Stereotaxic surgical targeting of the non-human primate caudate and putamen: gene therapy for Huntington’s disease

Summary

Stereotaxic surgery is an invaluable tool to deliver a variety of gene therapy constructs to the non-human primate caudate and putamen in pre-clinical studies for the genetic, neurodegenerative disorder, Huntington’s disease (HD). Here we describe in detail how to perform this technique beginning with a pre-surgical magnetic resonance imaging scan to determine surgical coordinates followed by the stereotaxic surgical injection technique. In addition, we include methodology of a full necropsy including brain and peripheral tissue removal and a standard immunohistochemical technique to visualize the injected gene therapy agent.

1. Introduction

Huntington’s disease (HD) is a genetic, neurological disorder that is characterized by a devastating array of symptoms including a hyperkinetic movement disorder, changes in cognitive capabilities, psychiatric manifestations and a robust metabolic syndrome (1). Many of these symptoms are caused by neuronal loss in the caudate and putamen that results from a mutated HTT gene containing an abnormally long stretch of the DNA bases cytosine, adenine and guanine (CAG) (1, 2). Because of the dramatic cell loss observed in these brain regions, the caudate and putamen have historically been the go-to target structures in both pre-clinical studies as well as clinical trials investigating potential therapeutic strategies to treat HD.

Stereotaxic surgery is a powerful tool that has been used for decades to accurately inject therapeutic agents into different brain regions in a variety of different animal species. Prior to the identification of the mutant, disease-causing HTT gene in 1993 (1), stereotaxic surgery was commonly used to deliver excitotoxins into the non-human primate (NHP) caudate and putamen to create a large animal model of HD that replicated some of the key symptoms of the disease (3–6). More recently, lentiviral vectors (LVs) (7) and adeno-associated viral vectors (AAVs) (ongoing studies in our laboratory) have been used to deliver mutant HTT into the caudate and putamen, replicating many of the neuropathological and behavioral changes seen in humans with HD. Pre-clinical studies have employed stereotaxic surgical targeting of the caudate and putamen to evaluate potential therapeutic strategies in some of these NHP models including fetal tissue transplants and trophic factor administration (8–11). More recently, stereotaxic surgery has been used to target the naïve NHP putamen to assess the safety of partially reducing HTT expression via 1) AAVs expressing microRNA constructs (12, 13) or siRNAs delivered through a cannula placed into the putamen (14). With the recent creation of a transgenic HD monkey bearing a fragment of human mutant HTT (15), it is anticipated that stereotaxic surgery will be used to evaluate a variety of therapeutics in this NHP model in the near future.

Here, we describe in detail the methodology that our laboratory uses to surgically target the NHP caudate and putamen to deliver both mutant HTT (disease modeling) as well as gene therapeutics (AAV-RNAi) including 1) a pre-surgical magnetic resonance imaging (MRI) scan to determine surgical coordinates, 2) the stereotaxic surgical injection procedure including anesthesia monitoring and pain medication delivery, 3) post-surgical necropsy including brain, blood and peripheral tissue collection and 4) immunohistochemical processing of brain sections to evaluate the expression and spread of the injected agent.

2. Materials

2.1 Pre-surgical magnetic resonance imaging (MRI)

1. Personal protective equipment including scrubs, a surgical face mask, protective eye wear, nitrile gloves, a hair net, shoe covers and a water-resistant gown

2. MRI machine (1.5T or 3T) and associated imaging software

3. Ketamine HCl

4. Glycopyrrolate

5. Isoflurane anesthesia machine on a mobile, wheeled cart or transport board

6. Shaver

7. Endotracheal tube (4.0 to 5.5 mm i.d.) and umbilical tape

8. Ophthalmic ointment

9. 22G cephalic vein catheter

10. Pulse oximeter

11. Heart rate monitor

12. Animal heating mechanisms (towels, water blanket, etc)

13. MRI-compatible non-human primate head frame containing ear bars, eye bars and a palate bar

14. Mineral oil to fill the ear bars

15. Allen wrenches of various sizes

16. Fiducial marker (Vitamin E capsule)

17. Surgical coordinate sheets

18. MRI surface/head coil

2.2 Stereotaxic injection into the caudate and putamen

1. PPE including scrubs, hair bonnets, water resistant gown, surgical masks, protective eyewear and gloves (surgical staff) as well as sterile gowns and sterile surgical gloves (surgeons)

2. Anesthesia cart and physiologic monitoring system

3. Ethylene oxide sterilizing system

4. Infusate (viral vector, cells, etc)

5. Bucket of ice or cooling block if infusate must remain cold prior to injection

6. Primate head frame

7. Sterile micromanipulator

8. Sterile Allen wrenches

9. Sterile T-square (PolySquare™)

10. Sterile Hamilton infusion syringes (10–100 ul or appropriate volume for infusion)

11. Sterile Hamilton surgical needles (22–25G, 25–38mm in length, blunt tip style)

12. Sterile stereotaxic infusion pump and controller box

13. Sterile AP Zeroing bar

14. Sterile towels and a sterile craniotomy drape

15. ChloraPrep preoperative skin preparation

16. Hydromorphone (2 mg/ml)

17. Cafazolin (250 mg/ml)

18. Buprenorphine (0.3 mg/ml)

19. Lidocaine (1%) and epinephrine

20. Bupivicaine (0.5%)

21. 1 and 3 cc syringes with 25G needles

22. Lactated Ringer’s solution

23. Warm forced air system (Bair hugger) and warm water bags

24. Sterile Parafilm (several 2”x2” squares)

25. Sterile surgical drill and drill bit (2mm round, carbide burr)

26. Sterile gauze

27. Sterile surgical cotton swabs

28. Sterile pipettors and appropriately sized sterile pipette tips

29. Sterile surgical instruments (Adson forceps, scalpels, scalpel holders, hemostats, scissors, micro-dissection scissors, rongeurs, needle holders, elevators, Gelpi retractors

30. Sterile 22G needle

31. Sterile Sharpie marker

32. Sterile saline

33. Sterile 30ml syringe

34. Sterile suction tip and suction machine

35. Sterile Gelfoam

36. Sterile 4-0 Monocryl and 3-0 Vicryl suture

37. Biohazard waste container

38. Calculator, pencils and surgical coordinate sheets

2.3 Necropsy and tissue collection

1. PPE: Scrubs, water resistant gown, hair bonnet, nitrile gloves, facemask, protective eye wear

2. Peristaltic perfusion pump

3. Polyethylene tubing for perfusion pump (inside diameter: ¼ inch) attached to a 13G cannula

4. Board & masking tape for securing animal

5. Scalpel and scalpel blades, Metzenbaum scissors, forceps, assorted sizes of hemostats, heavy duty scissors, bone lever

6. Bottles of 70% EtOH and saline

7. Cutting boards

8. Weigh boats for organs

9. 60cc syringe with 18 gauge needle for drawing terminal serum

10. Supply of scalpel blades, #22, blunt & sharp

11. Supply of 4×4 gauze

12. Pruning shears

13. T-bar

14. Bone cutting forceps

15. Pens (sharpie)/pencils for labeling tubes and foil packs

16. 2 4ml serum blood tubes

17. Assorted syringes (1cc, 3cc, 5cc, 20cc, 30cc, 60cc) 7 needles (18–25G, 1” to 1.5”)

18. Tourniquet

19. Sharps containers

20. Waste disposal bags

21. Hand saw

22. Sodium pentobarbital

23. Ketamine hydrochloride

24. 0.9% sterile saline

25. Paraformaldehyde

26. Sodium Phosphate Monobasic Monohydrate NaH₂PO₄ (FW137.99)

27. Sodium Phosphate Dibasic Anhydrous Na₂HPO₄ (FW141.96)

28. 4% paraformaldehyde (40g Paraformaldehyde, 3.2g monobasic, 10.9g dibasic, 1L dH₂O)

29. Ice bucket with wet ice

30. Dewar with liquid nitrogen

31. Aluminum foil packs labeled with ID, date & tissue for tissue bank

32. Microcentrifuge tubes (2.0ml with screw cap & o-ring/sterilized)

33. Brain jar on ice filled with sterile saline

34. Small jars on ice filled with sterile saline

35. Tissue biopsy cores (2mm)

36. Rhesus brain matrix

37. Tissue slicer blades

38. Sterile petri dishes

2.4 Cutting and immunohistochemical processing of brain

1. PPE: White lab coat, nitrile gloves, protective eye wear

2. Sliding microtome and microtome blade

3. Paint brushes for tissue cutting

4. Dry ice

5. Trizma® pre-set crystals (pH7.4, avg Mw: 151.6)

6. NaCl (FW58.44)

7. Triton®X-100

8. Sodium Phosphate Dibasic Anhydrous Na₂HPO₄ (FW141.96)

9. Sodium Phosphate Monobasic Monohydrate NaH₂PO₄ (FW137.99)

10. Serum (goat or donkey)

11. Sodium meta-Periodate INaO₄ (FW213.89)

12. DAB (FW360.11)

13. Nickel(II) sulfate hexahydrate (FW262.85)

14. Vectastain® Elite® ABC Kit

15. Dilution media (7.46g Trizma, 8.77g NaCl, 0.5 ml TritonX-100, 1L dH₂O)

16. TBS (7.46g Trizma, 8.77g NaCl, 1L dH₂O)

17. PBS (5.47g Dibasic, 1.60 Monobasic, 9.26g NaCl, 1L dH₂O)

18. Cryoprotectant solution (300g Sucrose, 300 mls ethylene glycol, 0.2g Sozium Azide, 500 mls PBS)

19. Netted staining dishes (with glass dishes to contain fluid)

20. Orbital shaker

21. Appropriate primary and secondary antibodies

22. 6 well tissue culture plates

23. Large 24 well (4×6) compartmented tissue collection box

3. Methods

3.1 Pre-surgical magnetic resonance imaging (MRI)

1. Sedate the animal while in its home cage with 10–20 mg/kg Ketamine combined with 0.01–0.02 mg/kg Glycopyrrolate (intra-muscular-IM) and bring to the pre-operative procedure room (food withheld for 12 hours prior to MRI and surgery). Shave animal’s head from the brow ridge caudally to the foramen magnum and laterally down to the lateral canthus of each eye, place ophthalmic ointment in each eye, shave the left forearm and establish a 22G cephalic vein catheter for agent administration during surgery.

2. Place an endotracheal tube (for adult rhesus macaques, 4.0 to 5.5 mm i.d. endotracheal tubes are typically used), secure tube with umbilical tape and start the animal on 1.5% isoflurane. While waiting to go into MRI, monitor the animal’s pulse rate, blood oxygen saturation, and ETC0₂ levels.

3. Bring the animal to the MRI facility on a wheeled cart under isoflurane anesthesia.

4. In the MRI procedure room, place the animal’s head into the MRI-compatible stereotaxic head frame (Figure 1A) while still connected to the breathing tube and under anesthesia (Note 1). First place the animal in the earbars (filled with contrast dye or mineral oil) such that there is free movement of the animals head up and down (nodding motion), but no movement laterally. Center animal in the head frame both medial-laterally and dorso-ventrally. Next, place the palate bar into the animal’s mouth, resting against the hard palate, and secure into position such that the animal’s head is straight and its eyes are facing forward. Finally, secure the eye bars such that they fit tightly in the ventral grooves of the orbital socket. Ensure that ear bars, palate bar and eye bars are all secured in place.

5. Disconnect the animal’s breathing tube from the anesthesia machine and quickly transfer the animal placed in the head frame to the MRI table (still prone and head first) and reconnect to the anesthesia machine located in the MRI unit (Note 2).

6. Place a circulating warm water pad and towels under the animal on the MRI table and place extra towels over the animal order to maintain body temperature.

7. Attach peripheral physiological (pulse rate/oxygen saturation, ETCO2, respiration, and/or non-invasive blood pressure) monitors to the animal (digit cuffs) and observe for accuracy and reliability of readings. From the control room, an MRI technician should continue to monitor the animal’s respiratory rate, pulse rate, end tidal CO2 and oxygen saturation (Note 3).

8. Tape a fiducial marker to right side of the animal’s head to use as a reference on the MR image (Note 4).

9. Place the surface/head coil (Figure 1B) directly above and parallel to the animal’s head such that it is centered over the scanning area of interest (caudate and/or putamen, approximately 12–25 mm in front of the ear bars) for HD gene therapy studies. The coil should be approximately 0.5–1.0 cm above the surface of the head but not touching the skin. (Note 5). Center the animal’s head on MRI table using the laser crosshair feature on the MRI scanner and then send the animal into the center of the magnet.

10. First collect a quick, single-slice, three-axis localizer scan that gives a view of the animal’s head in the three scanner-frame axes (voxel size1.0 × 1.0 × 1.0). This ensures that the coil is functioning properly and allows for further refinement around the scanning area of interest (1–2 minute scan).

11. Next collect a T1-weighted structural scan (MPRAGE) scan to establish the surgical coordinates (voxel size: 0.5 × 0.5 × .05, 12 minute scan, Note 6). Coordinates are determined from coronal scans using the MRI software Distance Tool as follows: 1) Anterior/Posterior (AP) determined by establishing the MR image containing your injection region of interest (ie caudate or putamen) and calculating the distance (in millimeters (mm)) in front of or behind the MR image containing the earbars (earbar contrast dye is evident on scan), 2) Medial/Lateral (ML) established by determining the distance (mm) from the center of the two hemispheres to the proposed injection site (draw a vertical line through the center of the sagittal sinus vein as a reference) and 3) Dorsal/Ventral (DV) established by determining the distance from the pial surface to the proposed injection site (Figure 2, red asterisks indicates the sagittal sinus vein and the blue dot indicates the injection site in the rhesus putamen, Note 7). Record surgical coordinates on a piece of paper that will go with you into the operating room.

12. When the scan is finished, the animal can be removed from the magnet, disconnected from physiological monitors and anesthesia tubing, placed back onto the mobile anesthesia cart, reconnected to anesthesia on cart and wheeled into the operating room.

Figure 1.

Pre-surgical MRI to determine surgical coordinates. (A) Animals are anesthetized and fitted into an MRI-compatible stereotaxic head frame by first securing the animal into the earbars, followed by the palate and eye bars. (B) The scan is obtained using a surface coil that sits directly above the animal’s head.

Figure 2.

Determining surgical coordinates from the MRI scan. Surgical coordinates an for injection site into the putamen (shown by a blue dot) are created by using the MRI software to establish 1) the distance in mm in front of the earbars (AP coordinate), 2) the distance in mm lateral to the center of the sagittal sinus vein (ML coordinate, shown by a red asterisks) and 3) the distance in mm ventral to the pial surface of the brain (DV coordinate).

3.2 Stereotaxic injection into the caudate and putamen

1. Disconnect animal from anesthesia and remove from mobile anesthesia cart. Place animal onto the surgical table and connect endotracheal tube to surgical anesthesia unit. Maintain animal on 1–2% Isoflurane combined with 100% oxygen administered at a rate of 1–1.5 L/min (Note 8). Secure the base of stereotaxic frame to the table with surgical tape (animal’s torso and legs must be delicately lifted up to achieve this, while head remains locked into frame) and place physiologic monitoring equipment on the animal.

2. Administer pre-operative doses of Hydromorphone HCl (2 mg/ml) (range of doses: <3kgs bodyweight, 0.5mgs, 0.25ml; 3–10kgs bodyweight, 1mg, 0.5ml; >10kgs bodyweight, 2mg, 1ml) intravenously and 25 mg/kg Cefazolin intravenously as well as every 2 hours intra-operatively (for both drugs).

3. Establish intravenous fluids (Lactated Ringers solution) @ 10–20 ml/kg/hr. Fluids may be increased based on hydration status of patient or due to blood loss during surgery.

4. Apply a ChlorPrep (a chlorhexadine/alcohol instant solution) to the entire surgical site and allow it to dry. Drape surgical areas with sterile towels.

5. Place a warm forced air system (Bair hugger) around the animal, as well as warm water bags, for the duration of the surgical procedure. Place a full table drape over the animal, towels and Bair hugger to avoid any potential breaks in aseptic technique due to the necessary manipulation of the stereotaxic manipulator.

6. Surgical staff should wear hair bonnets, surgical masks, protective eyewear and gloves prior to entering the operating room. Surgeons should perform a full pre-surgical sterile scrub with a chlorhexadine or betadine solution. Surgeons should wear sterile gowns and sterile surgical gloves prior to draping the patient and initiating the surgical procedure.

7. Open sterile (ethylene oxide sterilized) surgical packs containing instruments, drill, micromanipulator, AP zeroing bar, surgical pump, needles/syringes and supplies on a flat surface near the operating table.

8. Assemble needles and syringes and wet the syringe barrels by withdrawing and expelling injection vehicle (saline, PBS, buffer, etc) several times (Note 9).

9. Slide the micromanipulator onto the AP zeroing bar and tighten down anywhere.

10. Screw the AP slide attachment onto the micromanipulator and tighten down (use Allen wrench if necessary).

11. Screw the infusion pump onto the AP slide attachment and set/lock AP slide attachment to a set number (example- 50 or 0; all AP coordinate measurements from the MRI will be added or subtracted from this number, depending on whether the micromanipulator is placed on the left or right AP bar on the animal’s head frame).

12. Place the syringe/needle into the infusion pump and lock into place. Use a T-square (ex PolySquare) to make certain that the micromanipulator, pump, syringe and needle are straight and make adjustments if not (Figure 3).

13. Ensure that all components of the micromanipulator are tightened appropriately (use Allen wrenches).

14. Carefully loosen the micromanipulator from the AP zeroing bar and slide laterally to line up the tip of the needle with the point structure on the AP zeroing plate. Record the number on the AP zeroing bar (not the AP slide attachment). When the micromanipulator is placed at the same number on the animal’s head frame AP bar, the needle will be correctly positioned at the ear bars (ear bar zero).

15. Administer 25 mg/kg of Cefazolin intravenously prior to making the initial scalp incision for antibiotic coverage. Inject an intradermal, local block at the proposed incision site consisting of 0.4 ml (up to 0.8 ml depending on size of incision) Bupivicaine (0.5%) combined with 0.1 ml (up to 0.2 ml depending on size of incision) and Lidocaine (1%) with epinephrine.

16. Make a linear sagittal incision of approximately 6 cm length across the top of the animal’s scalp. Elevate the subcutis and periosteum and retract laterally to expose the skull. Sharply dissect muscle attachments and retract laterally.

17. Drill a 1 cm long line close to the planned injection site (caudate and/or putamen) perpendicular to the sagittal sinus. Extend the drill line down to the level of the dura to expose the dark sagittal sinus vein. The exposed sagittal sinus vein will be used to establish the ML zero point.

18. Bring the micromanipulator (connected to the pump, syringe and needle) from the AP zero bar to the animal’s head frame and lock into place on either the left or right AP bar at the number recorded earlier (Note 10). Use the AP slide attachement and the ML dials on the micromanipulator to bring the needle to the center of the saggital sinus vein and record the ML number as the zero point.

19. Calculate the AP and ML coordinates for each injection site (Note 11) and use the micromanipulator to bring the needle to those sites and mark the sites on the skull with a pencil or permanent marker (Sharpie).

20. Create oval craniotomies over the proposed injection sites using a high speed surgical drill with a 2 mm carbide burr (Figure 4A). The length and diameter of the craniotomies will vary depending on the number of proposed injection sites in each hemisphere. Use saline and suction to cool the drill bit and clear bone debris from the craniotomy area. Smooth the edges of the craniotomies with ronguers and incise the dura with a 22 gauge needle followed by micro-dissection scissors (Figure 4B).

21. Load the syringe with infusate without removing the syringe from the infusion pump (Note 12) and prime the needle by using the infusion pump to infuse until a small amount of liquid is visible at the tip of the needle. Swab the infusate away with gauze or a surgical swab.

22. At each injection site, carefully lower the needle down to the pial surface and record the DV value on the micromanipulator, which will serve as the DV zero value. Calculate the DV coordinate by subtracting the DV value obtained from the MRI from the DV zero value.

23. Slowly lower the needle to the DV coordinate and infuse at the desired rate (Figure 4C, Note 13).

24. After the infusion is complete, allow the needle to rest in place for an additional 3–10 minutes to allow the infusate to disperse from the needle tip. Slowly raise the needle out of the brain and proceed to the next injection site until all injections have been made.

25. Place Gelfoam sponge into the craniotomy sites (Figure 4D) and use Cruciate 3-0 Vicryl sutures to bring the muscle back to its original lateral position along the skull. Close the subcutis and appose the skin edges with simple interrupted and intradermal 4-0 Monocryl sutures. Take the animal out of the stereotaxic head frame, discontinue the isoflurane gas anesthesia while continuing oxygen support and then release the animal from the stereotaxic head frame. Allow the animal to recover on the OR table until extubation.

26. Administer the following post-surgical medications: Hydromorphone on the day of surgery every 4 hours IM until 2000 hours, Buprenorphine for 2 days IM SID at 2000 hours and Cefazolin for 3 days IM BID (25mg/kg, 250mg/ml) (Table 1).

Figure 3.

Zeroing the AP slide bar. Prior to securing the micromanipulator on the animal’s head frame in the surgery room, the AP slide bar must be zeroed out by setting the AP slide bar to a pre-determined number (example, 50) and lining up the tip of the needle/cannula with the point structure on the AP zeroing plate. When the micromanipulator is placed into the exact position on the AP bar of the head frame, the needle tip will be located at the earbars. Using a T-square when zeroing the AP slide bar helps to ensure that the needle is straight.

Figure 4.

Stereotaxic injection into the caudate and putamen. (A) A craniotomy that encompasses the proposed injection site(s) is established using a high speed drill and a 2 mm carbide burr. The edges are rounded using a rongeur. (B) The dura is excised using a micro-scissor and folded back to expose the pial surface of the brain. (C) The needle is slowly lowered to the calculated injection site coordinate and the infusate delivered using an infusion pump set to either a constant or ramping injection rate. (D) Post-injection, the needle is slowly raised and the craniotomy is filled with Gelfoam prior to suturing.

Table 1.

Pain medications administered post-surgery. Different doses of Hydromorphone and Buprenorphine are listed according to rhesus macaque weight. Hydromorphone is administered post-surgery every 4 hours IM until 2000 hours and Buprenorphine (IM) for 2 days post-surgery at 2000 hours (once per day).

Animal Weight	Hydromorphone (2mg/ml)	Buprenorphine (0.3mg/ml)
<3kg	0.5mg (0.25ml)	0.15mg (0.5ml)
3–10kg	1mg (0.5ml)	0.3mg (1ml)
>10kg	2mg (1ml)	0.3mg (1ml)

3.3 Necropsy and tissue collection

Necropsy

• 1Sedate the animal while in its home cage with 10–20 mg/kg ketamine (IM) and bring to the necropsy room. Secure animal to the board on the necropsy table with masking tape and anesthetize animal with sodium pentobarbital administered IV at 25mg/kg. Assess depth of anesthesia by loss of palpebral, corneal, pain and gag reflexes
• 2Incise abdomen with scalpel after adequate anesthesia has been established and collect terminal blood samples from abdominal aorta or caudal vena cava if needed. Place blood in red top serum tubes. Sever aorta to effect exsanguination and euthanasia.
• 3If necessary, collect cerebrospinal fluid (3cc syringe with 1.5” 22 gauge sterile needle) and place into sterile Microcentrifuge collection tubes.
• 4Open the thoracic cavity and the perfuse brain and spinal cord with 1–2 liters of ice-cold, 0.9% sterile saline via the right carotid artery (Note 14).
• 5Carefully saw around the entire skull in the axial plane using a hand saw and remove the skull cap using a prying motion with a bone lever. Gently remove the brain from the calvarium and place it in a brain jar (in saline) on ice. It will be necessary to dissect through the cranial nerves and spinal cord prior to removing the brain.
• 6Collect relevant samples of tissues/organs (liver, spleen, gastroc, kidney, lung, lymph nodes, pancreas, intestine, adrenal, gonads) in microcentifuge tubes (smaller samples) and foil packs (larger samples for banking). Immediately place the samples in microfuge tubes on dry ice and the samples in foil packs in the Dewar filled with liquid nitrogen (Note 15).
• 7If of interest, remove spinal cord, divide into 3 sections (Cervical, Thoracic & Lumbar), remove meninges, cut a 2 cm long segment from the middle of each section and place each in small jars with sterile saline on ice.
• 8Transport the brain & spinal cord on ice back to the laboratory in an enclosed biohazard container. Transport the Dewar full of frozen tissue samples back to laboratory. Transport blood samples in red-top tubes back to the laboratory at room temperature in a biohazard container.
• 9Place the brain into a non-human primate brain matrix and carefully cut into slabs (2–8mm thick slabs recommended) using tissue blades (Figure 5A, showing a rhesus macaque brain matrix). Place each brain slab into a separate, sterile saline-filled petri dish on ice.
• 10Use tissue biopsy cores (2mm) to take samples from regions of interest for future molecular and/or biochemical analyses (Figure 5B, Note 16). Place samples in microcentrifuge tubes and place immediately on dry ice. Store samples at −80 degrees.
• 11After collecting brain samples, place slabs of brain tissue in 4% paraformaldehyde for 48 hours for post-fixing. After post-fixing in 4% paraformaldehyde, place slabs of brain tissue in 30% sucrose until they have completely sunken to the bottom of the jar.
• 24Cut brain slabs using a frozen microtome at a thickness of 40 um and collect tissues in large 24 well (4×6) compartmented tissue collection box filled with cryoprotectant solution.

Figure 5.

Brain processing post-necropsy. (A) Following removal at necropsy, the brain is placed in to a matrix and sectioned into slabs of varying thickness using tissue slicer blades. (B) The slabs are placed into petri dishes containing sterile saline and biopsy punches are taken from key areas to be used for molecular and biochemical studies. Brain slabs are then post-fixed in 4% paraformaldehyde for 48 hours and cryoprotected in 30% sucrose prior to immunohistochemical staining.

Blood processing

1. Allow blood to coagulate for 2 hours at room temperature.

2. Centrifuge red top serum tubes at 2500 rpm for 20 minutes at room temperature (25°C) using a tabletop centrifuge.

3. Pipet off serum and transfer to microcentrifuge tubes. Store serum samples at −80 degrees.

3.4 Immunohistochemical processing of brain to verify expression of gene therapy agent

Day 1

1. Wash tissue in dilution media in netted staining dishes (5 × 8 minutes).

2. Block endogenous peroxidase (2.13 g of sodium meta-periodate per 100 mls TBS) for 20 minutes in netted staining dishes.

3. Wash tissue in dilution media in netted staining dishes (5 × 8 minutes).

4. Block tissue in 5% of the appropriate serum (goat, donkey) in netted staining dishes (5 mls serum per 100 mls dilution media).

5. Incubate tissue in primary antibody solution (primary antibody solution: 3 mls serum and 400 ul triton-X per 100 mls PBS in 6 well tissue culture plates (either shaking at room temp overnight or at 4 degrees for 48 hours). Each primary antibody should be used at its own concentration.

Day 2

• 1Wash tissue in dilution media in netted staining dishes (5 × 8 minutes).
• 2Incubate tissue in secondary antibody solution in 6 well tissue culture plates (shaking at room temperature for 1 hour). Secondary antibody solution: 3 mls normal serum per 100 mls dilution media. Each secondary ab should be used at its own required concentration.
• 3Wash tissue in dilution media in netted staining dishes (5 × 8 minutes).
• 4Incubate tissue in ABC solution in netted staining dishes for 1 hour. (First add 3 mls serum per 100 mls of dilution media. Next add 4 drops of bottle A and 4 drops of bottle B to 10 mls of serum/dilution media solution and mix. After waiting 30 minutes, mix those 10 mls back with the other 90 mls and place tissue into the solution).
• 5Wash tissue in TBS in netted staining dishes (3 × 8 minutes).
• 6DAB reaction to develop color on the tissue: First add 50 mg DAB to 100 mls of TBS. Next add 2.5 g Nickel II Sulfate if nickel intensification needed (makes stain purple versus brown). Immediately prior to tissue incubation, add 20 ul of the 30% hydrogen peroxide solution. Incubate tissue in DAB solution until the stain develops (depends on the antibody, ranges from ∼ 1–10 minutes) (Note 17, Figure 6).
• 6Wash tissue in TBS in netted staining dishes (3 × 8 minutes).
• 7Store tissue at 4 degrees in PBS in netted staining dishes until able to mount onto slides.

Figure 6.

Immunohistochemcal staining of brain sections to identify the injected agent. Anti-eGFP staining demonstrates accurate expression of the transgene following a 50 ul injection of AAV2/1-eGFP (5e12 vg/ml) into the putamen of an adult rhesus macaque.