Using Optogenetics to Reverse Neuroplasticity and Inhibit Cocaine Seeking in Rats

The experiments described in this protocol were consistent with the guidelines set forth by the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the University of Pittsburgh's Institutional Animal Care and Use Committee. All procedures were performed using adult, naïve Sprague-Dawley rats that weighed 275-325 g upon arrival.

1. Construction of Optic Fiber Implants and Patch Cables

1. Prepare optic fiber implants following previously published protocols⁹. Experiments described in this protocol used 200 µm core fiber (0.5 NA) and Ø1.25 mm Multimode LC/PC Ceramic ferrule, Ø230 µm hole size.
   1. Use a Dremel tool to score the lower third of a ferrule (closest to the flat end of the ferrule). Scoring the ferrules helps them stay attached to dental cement, increasing the likelihood that they will remain secure throughout the entire extent of experimentation.
   2. Use wire cutters to cut ~35 mm of fiber. Use fiber stripping tool to strip ~25 mm of fiber, leaving 10 mm unexposed.
   3. Prepare heat-curable epoxy according to manufacturer's instructions. Dissolve 1 g of resin powder in 100 mg of hardener compound. Attach a blunted 25-gauge needle to a 1 mL syringe. Fill the syringe with epoxy and attach a blunted 25-gauge needle tip.
   4. Use a vice or clamp to securely hold the ferrule with the flat side facing up and the convex side facing down. With the epoxy-filled syringe, add one drop of epoxy to the flat side of the ferrule, using caution to wipe excess epoxy from the sides of the ferrule.
   5. Insert the stripped portion of fiber through the ferrule allowing an extra 5 mm of stripped fiber exposed. In the case of LA implants, the fiber will be implanted 7.9 mm ventral to bregma, so the exposed length of unstripped fiber should be ~13 mm.
   6. Cure the epoxy with a heat gun for about 30-40 s until it turns black/amber in color.
   7. Score the fiber directly at the interface of the convex end of the ferrule with diamond knife and use a finger to gently tap the fiber off.
   8. Polish the convex end of the ferrule by holding with a hemostat, being sure to apply even pressure, and making 20 circular rotations each on a series of polishing paper (from high grade to low grade; 5, 3, 1, 0.3 µm).
   9. Secure unstripped fiber to table using tape and score stripped fiber, leaving an extra 2 mm beyond the ventral coordinate (For LA implants, final length of fiber is ~10 mm). Use a hemostat to pull on the ferrule and evenly break the fiber where it has been scored. Be careful not to cut fiber completely when scoring, or else the core of the fiber will be damaged.
2. Build patch cables that are compatible with optic fiber implants. Custom designed patch cables were purchased (See Table of Materials). Alternatively, patch cables can be constructed following previously published protocols⁹.
   NOTE: The diameter and NA of the ferrule fiber and patch cable fiber must match at the coupling junction to prevent excess loss of light, which can result in a failure to sufficiently stimulate neural activity.
3. Measure the light output through the patch cable and optic fiber implants by attaching patch cable/optic fiber to an appropriate laser light source (473 nm, 1 mW output) and measuring output with a light sensor. A successfully constructed fiber will emit a concentric circle of light and have no more than 30% light loss.

2. Rodent Intravenous Catheterization, Virus Delivery, and Optic Fiber Implantation

1. Prepare animal for surgery.
   1. Fully anesthetize rats with anesthetic of choice based on institutional guidelines. One option is ketamine hydrochloride (87.5-100 mg/kg, i.m.) and xylazine hydrochloride (5 mg/kg, i.m.). Ensure that the rat is fully anesthetized by checking for lack of a toe pinch reflex.
      CAUTION: Ketamine is a controlled substance that must be handled according to institutional guidelines. 
      NOTE: Intramuscular injection of anesthetics is used in this study as it produced a more rapid and reliable anesthesia induction than intraperitoneal injection. Continuously monitor the rat's respiration and responsiveness and provide thermal support throughout the surgery. 
   2. Shave a large area of the rat's back (upper back from just above the shoulder blades to the middle of the back) as well as the area of the neck underneath the right forelimb, and the scalp.
   3. Place the rat in the surgical area and apply puralube (artificial tears) to the eyes. Inject a body weight volume of carprofen (analgesic) subcutaneously (s.c.) through the skin of the upper back, then inject 5 mL of Lactated Ringer's solution s.c. through the skin of the lower back.
   4. Sanitize all surgical sites by wetting a piece of sterile gauze with betadine and wiping it down the shaved surgical area using a circular motion. Then repeat the process with 70% ethanol. Repeat this alternating cycle three times. 
2. Perform intravenous catheter implantation according to previously published protocols^4,10.
   NOTE: A surgical drape is not used during this surgery to avoid irritation during catheter implantation. Sterilize all instruments and equipment before use. Use sterile gloves and change the gloves if any non-sterile surface is contacted.
3. Immediately following catheter implantations, secure the rat in a stereotaxic frame to perform AAV injections.
   1. Deliver a subcutaneous (s.c) injection of 2% lidocaine (0.2-0.3 mL) to the scalp as a local anesthetic.
      NOTE: Local anesthetic is not used during the intravenous catheter implantation to avoid alterations to the surgical outcomes. 
   2. Connect a 26-gauge stainless steel injection cannula to a Hamilton syringe filled with 1 µL of concentrated AAV solution: either AAV5-hSyn-tdTomato or AAV5-hSyn-oChIEF-tdTomato
      NOTE: oChIEF is a variant of the blue-light sensitive opsin channelrhodopsin (ChR2), that can respond to a wide range of frequencies^8,11, and therefore has utility for the low-frequency LTD experiments discussed in this protocol, but also for high-frequency LTP experiments (not discussed here). The oChIEF construct was donated by Dr. Roger Tsien and processed for packaging and purification by the Duke Viral Vector Core. At least 3-4 weeks is needed between the injection day and the day of LTD induction to allow for optimal virus expression in MGN axon terminals.
      CAUTION: Generally, AAV is regarded as a Biosafety Level 1 (BSL-1) organism, with low risk of self-infection unless a helper virus is used in its production. Its use requires IACUC approval and proper PPE must be used at all times in accordance with institutional guidelines to limit unnecessary exposure.
   3. Using a scalpel, make a 0.5 mm incision from the front to the rear of the skull, and remove overlying tissue to expose the surface of the skull.
   4. Level the rat's head in the anterior/posterior axis and zero stereotaxic coordinates to bregma.
   5. Drill three small holes through the skull using a Dremel tool equipped with a small drill bit. Use screwdriver to firmly mount stainless steel screws (M2x4 965-A2) in place.
      NOTE: Screws are necessary for proper binding of dental cement and creation of sturdy, long-lasting headcaps. Position of screws should be spread across the anterior-posterior axis of the skull, and away from AAV injection site.
   6. Drill bilateral holes for injection of AAV based on the coordinates from the Rat Brain Atlas (Watson and Paxinos)¹² for the medial portion of the medial geniculate nucleus (MGN); in mm from bregma, AP: -5.4; ML: ±3.0; DV: -6.6. Slowly lower injection cannulae (4 mm/min) until positioned in the MGN. Inject concentrated AAV solution at a rate of 0.1 µL/min.
   7. Leave injection cannula in place for 5 min after infusions are complete to allow for diffusion away from the cannula and then slowly withdraw cannula from the brain.
4. Immediately following virus injections, continue to implant optic fibers^4,9 targeting MGN-LA terminals.
   1. Use a Dremel tool to drill bilateral holes for optic fiber implants targeting the lateral amygdala (in mm from bregma, AP: -3.0; ML ±5.1).
   2. Use forceps to grab the ferrule of the optic fiber implant and fix it to the stereotaxic adapters so that they are securely held in place.
   3. Slowly lower fibers at a rate of 2 mm/min, until the tip of the fiber sits in the dorsal portion of the LA (DV: -7.9 mm).
   4. Secure ferrules to the skull first using a thin layer of Loctite instant adhesive followed by dental cement and cover ferrules with 1.25 mm diameter ferrule sleeves and dust covers.
      NOTE: The choice of adhesive for securing the ferrules to the skull should be approved by the local or institutional animal care and use committee. Loctite is used for this study to reliably secure the ferrules to the skull after trial and error with multiple adhesives; however, available alternatives can be considered. 
5. Following surgical procedures, house rats individually, and provide free access to food and water. Provide postoperative care consistent with institutional guidelines. Flush catheters daily with saline containing gentamicin (5 mg/mL) and heparin (30 USP/mL) to maintain patency. At least 5 days post-surgery and 24 h prior to the start of behavioral experiments, food restrict rats to ~90% of their free-feeding weight.

3. Rodent Cocaine Self-Administration and Instrumental Lever Extinction

NOTE: All behavioral procedures are conducted in standard operant conditioning chambers, equipped with two retractable levers on one wall, a stimulus light above each lever, a tone generator, a house light, and an infusion pump.

1. Subject rats to daily 1-h cocaine (2 mg/mL) self-administration training sessions under an FR1 schedule of reinforcement.
   1. Place rats in operant chamber each day and allow rats to lever press. A press on the designated 'active lever' (counterbalanced across left and right levers) results in a cocaine infusion (1.0 mg/kg/infusion) and a 10 s presentation of a compound light and tone cue. A press on the designated 'inactive lever' has no programmed effects.
   2. Continue self-administration experiments for at least 10 d and until rats successfully earn at least 8 infusions/day across 3 consecutive days. Failure to reach acquisition criteria by day 20 results in exclusion from the study.
2. After acquisition criteria are successfully met, subject rats to 1 h instrumental extinction sessions for 6-10 d.
   1. Place rats in operant chambers and allow rats to freely lever press. However, responses on both the active and inactive levers have no programmed consequences.
   2. Have rats continue instrumental extinction daily until an average of < 25 lever presses over two consecutive days occurs.

4. In vivo Optogenetic Induction of LTD

NOTE: Optogenetic inhibition experiments take place 24 h after the final day of instrumental extinction.

1. Connect patch cords to a 473-nm blue laser diode via a rotary joint suspended above a clean, standard rodent housing cage with the cover removed. This setup allows rodents to freely move around the cage during the optogenetic stimulation.
2. Turn on laser diode according to operating instructions and connect to a pulse generator. Adjust settings, so that when turned on the rat will receive 900 2-ms pulses of light at 1 Hz.
   CAUTION: Proper eye protection must be used at all times while operating laser.
3. Measure the light intensity through the patch cord using a light sensor. Adjust the intensity of the laser so that light output through the patch cable is ~5-7 mW.
4. Place rats in a clean housing cage. Remove dust covers and ferrule sleeves, exposing the ferrules. Connect patch cords bilaterally to the optic fiber implants. Allow rats to explore the environment for 3 min prior to LTD induction.
5. Turn on the pulse generator to initiate optogenetic stimulation.
   NOTE: Although unlikely, if rat experiences any adverse reactions to stimulation, the experiment is immediately terminated, and rats are properly euthanized based on institutional guidelines.
6. Following LTD induction, keep rats in the cage for 3 min, and then place back in their home cages.
7. For control experiments, use the same stimulation procedure on rats that express the AAV5-tdTomato control virus. For sham experiments, attach a patch cord to the optic fiber of rats that express the AAV5-oChIEF virus, but no stimulation is delivered during a 15 min session.

5. Test the Effect of Optogenetic Stimulation on Cue-Induced Cocaine Seeking

1. 24 h after in vivo optogenetic stimulations, place rats back in operant conditioning chambers. Rats are subjected to a 1-h standard cue-induced reinstatement session to assess cocaine seeking behavior.
   NOTE: During cue-induced reinstatement, a response on the active lever yields a 10-s presentation of the cocaine-associated cue, but no cocaine infusions.
2. Give a second reinstatement test at least 1 week after the first test to determine if optogenetic LTD induction results in a long-term suppression of cocaine seeking

6. Staining, Fluorescence, and Imaging for Histological Verification of Viral Expression and Optic Fiber Placement

1. Make 1x phosphate buffered saline (PBS) and 4% paraformaldehyde (PFA). Store both solutions on ice. Total volume will depend on the number of rats in the study (~100 mL of PBS and 200 mL of PFA will be used per rat).
   CAUTION: PFA is a toxic chemical and known carcinogen. Take proper care to avoid inhalation as well as contact with the skin. Its use and disposal should be in accordance with institutional guidelines, including the use of proper PPE and a chemical flow hood.
2. Setup peristaltic pump at a flow rate of 20 mL/min. Fill tubing of pump with 1x PBS. Attach a blunted 20-gauge needle to the end of the tubing.
3. Deeply anesthetize rats with sodium pentobarbital (100 mg/kg, i.p.). Confirm the depth of anesthesia by lack of response to toe pinch before proceeding further.
   NOTE: Sodium pentobarbital is used since the perfusion is a terminal procedure.
4. Use surgical scissors to cut open the abdominal cavity of the rat below the diaphragm. Cut through the rib cage rostrally along the lateral edges to expose the rat's heart. Use hemostat to clamp the rostral portion of the rib cage away from the heart. Cut away any overlying fat tissue that surrounds the heart.
5. Insert the blunted needle through the left ventricle and up into the aorta. Cut a small hole in the right atrium to drain solution as it returns to the heart.
6. Perfuse each rat with 1x PBS for 5 min followed by 4% PFA, pH 7.4 for 10 min.
7. Decapitate the rat, extract the brain, and postfix it in 4% PFA for 24 h. Then transfer the brain to 30% sucrose solution for 2-3 d.
8. Section brains at 50 µm using a cryostat.
9. Mount all slices containing the LA or MGN onto glass slides and coverslip.
10. Image slices using an epifluorescent microscope to verify AAV-oChIEF-tdTomato expression in the MGN and its projections to the LA, as well as placement of the optic fiber above the LA.

7. Perfusion and Acute Brain Slice Preparation for Electrophysiology Experiments

NOTE: Electrophysiological experiments are performed on a subset of animals to validate the success of in vivo LTD.

1. Prepare electrophysiology solutions using the reagents listed in Tables 1-3^4,13. Adjust the pH of all solutions to 7.4 with HCl and adjust osmolarity to 300-310 mOsm/kg H₂O. Make solutions fresh prior to experiments and store at 4 °C for up to 1 week. Saturate all solutions with carbogen (95% O₂/5% CO₂) at all times during use.
2. Using isoflurane according to the local or institutional animal care and use guidelines, deeply anesthetize the rat in an enclosed euthanasia chamber.  Confirm that the animal is fully anesthetized via the toe-pinch reflex.
3. Fill a small beaker with 50 mL of ice-cold cutting solution. Fill tubing of peristaltic pump with solution and adjust flow rate to 20 mL/min. Attach a blunted 20-gauge needle to the end of the tubing.
4. Open the abdominal cavity (See steps 6.4 and 6.5) and briefly perfuse rat with cutting solution (maximum 1-2 min).
5. Following perfusion, immediately decapitate rat. Remove the brain and place it in a small beaker filled with 4 °C cutting solution for 30 s-1 min.
6. Transfer brains with a spatula and quickly fix them to the chamber of a vibratome. Remove the pia using fine forceps. Fill the chamber with 4 °C cutting solution and prepare acute coronal slices (250 µm thick) of the amygdala at a velocity of 0.37 mm/sec and a frequency of 70 Hz.
7. As slices are obtained, place each one in a holding chamber filled with cutting solution and incubate at 37 °C for 10-12 min. About 5-7 slices containing the LA can be obtained per animal.
8. Transfer slices to a beaker of room temperature (RT) holding solution and allow to recover for >30 min prior to experimentation.
   ​NOTE: Slices generally remain healthy while kept in holding solution for 4-6 h. Due to the fluorescent nature of the AAV, slices are kept in low-light conditions.

8. Ex vivo Electrophysiological Recordings

1. Prepare intracellular solutions using the reagents listed in Tables 4-5.
   NOTE: Intracellular solutions should be made in advance of experiments and can be stored long-term (3-12 months) at -80 °C or short-term (1-2 months) at -20 °C. Solutions are pH adjusted to 7.3 (with CsOH for Cs-based intracellular solution and with KOH for K-based intracellular solution). Adjust to a final osmolarity of 290-300 mOsm/kg H₂O.
2. Prepare 500 mM picrotoxin stock solution dissolved in dimethylsulfoxide (DMSO).
   NOTE: Picrotoxin stocks are aliquoted and stored at -20 °C. The day of use, aliquots are thawed and added to recording solution to a final concentration of 100 µM.
   CAUTION: Picrotoxin is a non-competitive antagonist of GABA_A receptors, so infusion of picrotoxin has a stimulative effect. It is severely toxic by oral ingestion or skin absorption. Proper PPE must be used at all times when working with picrotoxin.
3. Transfer slices to an upright microscope designed for electrophysiology experiments.
4. During experiments, continuously bath perfuse slices with recording solution that is heated to 31-33 °C.
5. Magnify the LA using a 4x objective. Identify principal neurons by morphology with a 40x water immersion lens.
6. Use a glass pipette (3-5 MΩ) filled with either a Cs-based intracellular solution (for voltage clamp experiments) or a K-based intracellular solution (for current clamp experiments) to obtain whole-cell patch clamp recordings.
7. Identify AAV-infected MGN axonal projections under fluorescence (using an RFP filter). Stimulate projections using a blue light (473 nm) DPSS laser connected to a pulse generator.
   CAUTION: To limit laser exposure, collimated laser light is coupled to a fluorescent port on the microscope and focused onto the slice through the objective.
   NOTE: In voltage clamp mode, excitatory postsynaptic currents (EPSCs) are optically-evoked at 0.1 Hz. Neurons receiving inputs from AAV-infected MGN neurons will exhibit reliable EPSCs.
8. To induce ex vivo LTD, in current clamp mode, record a stable baseline of excitatory postsynaptic potentials (EPSPs) for at least 10 min. Next, deliver 900 2-ms pulses of 473-nm light at a frequency of 1 Hz (total time = 15 min). Then continuously record EPSPs at 0.1 Hz for ≥60 min.