. When used to deliver high-quality viral vectors, this procedure can produce robust expression of optogenetic proteins, allowing optical control of neural activity and behavior in NHPs.

As currently implemented, optogenetics in non-human primates requires injection of viral vectors into the brain. An optimal injection method should be reliable and, for many applications, capable of targeting individual sites of arbitrary depth that are readily and unambiguously identified in postmortem histology. An injection method with these properties is presented.

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in the brain sense various biological conditions in the periphery and transmit the signals to neurons. Microglia, immune cells in the brain, are involved in synaptic development and plasticity. Therefore, the contribution of microglia to neural circuit construction in response to the internal state of the body should be tested critically by intravital imaging of the relationship between microglial dynamics and neuronal activity.
Here, we describe a technique for the simultaneous imaging of microglial dynamics and neuronal activity in awake mice. Adeno-associated virus encoding R-CaMP, a gene-encoded calcium indicator of red fluorescence protein, was injected into layer 2/3 of the primary visual cortex in CX3CR1-EGFP transgenic mice expressing EGFP in microglia. After viral injection, a cranial window was installed onto the brain surface of the injected region. In vivo two-photon imaging in awake mice 4 weeks after the surgery demonstrated that neural activity and microglial dynamics could be recorded simultaneously at the sub-second temporal resolution. This technique can uncover the coordination between microglial dynamics and neuronal activity, with the former responding to peripheral immunological states and the latter encoding the internal brain states.

Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in ...

Longitudinal In Vivo Imaging of Retinal Cells with Tailored Adeno-Associated Virus Serotypes via Confocal Scanning Laser Ophthalmoscopy

The dynamic nature of retinal cellular processes necessitates advancements in gene delivery and live monitoring techniques to enhance the understanding and treatment of ocular diseases. This study introduces an optimized adeno-associated virus (AAV) approach, utilizing specific serotypes and promoters to achieve optimal transfection efficiency in targeted retinal cells, including retinal ganglion cells (RGCs) and M&#252;ller glia. Leveraging the precision of confocal scanning laser ophthalmoscopy (CSLO), this work presents a non-invasive method for in vivo imaging that captures the longitudinal expression of AAV-mediated green fluorescent protein (GFP). This approach eliminates the need for terminal procedures, preserving the continuity of observation and the well-being of the subject. Furthermore, the GFP signal can be traced in AAV-infected RGCs along the visual pathway to the superior colliculus (SC) and lateral geniculate nucleus (LGN), enabling the potential for direct visual pathway mapping. These findings provide a detailed protocol and demonstrate the application of this powerful tool for real-time studies of retinal cell behavior, disease pathogenesis, and the efficacy of gene therapy interventions, offering valuable insights into the living retina and its connections.

This protocol describes the use of adeno-associated virus (AAV) vectors for cell-specific labeling and in vivo imaging using a confocal scanning laser ophthalmoscope (CSLO). This method enables the investigation of different retinal cell types and their contributions to retinal function and disease.

The dynamic nature of retinal cellular processes necessitates advancements in gene delivery and live monitoring techniques to enhance the understanding ...

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

Summary

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Chapters in this video

Intracranial Injection of Adeno-associated Viral Vectors

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic Functional MRI

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

A Neural Implant Design Toolbox for Nonhuman Primates

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Longitudinal In Vivo Imaging of Retinal Cells with Tailored Adeno-Associated Virus Serotypes via Confocal Scanning Laser Ophthalmoscopy