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

Summary

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.

Abstract

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 (>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. 

Introduction

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 years^{1,2,3,4,5,6,7}, 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 recording^8,9 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 system^11,12,13. 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.

Protocol

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

1. 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’s temperature, heart rate, oxygen saturation, electrocardiographic responses, and end-tidal partial pressure of CO₂ throughout the procedure.
2. 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).
3. Acquire standard T1 (flip angle = 9°, repetition time/echo time = 668.6, matrix size = 192 x 192 x 80 , slice thickness = 1 mm) and T2 (flip angle = 130°, 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.
4. Recover the animal from anesthesia and transport it to the animal housing area.
5. 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 2^nd 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.

2. MR-Compatible Chamber Implantation

1. Sedate and intubate the animal and maintain general anesthesia with standard anesthetic monitoring as in 1.1.
2. 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.
3. Create a sagittal incision approximately 2 cm from the midline with a length of about 5 cm with a scalpel.
4. Remove the underlying soft tissue from the skull using elevators (see Table of Materials).
5. Create a circular craniotomy (2.5 cm diameter) to cover the pre-planned trajectories for injections using a trephine (see Table of Materials).
   1. 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.
   2. Periodically flush the area with saline as needed to maintain tissue moisture throughout the craniotomy.
   3. 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.
6. Thread a fine suture (size 6-0) through 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.
7. 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.
8. 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.
9. Secure the implants to the skull using either plastic screws and dental acrylic or a few titanium screws.

3. Viral Vector Delivery

1. 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).
2. 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.
3. 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.
4. Acquire T1 images (flip angle = 9°, 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’s ruler tool to measure the distance between the top of the injection grid and the cortical surface.
5. Acquire T2 images (flip angle = 130°, 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.
6. 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).
7. Load the mixed virus into a 0.2 mL high pressure IV tubing (see Table of Materials).
8. Establish a sterile field outside the MR scanner for preparing the viral infusion line.
9. 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.
10. 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).
11. 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.
12. 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.
13. Begin the infusion at a rate of 1 μL/min and visually validate the flow of the fluid in the infusion line by observing the ejection of fluid from the cannula tip.
14. 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.
15. Acquire fast (2 minute) flash T1 weighted images (flip angle = 30°, 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.
16. After infusing ~10 μL 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.
17. Monitor the infusion via guidance of online MR images (flash T1 weighted) and increase the infusion rate to 5 µL/min by 1 µL/min steps every few minutes (Figure 3A).
18. After infusing approximately 40 μL of the viral vector, begin reducing the infusion rate by 1 μL/min steps and stop the infusion after ~50 μL is injected and leave the cannula in place for 10 minutes.
19. Slowly remove the cannula from the brain and move to the next location. Repeat steps 3.9 to 3.19 for each location.
20. Cover the cylinder with a sterile drape at the end of injections, before animal transport. Then transport the animal back to the operating room.
21. 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.

4. MR-compatible Chamber Production

1. Fabricate the nylon cannula injection grid (Figure 1A-B) by machining according to the specified dimensions (Supplementary Figure 1).
2. 3D print the MR-compatible cylinder out of acrylonitrile butadiene styrene (ABS0 plastic (Figure 1C-E) (see Supplementary File 1–3; 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).
3. 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.
4. Insert the nylon cannula injection grid by screwing into the tapped hole of the MR-compatible cylinder.

5. Reflux-resistant Cannula Production¹³

1. 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.
2. 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).
3. 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.
4. 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).

Results

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 (

Discussion

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 µL of the vector yield expression in a 2-3 mm diameter region

Materials

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