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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Rodent Intravenous Catheterization, Virus Delivery, and Optic Fiber Implantation
- Prepare animals for surgery.
- Fully anesthetize rats with the anesthetic of choice based on institutional guidelines. One option is ketamine hydrochloride (87.5-100 mg/kg, intramuscular [i.m.]) and xylazine hydrochloride (5 mg/kg, i.m.). Ensure that the rat is fully anesthetized by checking for the lack of a toe pinch reflex.
CAUTION: Ketamine is a controlled substance that must be handled according to institutional guidelines.
NOTE: Intramuscular injections of anesthetics are used in this study as they produce 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. - 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.
- Place the rat in the surgical area and apply puralube (artificial tears) to the eyes. Inject a body weight volume of carprofen (analgesic) subcutaneously (subcutaneous [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.
- 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.
- Perform intravenous catheter implantation according to previously published protocols.
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. - Immediately following catheter implantations, secure the rat in a stereotaxic frame to perform Adeno-associated virus (AAV) injections.
- 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. - 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), which can respond to a wide range of frequencies, and therefore has utility for the low-frequency long-term depression (LTD) experiments discussed in this protocol, but also for high-frequency long-term potentiation (LTP) experiments. 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 the medial geniculate nucleus of the thalamus (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 Institutional Animal Care and Use Committee (IACUC) approval, and proper personal protective equipment (PPE) must be used at all times in accordance with institutional guidelines to limit unnecessary exposure. - 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.
- Level the rat's head in the anterior/posterior axis and zero stereotaxic coordinates to bregma.
- 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. The position of screws should be spread across the anterior-posterior axis of the skull, and away from AAV injection site. - Drill bilateral holes for injection of AAV based on the coordinates from the Rat Brain Atlas (Watson and Paxinos) for the medial portion of the medial geniculate nucleus (MGN); in mm from bregma, anteroposterior (AP): -5.4; mediolateral (ML): ±3.0; dorsoventral (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.
- Leave injection cannula in place for 5 min after infusions are complete to allow for diffusion away from the cannula and then slowly withdraw the cannula from the brain.
- Immediately following virus injections, continue to implant optic fibers targeting MGN-LA terminals.
- 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).
- 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.
- Slowly lower the fibers at a rate of 2 mm/min, until the tip of the fiber sits in the dorsal portion of the LA (dorsoventral [DV]: -7.9 mm).
- 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.
- 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.
2. 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.
- Subject rats to daily 1-h cocaine (2 mg/mL) self-administration training sessions under a fixed-ratio schedule 1 (FR1) schedule of reinforcement.
- Place rats in an 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.
- Continue self-administration experiments for at least 10 days 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.
- After acquisition criteria are successfully met, subject rats to 1 h instrumental extinction sessions for 6-10 d.
- 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.
- Have rats continue instrumental extinction daily until an average of < 25 lever presses over two consecutive days occurs.
3. In vivo Optogenetic Induction of LTD
NOTE: Optogenetic inhibition experiments take place 24 h after the final day of instrumental extinction.
- 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.
- Turn on the laser diode according to the 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 the laser. - Measure the light intensity through the patch cord using a light sensor. Adjust the intensity of the laser so that the light output through the patch cable is ~5-7 mW.
- 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.
- Turn on the pulse generator to initiate optogenetic stimulation.
NOTE: Although unlikely, if rats experience any adverse reactions to stimulation, the experiment is immediately terminated, and rats are properly euthanized based on institutional guidelines. - Following LTD induction, keep rats in the cage for 3 min, and then place them back in their home cages.
- 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-minute session.
4. Test the Effect of Optogenetic Stimulation on Cue-Induced Cocaine Seeking
- 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. - 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