Combined Infusion and Stimulation with Fast-Scan Cyclic Voltammetry (CIS-FSCV) to Assess Ventral Tegmental Area Receptor Regulation of Phasic Dopamine

All experiments were conducted according to the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals and were approved by both Elizabethtown College and Yale University Institutional Animal Care and Use Committee (IACUC). This protocol is specific to the anesthetized rat preparation of utilizing CIS-FSCV.

1. Presurgical preparations

1. Electrode solution preparation
   1. To make the electrode backfill solution, prepare a solution of 4 M potassium acetate with 140 mM potassium chloride¹⁶.
2. Electrode preparation
   1. Using vacuum suction, insert a T-650 carbon fiber (7 µm in diameter) into a borosilicate glass capillary (length = 100 mm, diameter = 1.0 mm, inside diameter = 0.5 mm).
   2. Once the carbon fiber has been placed inside the glass capillary, place the glass capillary into a vertical electrode puller, with the heat element roughly in the middle of the capillary. Set the heater to 55 with the magnet turned off.
   3. After the capillary is pulled, carefully raise the upper capillary holder so that the tip of the electrode is not surrounded by the heating element.
   4. Using sharp scissors, cut the carbon fiber that is still connecting the two pieces of the capillary. This will result in two separate carbon fiber microelectrodes.
   5. Under a light microscope, carefully cut the exposed carbon fiber with a sharp scalpel, so that the carbon fiber extends approximately 75-100 µm beyond the end of the glass.
   6. Using a light microscope, ensure that the electrode is free of cracks along the capillary. Also ensure that the seal, where the carbon fiber exits the capillary, is difficult to notice and free from cracks.
      NOTE: A good seal will help reduce noise during recordings. See published studies^17,18,19 for a more detailed protocol.
3. Reference electrode fabrication
   1. Solder a gold pin to a 5 cm silver wire.
   2. Attach the anode to a metal paper clip or other conductor, the cathode to a pin, and apply a voltage (~2 V) while the paper clip and silver wire is submerged in 0.1 M HCl.
   3. Cease the voltage once a white coating (AgCl) appears on the silver wire.
4. Preparing electrode for implantation
   1. Solder a gold pin to a thin insulated wire (~10 cm in length, <0.50 mm diameter).
   2. Remove ~5 cm of insulation from the wire opposite to the gold pin.
   3. Fill the electrode approximately halfway with electrode solution.
   4. Insert insulated wire into the electrode.
      NOTE: The wire should make contact with the carbon fiber inside the electrode.

2. Electrode implantations

1. Give adult, male, Sprague Dawley rats (250−450 g) an intraperitoneal injection (1.5 g/kg or 1 mL/kg volume) of 0.5 g/mL urethane dissolved in sterile saline. Start with an initial urethane dose of 1.0−1.2 g/kg. If the animal is still responsive to the noxious stimulus test (tail pinch) 20 min after urethane administration, administer an additional 0.3−0.5 g/kg urethane for a 1.5 g/kg total dose.
   NOTE: For preparation of the 0.5 g/mL urethane solution, add 10 g of urethane to 10 g (~10 mL) of saline. Urethane is a carcinogen and must be handled with care. Urethane is an important anesthetic, as it does not alter levels of dopamine, as do other anesthetics such as ketamine/ xylazine and chloral hydrate^20,21.
2. Once the animal is deeply anesthetized and is not responsive to noxious stimuli (e.g., toe pinch), place it in the stereotaxic frame. Apply ophthalmic lubricant to each eye of the rat.
   NOTE: This is a non-survival surgery, but good aseptic technique is encouraged.
3. Clean the rat's scalp using a two-stage scrub (i.e., an iodopovidone scrub followed by a 70% ethanol scrub; perform with a 3 cycle repetition).
4. Cut away the scalp tissue using sterilized needle nose tweezers and surgical scissors. Remove a significant amount of tissue to make room for the various implantations outlined below.
5. Gently clean the skull surface using sterilized cotton tip applicators. Then apply 2−3 drops of 3% hydrogen peroxide to help identify the lambda and bregma.
6. Using a stereotaxic or hand drill (1.0 mm, ~20,000 rpm), drill a 1.5 mm diameter hole 2.5 mm anterior to the bregma and 3.5 mm lateral to the bregma. Partially (about halfway, until it is firmly in place) implant a screw (1.59 mm O.D., 3.2 mm long) in this hole. It is recommended to use sterile saline to irrigate while drilling to prevent thermal injury.
7. For the reference electrode, drill a 1.0 mm diameter hole 1.5 mm anterior and 3.5 mm lateral to the bregma, in the left hemisphere.
8. By hand, insert ~2 mm of reference wire into this hole, while wrapping the reference wire around and under the head of the implanted screw.
9. Fully implant the screw, pinning down the reference electrode in place.
10. In the right hemisphere, drill a 1.5 mm diameter hole 1.2 mm anterior and 1.4 mm lateral to the bregma.
11. Gently remove the dura using tweezers.
12. For the stimulating electrode, drill a square hole (2 mm anterior-posterior, 5 mm medial-lateral) centered at 5.2 mm posterior and 1.0 mm lateral to the bregma.
13. Using the stereotactic arm bars, lower the bipolar stimulating electrode/guide cannula 5 mm below the dura. In case of bleeding during the implantation of the electrode, use sterile cotton swabs and gauze to minimize bleeding.
    NOTE: The bipolar stimulating electrode used in this method is prefitted with a guide cannula (Table of Materials). The internal cannula used with this item should be flushed with the prongs on the bipolar stimulating electrode when fully inserted into the guide cannula. This will allow the internal cannula to sit directly in between the two prongs of the stimulator, which sit about 1 mm apart. A similar protocol is described elsewhere¹⁴.
14. Using the stereotactic arm bars, lower the carbon fiber microelectrode 4 mm below the dura. This location is at the most dorsal portion of the striatum.
15. Connect the reference wire and carbon fiber to a potentiostat.
16. Apply a triangular wave form (-0.4−1.3 V, 400 V/s) for 15 min at 60 Hz, and again for 10 min at 10 Hz.
    NOTE: Typically, when applying waveforms to carbon fiber microelectrodes in the brain, oxide groups are added to the surface of the carbon fiber. Equilibrium of this reaction must be reached prior to recording; otherwise significant drift will occur¹⁹. Cycling the electrode at higher frequencies (60 Hz) allows the carbon fiber to achieve equilibrium faster.

3. Optimizing carbon fiber and stimulating electrode/guide cannula locations

1. Set the stimulator to produce a bipolar electrical waveform, with a frequency of 60 Hz, 24 pulses, 300 µA current, and pulse width of 2 ms/phase.
2. Gently lower the stimulator in increments of 0.2 mm from 5 mm to 7.8 mm below the dura. At each increment, stimulate the VTA.
   NOTE: At more dorsal depths (5−6 mm), stimulation of the brain will typically (~80% of the time) cause the whiskers of the rat to twitch. At further depths, the whiskers will cease twitching, which occurs between 7.5−8.2 mm below the dura. When the whiskers cease twitching, the stimulating electrode will be near or at the VTA. This will not occur in every rat, and lack of whisker twitching should not be taken as a sign that the bipolar stimulating electrode/infusion cannula is misplaced. Whisker twitching may not occur for all anesthetics (e.g., isoflurane).
3. Continue to lower the bipolar stimulating electrode/guide cannula until a stimulation produces phasic DA release at the carbon fiber microelectrode (currently in the dorsal striatum).
   NOTE: DA release in the dorsal striatum will not always occur if the bipolar electrode is implanted in the VTA, but observation of DA release in the dorsal striatum upon VTA stimulation is usually a good sign that a good signal will be observed in the NAc core.
4. Lower the carbon fiber microelectrode until it is at least 6.0 mm below the dura. This is the most dorsal part of the NAc core.
5. Stimulate the VTA and record the peak amplitude of the DA peak.
6. Lower or raise the carbon fiber microelectrode at the site that produces the greatest DA release.
7. Ensure that the peak of the DA response is a clear oxidation peak at 0.6 V and a reduction peak at -0.2 V. These peaks are indicative of DA.

4. Combination infusion and stimulation FSCV recording

NOTE: Figure 1 shows the timeline for recording before and after VTA microinfusion.

1. Once the carbon fiber and stimulating electrode/guide cannula location has been optimized, stimulate for ~20−30 min.
   NOTE: Under the current stimulation parameters, do not stimulate any more than once every 3 min, to allow for vesicular reloading²².
2. After achieving a stable baseline (<20% variation over five stimulations), gently lower the internal cannula by hand into the guide cannula that is prefitted into the bipolar stimulator.
3. Take an additional 2−3 baseline recordings to ensure that the cannula insertion itself did not cause a change in the evoked signal. In some cases, insertion and removal of the internal cannula can cause damage to the VTA. If the signal drastically changes over this baseline period (>20%), then take an additional 3−4 recordings until the baseline restabilizes.
4. Using a syringe pump and microsyringe, infuse 0.5 µL of solution (e.g., 0.9% saline, N-methyl-D-aspartate [NMDA], (2R)-amino-5-phosphonovaleric acid [AP5]) into the VTA over a 2 min period.
5. Postinfusion, leave the internal cannula for at least 1 min prior to removal.
   NOTE: Some drugs may require leaving the internal cannula for a longer time based on the drug kinetics, and removal of the internal cannula may cause the drug to travel back up through the internal cannula. If there is concern, one could leave the internal cannula in the guide cannula during the entirety of the recording. Otherwise, recording can begin after this 1 min interval.
6. Continue recording every 3 min to measure postinfusion effects.
   NOTE: If infusing a control solution, and no effect is observed, it is possible to infuse a second time¹⁰. If there is altered DA release caused by inserting the internal cannula or saline infusion, the signal typically recovers to baseline within 30 min.

5. Histological verification of electrode placement

1. At the end of the experiment, create a small lesion at the recording site using the carbon fiber microelectrode.
   1. If the electrode must be preserved for postexperiment calibration, then use a tungsten wire placed in a glass capillary protruding ~100 µm beyond the capillary tip. In this case, raise the electrode from the brain, replace the recording electrode with the tungsten electrode, and lower it to the same dorsoventral coordinate.
      NOTE: The carbon fiber may be used to lesion the brain as well and will provide a more accurate representation of the location of the recording site; however, the experimenter will lose the ability to calibrate these electrodes.
2. To lesion the recording site, apply voltage using a power supply. Start at 1 V and increase by 1 V every 10 s until 10 V is reached.
3. Euthanize the animal using a lethal intraperitoneal injection of pentobarbital (150 mg/kg).
4. Perfuse the rat using a 4% formalin solution.
5. Remove the head from the rat using a sharpened guillotine.
6. Using rongeurs, remove the connective tissue and skull surrounding the brain, and gently dislodge the brain from any remaining tissue.
7. Store the brain in 4% formalin for 1 day and then transfer it to 30% sucrose.
   NOTE: Perfusion with 4% formalin is not necessary to see the lesion site, although as a best practice it will improve the reconstruction of the lesion site.
8. Create 30 µm slices of the brain using a cryostat.
9. Mount the slices on slides and cover with a cover slip.
10. Denote the location of the carbon fiber microelectrode lesion and bipolar stimulator/infusion cannula location using a light microscope.