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

Summary

The goal of this protocol is to directly manipulate ventral tegmental area receptors to study their contribution to subsecond dopamine release.

Abstract

Phasic dopamine (DA) release from the ventral tegmental area (VTA) to the nucleus accumbens plays a pivotal role in reward processing and reinforcement learning. Understanding how the diverse neuronal inputs into the VTA control phasic DA release can provide a better picture of the circuitry that controls reward processing and reinforcement learning. Here, we describe a method that combines intra-VTA cannula infusions of pharmacological agonists and antagonists with stimulation-evoked phasic DA release (combined infusion and stimulation, or CIS) as measured by in vivo fast-scan cyclic voltammetry (FSCV). Using CIS-FSCV in anesthetized rats, a phasic DA response can be evoked by electrically stimulating the VTA with a bipolar electrode fitted with a cannula while recording in the nucleus accumbens core. Pharmacological agonists or antagonists can be infused directly at the stimulation site to investigate specific VTA receptors' roles in driving phasic DA release. A major benefit of CIS-FSCV is that VTA receptor function can be studied in vivo, building on in vitro studies.

Introduction

Phasic dopamine (DA) release from the ventral tegmental area (VTA) to the nucleus accumbens (NAc) plays a vital role in reward-related behaviors. VTA DA neurons switch from a tonic-like firing (3-8 Hz) to a burst-like firing (>14 Hz)¹, which produces phasic DA release in the NAc. The VTA expresses a variety of somatodendritic receptors that are well-positioned to control the switch from tonic to burst-firing^2,3,4,5. Identifying which of these receptors, and their respective inputs, control phasic DA release will deepen our understanding of how the reward-related circuitry is organized. The purpose of the methodology described here, combined infusion and stimulation with fast-scan cyclic voltammetry (CIS-FSCV), is to quickly and robustly assess the functionality of VTA receptors in driving phasic DA release.

The term combined infusion and stimulation (CIS) refers to pharmacologically manipulating receptors on a group of neurons (here the VTA) and stimulating those neurons to study the receptor's function. In the anesthetized rat, we electrically stimulate the VTA to evoke a large phasic DA signal (1-2 µM) in the NAc core, as measured by fast-scan cyclic voltammetry (FSCV). Infusions of pharmacological drugs (i.e., receptor agonists/antagonists) at the stimulation site can be used to measure the function of VTA receptors by observing the subsequent change in evoked phasic DA release. FSCV is an electrochemical approach that enjoys both high spatial (50-100 µm) and temporal (10 Hz) resolution, and is well-suited to measure reward-related, phasic DA events^6,7. This resolution is finer than other in vivo neurochemical measurements, such as microdialysis. Thus, together, CIS-FSCV is well-suited to assess VTA receptor regulation of phasic dopamine release.

One common way to investigate VTA receptor function is by using a combination of electrophysiological approaches that address how those receptors alter the firing rate of neurons^1,8. These studies are highly valuable in understanding what receptors are involved in driving DA firing upon activation. However, these studies can only suggest what might happen downstream at the axon terminal (i.e., release of a neurotransmitter). CIS-FSCV builds on these electrophysiological studies by answering how the output of VTA burst-firing, phasic DA release, is regulated by receptors located on VTA dendrites and cell bodies. Thus, CIS-FSCV is well-suited to build on these electrophysiology studies. As an example, nicotinic receptor activation can induce burst-firing in the VTA⁹, and CIS-FSCV in the anesthetized rat was used to show that nicotinic acetylcholine receptor (nAChR) activation in the VTA also controls phasic DA release in the NAc^10,11.

Mechanistic examination of phasic DA regulation is also commonly studied using slice preparations alongside with bath application of drugs. These studies often focus on the presynaptic regulation of phasic DA release from dopamine terminals, as the cell bodies are often removed from the slice¹². These preparations are valuable for studying presynaptic receptor effects on dopamine terminals, whereas CIS-FSCV is better suited to study somatodendritic receptor effects on dopamine neurons, as well as presynaptic inputs to the VTA. This distinction is important, because somatodendritic receptor activation in the VTA may have a different effect than NAc presynaptic receptor activation. Indeed, blocking dopaminergic presynaptic nAChRs in the NAc can elevate phasic dopamine release during burst-firing¹³, whereas the opposite is true at VTA somatodendritc nAChRs^10,11.

CIS-FSCV is an ideal approach for studying the ability of VTA receptors to regulate phasic DA release. Importantly, this approach can be performed in an intact rat, either anesthetized or free moving. This approach is suitable for acute studies, to study receptor function in its baseline state^10,14 as well as long-term studies that can assess functional changes in a receptor after drug exposure or behavioral manipulation^11,15.

Access restricted. Please log in or start a trial to view this content.

Protocol

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.

Access restricted. Please log in or start a trial to view this content.

Results

CIS-FSCV was used to study the function of VTA N-methyl-D-aspartate receptors (NMDAR), nicotinic acetylcholine receptors (nAChRs), and muscarinic acetylcholine receptors (mAChRs) in driving phasic DA release in the NAc core. Figure 2 shows representative data for a negative control, infusion of 0.9% saline, before (baseline) and 9 min postinfusion (saline). Figure 2 shows a color plot with potential on the y-axis, time on the x-a...

Access restricted. Please log in or start a trial to view this content.

Discussion

CIS-FSCV provides a unique opportunity to investigate VTA receptor mechanisms underlying phasic DA release. There are two critical steps in order to ensure a proper recording. First, a stable baseline recording must be achieved, with little drift in the evoked DA signal. An important way to increase the likelihood of establishing a stable recording is to ensure that the electrode has had plenty of time to cycle at both 60 Hz and 10 Hz (typically 15 min at 60 Hz, and 10 min at 10 Hz). As the carbon fiber is being cycled, ...

Access restricted. Please log in or start a trial to view this content.

Materials

Name	Company	Catalog Number	Comments
Electrode Filling Solution/Supplies
Micropipette	World Precision Instruments	MF286-5 (28 gauge)
Potassium Acetate	Sigma	236497-100G
Potassium Chloride	Sigma	P3911-25G
Electrode Supplies
Carbon fiber	Thornel	T650
Electrode puller	Narishige International	PE-22	Note: horizontal pullers can be used as well
Glass capillary	A-M systems	626000
Insulated wires for electrodes	Weico Wire and Cable Incorporated	UL 1423	Length; 10 cm; diameter,0.4mm; must get custom made; insulated material should cover 5 cm of the wire
Light Microscope (for viewing and cutting electrode)	Fischer Scientific	M3700
Pin	Phoenix Enterprises	HWS1646	To be soldered onto the insuled electrode wire and reference electrode; connects to headstage
Putty	Alcolin	23922-1003	Used to place electrode on while cutting the carbon fiber
Scalpal Blade	World Precision Instruments	500239	For cutting carbon fiber to the apprpriate length
Silver Wire	Sigma	327026-4G
FSCV Hardware/Software
Faraday Cage	U-Line	H-3618 (36" x 24" x 42")
Potentiostat	Univ. of N. Carolina, Electronics Facility
Stimulating electrode	PlasticsOne	MS303/2-A/SPC	when ordering, request a 22 mm cut below pedestal
TarHeel HDCV Software	University of North Carolina-Chapel Hill	-	https://chem.unc.edu/critcl-main/criticl-electronics/criticl-electronics-hardware/ for ordering information
UEI breakout box	Univ. of N. Carolina, Electronics Facility		https://chem.unc.edu/critcl-main/criticl-electronics/criticl-electronics-hardware/ for ordering information
UEI power supply	Univ. of N. Carolina, Electronics Facility		https://chem.unc.edu/critcl-main/criticl-electronics/criticl-electronics-hardware/ for ordering information
Stimulator Hardware
Neurolog stimulus isolator	Digitimer Ltd.	DS4	Neurolog 800A
Infusion/Stimulation Supplies
Infusion Pump	New Era Syringe Pump	NE-300
Internal Cannula	PlasticsOne	C315I/SPC INTERNAL 33GA
Microliter Syringe	Hamilton	80308
Tubing	PlasticsOne	C313CT/ PKG TUBING 023 X 050 PE50
Surgical Supplies
Cannula Holder	Kopf Instruments	1776 P-1
Cotton Tip Applicators	Vitality Medical	806
Electrode Holder	Kopf Instruments	1770
Heating Pad	Kent Scientific	RT-0501
Povidone Iodine	Vitality Medical	29906-004
Screws	Stoelting	Bone Anchor Screws/Pkg.of 100	1.59 mm O.D., 3.2 mm long
Silver wire reference with AgCl	InVivo Metric	E255A
Square Gauze	Vitality Medical	441408
Stereotax	Kopf Instruments	Model 902 (Dual Arm Bar)
Histological Supplies
Formulin	Sigma	1004960700
Power supply	BK Precision	9110
Sucrose	Sigma	80497
Tungsten microelectrode	MicroProbes	WE30030.5A3
Drugs for infusions
((2R)-amino-5-phosphonovaleric acid	Sigma Aldrich	A5282
N-methyl-D-aspartate	Sigma Aldrich	M3262
Mecamylamine hydrochloride (M9020-5mg)	Sigma Aldrich	M9020
Scopolamine hydrobromide (S0929-1g)	Sigma Aldrich	S0929