Separating Analgesia from Reward within the Ventral Tegmental Area

Abstract

Activation of the dopaminergic mesolimbic reward circuit that originates in the ventral tegmental area (VTA) is postulated to preferentially suppress emotional responses to noxious stimuli, and presumably contributes to the addictive liability of strong analgesics. VTA dopamine neurons are activated via cholinergic afferents and microinjection of carbachol (cholinergic agonist) into VTA is rewarding. Here, we evaluated regional differences within VTA in the capacity of carbachol to suppress rats' affective response to pain (vocalization afterdischarges, VADs) and to support conditioned place preference (CPP) learning. As carbachol is a non-specific agonist, muscarinic and nicotinic receptor involvement was assessed by administering atropine (muscarinic antagonist) and mecamylamine (nicotinic antagonist) into VTA prior to carbachol treatment. Unilateral injections of carbachol (4 μg) into anterior VTA (aVTA) and posterior VTA (pVTA) suppressed VADs and supported CPP; whereas, injections into midVTA failed to effect either VADs or CPP. These findings corroborate the hypothesis that the neural substrates underlying affective analgesia and reward overlap. However, the extent of the overlap was only partial. Whereas both nicotinic and muscarinic receptors contributed to carbachol-induced affective analgesia in aVTA, only muscarinic receptors mediated the analgesic action of carbachol in pVTA. The rewarding effects of carbachol are mediated by the activation of both nicotinic and muscarinic receptors in both aVTA and pVTA. The results indicate that analgesia and reward are mediated by separate cholinergic mechanisms within pVTA. Nicotinic receptor antagonism within pVTA failed to attenuate carbachol-induced analgesia, but prevented carbachol-induced reward. As addictive liability of analgesics stem from their rewarding properties, the present findings suggest that these processes can be neuropharmacologically separated within pVTA.

INTRODUCTION

The addictive liability of analgesics has persistently vexed the treatment of pain. More than a century ago, it was observed that strong analgesics, like opiates and psychostimulants, are highly abused and self-administered by both humans and animals (Spender, 1887; UKMH, 1926; Himmelsbach, 1942; May, 1953). Currently, the nonmedical use of prescription pain relievers is greater than the combined abuse of cocaine, hallucinogens, inhalants, and heroin (SAMHSA, 2011). Moreover, the number of overdose deaths from prescription pain relievers far outnumbered deaths from heroin and cocaine combined in the past decade (NIDA, 2011). Thus, identifying neural mechanisms by which strong analgesics alleviate pain while limiting the potential for addiction is a societal imperative.

The link between analgesia and addiction suggests that the neural substrates of antinociception and reward overlap (Oberst et al., 1943). Franklin (1989, 1998) proposed that the ability of opiates and psychostimulants to induce positive affect underlies their addictive liability and analgesic action. The positive affective state generated by these analgesics presumably reduces the distress that normally accompanies noxious stimulation and injury. This phenomenon is termed “affective analgesia” and reflects preferential suppression of the emotional response to pain.

Activation of dopamine neurons in the ventral tegmental area (VTA) that project to nucleus accumbens (NAc) contributes to reward produced by morphine, amphetamine, other drugs of abuse, and natural reinforcers (Schultz, 2000; Di Chiara, 2002; Kiyatkin, 2002; Wise, 2004; Koob and Volkow, 2010). Activation of this mesoaccumbal dopamine system also contributes to the antinociceptive action of morphine and amphetamine (Altier and Stewart, 1998). These neurons are endogenously activated via cholinergic projections from the laterodorsal tegmental (LTDg) and pedunculopontine tegmental (PPTg) nuclei (Blaha et al., 1996; Omelchenko and Sesack, 2005, 2006) acting on muscarinic and nicotinic receptors (Nisell et al., 1994; Yeomans and Baptista, 1997; Gronier and Rasmussen, 1998; Miller et al., 2005). Microinjecting nicotinic and muscarinic agonists into the VTA excite dopaminergic neurons via activation of local cholinergic receptors (Calabresi et al., 1989; Lacey et al., 1990) and increase the efflux of dopamine in NAc (Gronier et al., 2000; Nisell et al., 1994). Cholinergic activation of mesoaccumbal dopamine neurons contributes to reward (Rada et al., 2000). For example, intra-VTA administration of the nonspecific cholinergic agonist carbachol supports development of conditioned place preference (CPP) learning, and rats learn to self-administer carbachol into VTA, effects mediated by the local muscarinic and nicotinic receptors (Yeomans et al., 1985; Ikemoto and Wise, 2002). Additionally, the capacity of systemically administered morphine to support CPP learning and induce the efflux of dopamine into NAc is partially dependent on acetylcholine receptors in VTA (Miller et al., 2005; Rezayof et al., 2007).

Consistent with the affective analgesia hypothesis, we reported that microinjection of carbachol into VTA produced dose-dependent suppression of vocalization discharges (VADs) in rats (Kender et al., 2008). VADs occur immediately following application of noxious tail shock and are a validated rodent model of pain affect (Caroll and Lim, 1960; Borszcz, 1993, 1995, 2006; Borszcz and Spuz, 2009). However, regional differences within VTA were reported in the capacity of carbachol to activate the brain reward circuit. Carbachol supports development of CPP and is self-administered when delivered to posterior VTA (pVTA), but not the anterior VTA (aVTA) (Ikemoto and Wise, 2002). In our earlier study, affective analgesia followed administration of carbachol into pVTA, but its suppression of VADs when microinjected into aVTA was not assessed. Here we further evaluated the affective analgesia hypothesis by examining regional differences within VTA in the ability of carbachol to support affective analgesia and reward. We also analyzed the contribution of muscarinic and nicotinic receptors to the effects of carbachol within each VTA subregion. Our goal was to determine whether affective analgesia and reward can be neuropharmacologically separated within the VTA.

EXPERIMENTAL PROCEDURES

Animals

One hundred and thirty-six naïve male Long-Evans rats were housed as pairs in polypropylene cages (52cm × 28cm ×22cm) with hardwood chip bedding and given ad libidum access to Rodent Lab Diet 5001 (PMI, Nutrition International, Inc., Brentwood, MO) and water. Housing was provided in a climate-controlled vivarium maintained on a 12:12-hr circadian cycle with lights on at 0700 hrs. All testing was conducted between 0800 and 1700 hrs. Upon arrival, rats were given 5-7 days of acclimatization prior to handling. Rats were handled 2-3 times every third day for 1 week prior to surgery to minimize possible effects of stress from human contact. Following surgery, rats were handled once per day for at least one week before testing to check on their recovery and to further minimize the effects of stress from human contact. All experiments were performed following the guidelines of the United States National Institutes of Health Guide for the Care and Use of Laboratory Animals using protocols approved by the Wayne State University Institutional Animal Care and Use Committee.

Surgery

Rats were anesthetized with sodium pentobarbital (50 mg/kg, i.p.) following pretreatment with atropine sulfate (1 mg/kg, i.p.). A stainless steel 26-gauge cannula guide (Plastics One Inc., Roanoke, VA) was stereotaxically implanted unilaterally at a 15° angle, according to coordinates extrapolated from the rat brain atlases of Paxinos and Watson (1998, 2007), and from our analysis of tyrosine hydroxylase (TH) immunoreactivity within the ventral tegmentum (see below). Three sites along the rostrocaudal axis of the VTA were targeted. The coordinates (in mm) relative to the bregma suture and the top of the skull were for aVTA: AP = − 4.5, ML = + 2.5, DV = − 7.3, for midVTA: AP = − 5.0, ML = + 2.5, DV = −7.3, and for pVTA: AP = − 5.5, ML = + 2.5, DV = −7.3. Guides were affixed to the skull with 4 stainless steel bone screws and cranioplastic cement. Each guide cannula was fitted with a dummy obturator that extended the length of the guide to keep it clear of debris. Rats were given 7-10 days to recover before the initiation of testing.

Tyrosine Hydroxylase (TH) immunocytochemistry

Unless otherwise specified, all chemicals were purchased from Sigma-Aldrich (St. Louis, MO, USA). TH immunoreactivity was performed to localize cathecholaminergic cells within the ventral tegmentum. The immunoreaction was conducted according to the protocol described in Xavier et al. (2005) with nickel intensification. Briefly, serial coronal slices (45 μm) from 8 rats that underwent transcardial perfusion with 4% paraformaldehyde were pretreated with 0.3% H₂O_2, washed with 0.1 M PBS, blocked with goat and bovine albumin serum buffer, incubated in mouse monoclonal tyrosine hydroxylase primary antibody, then incubated in goat anti-mouse secondary antibody (Milipore, Billerica, MA), incubated in a Avidin-Biotin solution (Vector Laboratories, Burlingame, CA), and rinsed with 0.01 M Tris-HCl. The immunoreaction was developed by incubating each section in a diaminobenzene medium with nickel intensification. Finally, the sections were rinsed in distilled H₂O, mounted on microscope gelatin-coated glass slides, dehydrated in ethanol, cleared with CitriSolv™ (Thermo Fisher Scientific Inc., Waltham, MA) and xylene and then covered with Permount® (Thermo Fisher Scientific Inc., Waltham, MA) and coverslipped.

Histology and Microscopy

At the conclusion of testing, rats were killed by carbon dioxide asphyxiation. The injection sites were marked by an injection of 0.25 μl of safarin-O dye performed in the same fashion as drug injections. Brains were extracted and placed in 20% (w/v) sucrose formalin solution for 48-72 hours. Brains were then sectioned in slices of 45 [.proportional]m thickness on a freezing microtome (Leica SM2000R), and injection sites were localized under microscopes (Olympus SZX-ILLB2-100, Fisher Scientific Micromaster 12-561T) with the aid of Paxinos and Watson brain atlases (1998, 2007) and our analysis of TH immunoreactivity within ventral tegmentum by two experimenters, one of whom was unaware of the behavioral outcomes. Injection sites and TH immunoreactivity within VTA subregions were identified at low magnification (1.6 × and 4×). Individual immunoreactive cells were visualized with a 100× oil immersion objective.

Pain Testing

Assessment of pain affect

Research in this laboratory validated vocalization afterdischarges (VADs) as a rodent model of pain affect. These vocalizations occur immediately following application of noxious tailshock, are organized within the forebrain, and have distinct spectrographic characteristics compared to vocalizations that occur during shock (VDS) (Caroll and Lim, 1960; Hoffmeister, 1968; Borszcz, 1995, 2006). Systemically administered drug treatments that preferentially suppress the affective response of humans to pain (Gracely et al., 1978; Price et al., 1985) also preferentially suppress production of VADs (Borszcz et al., 1994). Generation of VADs is suppressed by damage of or drug treatments into forebrain sites known to contribute to production of the affective response of humans to clinical and experimental pain (Mark et al., 1961; Hoffmeister, 1968; Sweet, 1980; Borszcz, 1999; Harte et al., 2000; Zubieta et al., 2001; Borszcz and Leaton, 2003; Nandigama and Borszcz, 2003; Harte et al., 2004; Harte et al., 2011). Additionally, the capacity of noxious tailshock to support fear conditioning is directly related to its production of VADs (Borszcz, 1993, 1995, 2006; Borszcz and Leaton, 2003). In the present study, the effects of experimental treatments on VAD threshold were compared with their effects on the thresholds of other tail shock-elicited responses that are organized at medullary (vocalizations during shock, VDS) and spinal (spinal motor reflexes, SMR) levels of the neuraxis (Caroll and Lim, 1960; Borszcz et al., 1992).

Apparatus

Testing was controlled by custom computer programs via a multifunction interface board (DT-2801, Data Translation, Marlboro, MA) installed in a PC. Rats were placed into custom made Velcro body suits and restrained on a Plexiglas pedestal using Velcro strapping that passes through loops located on the underside of the suits. This design maintains the rat in a crouching posture throughout testing, permits normal respiration and vocalizing, and allows unobstructed access to the head for intracerebral injections (see photograph in Borszcz, 1995). Testing was conducted within a sound attenuating, lighted, and ventilated chamber equipped with a small window that enabled visual monitoring of rats during testing.

Tailshock (20 ms pulses at 25 Hz for 1,000 ms) was delivered by a computer controlled constant current shocker (STIMTEK, Arlington, MA) through electrodes (0-gauge stainless steel insect pins) placed intracutaneously on opposite sides of the tail, 7.0 cm (cathode) and 8.5 cm (anode) from the base. The intensity, duration, and timing of tailshocks were controlled by the computer. Current intensity was monitored by an analog-to-digital converter that digitized (500 Hz sampling rate) an output voltage of the shocker that was proportional to the current delivered.

Spinal motor reflexes (SMRs) were measured with a semi-isotonic displacement transducer (Lafayette Instruments Model 76614, Lafayette, IN) attached to the rat's tail with cotton thread. The arm of the transducer was positioned behind and perpendicular to the tail such that the thread extended in a straight line directly behind the rat. The output voltage of the transducer was amplified (×50) and then digitized (500 Hz sampling rate) by an analog-to-digital converter of the interface board. SMR was defined as movement of the transducer arm by at least 1.0 mm following shock onset. The computer recorded the latency (ms), peak amplitude (mm), and magnitude (cm x ms) of tail movement on each trial. Displacements up to 100 mm can be detected, and latencies in 2 ms increments can be measured.

Vocalizations were measured by a pressure-zone microphone (Realistic model 33-1090, Tandy, Ft. Worth, TX) located on the wall of the testing chamber 15 cm from the rat's head. The microphone was connected to an audio amplifier (Technics model SA-160, Tandy, Ft. Worth, TX) and a 10-band frequency equalizer adjusted to selectively amplify frequencies above 1500 Hz. The filtering of low frequencies prevented extraneous noise (i.e., rats’ respiration and movement artifacts) from contaminating vocalization records. The output of the amplifier was integrated by a Coulbourn Instruments (Allentown, PA) contour following integrator (2 ms time base) and digitized (500 Hz sampling rate) by a separate analog-to-digital converter of the interface board. The peak intensity (in decibels: SPL, B scale), latency (ms), and duration (ms) of vocalizations during the shock epoch (VDS) and for the 2,000 ms interval following shock termination (VAD), were recorded by the computer.

Procedure

For 2 consecutive days prior to pain testing, rats were adapted to the testing apparatus for a period of 20 min each day to minimize the effects of restraint stress. In all experiments, testing began 5 - 8 min following completion of intracerebral injections. Test sessions consisted of 20 randomly presented trials: 16 different intensities of tailshock (.02 – 2.5 mA) and 4 trials without tailshock that permitted the assessment of false alarm rates. Randomization was designed to control for the impact of any particular tailshock on subsequent response generation, and to prevent rats from anticipating the intensity of successive tailshocks. Trials were presented with a minimum intertrial interval of 30 sec and each test session concluded within 20 - 25 min. These procedures cause no observable damage to the tail. Following each test session, the testing apparatus was cleaned with 5% ammonia hydroxide (Fanselow, 1985) and Nilotron® (Nilodor, Inc., Bolivar, OH) to eliminate stress odors.

Drug Injections

Intracerebral injections were administered in a constant volume of 0.25 μl via 33-guage injectors that extended 1.7 mm beyond the end of the cannulae. All injections were made at a constant rate over 1 min via an infusion pump (Harvard Model PHD 2000), and injectors were left in place for 2 min after the completion of injections to aid the diffusion of drugs into tissue. Based on the evaluation of dye spread, behavioral effects from control injections outside the VTA (and within the midVTA), and previous Fos plume analysis (Smith and Berridge, 2005) it is estimated that drugs spread .2 - .3 mm from the site of injection.

In the dose-response experiment (Experiment 1), rats received unilateral microinjections of carbachol (1 μg/side, 2 μg/side, and 4 μg/side) as well as vehicle (normal saline = saline) into the VTA 5 - 8 min prior to the test sessions. In the antagonism experiment (Experiment 2), rats received either 30 and 60 μg of atropine, 15 and 45 μg mecamylamine, or vehicle into the VTA 7 - 10 min prior intra-VTA administration of carbachol (4 μg) or vehicle. Every animal in the muscarinic antagonism group received unilateral injections of saline + saline, saline + 4 μg carbachol, 30 μg atropine + 4 μg carbachol, 60 μg atropine + 4 μg carbachol, and saline + 60 μg atropine. Every animal in the nicotinic antagonism group received unilateral injections of saline + saline, saline + 4 μg carbachol, 15 μg mecamylamine + 4 μg carbachol, 45 μg mecamylamine + 4μg carbachol, and saline + 45 μg mecamylamine. Doses of drugs were taken from our earlier report (Kender et al., 2008) and from results of preliminary experiments. All drugs were dissolved in normal sterile saline solution. Carbamoylcholine chloride (carbachol), atropine sulfate (atropine), and mecamylamine hydrochloride (mecamylamine) were purchased from Sigma-Aldrich (St. Louis, MO, USA).

Test sessions were separated by 4 - 7 days. The order of injections was counterbalanced using a quasi-Latin square design that maintained the vehicle injection at either the beginning or the end of the test sequence. This design permits evaluation of the effects of repeated testing on baseline responding.

Conditioned Place Preference (CPP)

Apparatus

The place conditioning apparatus (Med Associates, Inc., St. Albans, VT) consisted of two dimly-lit Plexiglas chambers (43 cm long × 21.5 cm wide × 30.5 cm high) separated by an opaque black wall with a guillotine door in the middle (8 cm wide). One chamber differed from the other by wall pattern (horizontal versus vertical black and white lines, each 2.5 cm wide) and floor type (horizontal versus vertical bars). Each chamber was equipped with three sets of 16-beam infrared (I/R) emitter-detector arrays (Med Associates, St. Albans, VT): one emitter-detector array was mounted on each of the longer sides at a height of 12 cm and two emitter-detector arrays were mounted on each side at a height of 4.5 cm. Each photobeam array was spaced by 2.5 cm from one another. Rats’ movements resulted in the breaking of photobeams which was monitored by Activity Monitor software, version 5 (Med Associates, St. Albans, VT) that tracked the location of rats, recorded the amount of time spent in each side of the chamber, and provided indices of velocity of locomotion and distance traveled. Chambers were enclosed in sound-attenuating cabinets.

Procedure

An unbiased procedure was used to establish CPP. Each experiment consisted of three sessions: Habituation (Day 1), Conditioning (Days 2 to 7), and Testing (Day 8). On the Habituation day (H), the guillotine door was opened and the rats in a drug-free and naive state were given free access to both chambers for 15 min with time spent in each chamber recorded. In order to minimize novelty effects and to ensure that rats had equal access to both chambers, half of the rats in each group were placed in front of the open guillotine door facing one chamber whereas the other half faced the opposite chamber. On the first Conditioning day, either vehicle or drug(s) were administered into the rats’ VTA after which they were immediately confined to one chamber for 15 min with the guillotine door closed. The next day, rats were administered the opposite treatment into the VTA and restricted to the opposite chamber for 15 min. This procedure was repeated for the remaining days of conditioning. Thus, each rat was exposed to each chamber three times, in an alternate fashion. On Test day (T), each rat was placed in the opposite chamber than on the Habituation day, facing the opened guillotine door. Rats had access to both chambers for 15 min in a drug-free state and the time spent in each chamber was recorded. After each rat exposure, the chambers were cleaned with 0.75 % Alconox (VWR International LLC., Radnor, PA) and then aerated with Nilotron® (Nilodor, Inc., Bolivar, OH) to eliminate odors from other rats.

Drug Injections

Injection volume and procedure were identical to that performed prior to pain testing. To establish the capacity of carbachol to support CPP learning (Experiment 3), the highest dose used in pain testing experiments (4 μg) was administered into the aVTA, midVTA or pVTA during the conditioning phase. To evaluate acetylcholine receptor subtypes that contribute to carbachol-induced CPP learning the highest doses of antagonists used in the pain testing experiments (atropine = 60 μg, mecamylamine = 45 μg) were administered into the aVTA or pVTA 7 – 10 min prior to the intra-VTA injection of 4 μg carbachol during the conditioning phase (Experiment 4). Antagonists were not administered into the midVTA because carbachol failed to support CPP learning when injected into this subregion (see below).

Data analyses

Pain Testing

Some rats did not complete all testing sessions due to illness (n = 2) or blocked and damaged cannulas (n = 5). This attrition resulted in unequal sample sizes across doses of carbachol necessitating that data be considered from independent groups and analyzed accordingly.

Dose-Response Analysis

After each test session, data were reorganized in ascending order according to tail shock intensity. SMR, VDS, and VAD thresholds for each rat were calculated as the minimum current intensity from a string of at least 2 consecutive intensities that generated the response. For the aVTA (n = 6) and pVTA (n = 7), response thresholds were directly compared across doses of carbachol using repeated-measures multivariate analysis of variance (MANOVA). The effects of dose on individual responses were analyzed by one-way analysis of variance (ANOVA). The doses of carbachol that elevated thresholds above baseline were assessed by comparing thresholds after saline and carbachol treatments using Dunnett's t-test for multiple post-hoc comparisons.

Anatomical Specificity

Data from rats (n = 7) were analyzed separately as anatomic controls when histological evaluation revealed that their injection sites were outside of the VTA. Data from rats (n = 9) in which the injection sites were determined to be within the midVTA were also analyzed separately. Thresholds following saline and 4 μg carbachol injections were compared via Student's t-test of independent groups.

Pharmacological Specificity

The capacity of atropine (aVTA: 30 μg, n = 5, 60 μg, n = 6; pVTA: 30 μg, n = 5, 60 μg, n = 5) or mecamylamine (aVTA: 15 μg, n = 5, 45 μg, n = 5; pVTA: 15 μg, n = 6, 45 μg, n = 5) to reduce increases in response thresholds generated by injection of 4 μg carbachol into aVTA and pVTA was assessed via one-way ANOVA. Doses of antagonists that significantly reduced carbachol-induced increases in response thresholds were revealed via Dunnett's t-test for multiple post-hoc comparisons.

CPP

Development of CPP was defined as rats spending significantly more time in the drug-paired compartment on the Test day that followed conditioning compared to on the Habituation day that preceded conditioning. Comparison of the number of seconds spent in the drug-paired compartment on the Test day versus Habituation day was made using Student's t-test for paired samples for each VTA subregion (Experiment 3: aVTA, n = 6, midVTA, n = 8, pVTA, n = 7; Experiment 4: atropine groups aVTA, n = 9, pVTA, n = 6; mecamylamine groups aVTA, n = 7, pVTA, n = 6). Comparison of CPP learning across VTA subregions was made using the CPP score. The CPP score is defined as time spent in the carbachol-paired chamber after conditioning (T) minus time spent in the carbachol-paired chamber before conditioning (H). A comparison across all three subdivisions was made via one-way ANOVA with pairwise comparisons made via Student's t-test for independent groups.

RESULTS

TH Immunohistochemistry

Figure 1 shows the distribution of catecholaminergic neurons within aVTA (top), midVTA (middle) and pVTA (bottom) of the adult male Long-Evans rat. The aVTA and pVTA were defined as located within the ventral midbrain dorsal to the mammillary bodies (4.68 mm to 5.16 mm posterior to bregma) and dorsal to the interpeduncular nucleus (5.88 mm to 6.60 mm posterior to bregma), respectively. These coordinates are similar to those previously reported for aVTA and pVTA (Ikemoto and Wise, 2002). The midVTA had not been previously described but was defined based on its support of different behavioral outcomes as compared to aVTA and pVTA (see below). The midVTA was localized within the ventral midbrain beginning at the caudal tail of the mammillary bodies (5.28 mm posterior to bregma) and ending at the rostral extent of the interpeduncular nucleus (5.76 mm posterior to bregma). Stereotaxic coordinates for intracerebral implants into VTA subregions were adjusted accordingly.

Figure 1.

Catecholamine-producing neurons within VTA. aVTA (A), midVTA (B), and pVTA (C). The staining was obtained via Tyrosine Hydroxylase Immunohistochemistry.

The location of the injection sites throughout VTA are depicted in Figure 2.

Figure 2.

Histological reconstruction of microinjection sites within subregions of the VTA. A,B,C. Each symbol signifies a microinjection site in the pain experiment: red circles in the aVTA (A), black triangles in the midVTA (B), and red squares in the pVTA (C). D,E,F. Each symbol signifies an injection site in the reward experiments in the aVTA (D), midVTA (E), and pVTA (F): red circles and black triangles represent 4 μg carbachol injections, and the blue squares represent the location of an antagonist + 4 μg carbachol injections. The coronal diagrams are modified from Paxinos and Watson (2007). Numbers on the right side of diagrams represent coordinates in millimeters posterior to bregma.

Pain Testing

Response Profile

As demonstrated by Carroll and Lim (1960), SMR, VDS, and VAD reflect nociceptive processing at spinal, medullary, and forebrain levels of the neuraxis, respectively (also see Borszcz et al.1992; Borszcz and Leaton, 2003). Consistent with our previous reports (Borszcz, 1993; Harte et al., 2000; Nandigama and Borszcz, 2003; Kender et al., 2008), responses organized rostrally within the neuraxis were rarely generated without those integrated more caudally. VAD generation, without concomitant elicitation of VDS and SMR occurred on 1.14% of all trials. VDS was elicited without SMR on 2.81% of the trials in which VDS was the most rostrally elicited response. False alarm rates for each response were low: SMR = 1.44%, VDS = 0.12%, VAD = 0.12%. The low incidence of false alarms indicates that responses were not induced by drug administration, were not occurring spontaneously, and were not conditioned responses to the context, but instead were generated by tailshock.

Response Performance

The effects of carbachol on the performance of SMR, VDS and VAD were evaluated (data not shown). Performance variables at threshold obtained after saline treatment were compared with performance variables at threshold attained after administration of carbachol. The dose-response analysis (Experiment 1) revealed that administration of carbachol into aVTA or pVTA did not alter the latency, amplitude, and magnitude of SMRs, Fs < 1.73, ps > .20. The latency, amplitude, and duration of VDSs and VADs were also not affected by carbachol treatment (Fs < 1.78, ps > .15), except for VDS latency following administration of carbachol into the aVTA, F(3,20) = 5.37, p < .01. Post hoc analysis (Dunnett's test) indicated that VDS latency was significantly shorter after 4 μg carbachol administration than after saline administration (p < .01). This decrease in VDS reaction time reflects facilitation of performance. These findings indicate that carbachol-induced increases in response thresholds are unlikely to be the result of interference with the capacity of rats to fully perform the responses.

During pharmacological specificity analysis (Experiment 2), performance variables obtained after saline + saline and saline + 4 μg carbachol were compared. In the pVTA, carbachol treatment did not affect any performance variable, except VDS duration which was significantly shorter after carbachol treatment, F(1,19) = 10.21, p < .01. This reduction of VDS duration reflects a performance decrement that may have contributed to the increase in VDS threshold. In the aVTA, carbachol treatment did not affect performance of any response, except VAD amplitude which was significantly higher after carbachol treatment, F(1,12) = 9.90, p < .01. This increase in VAD amplitude is a facilitation of performance; therefore, it is unlikely that the increase in VAD threshold is the result of carbachol-induced interference with VAD performance.

Experiment 1: Dose-Response Analysis & Anatomical Specificity

The dose-dependent effects of carbachol administered into aVTA and pVTA on SMR, VDS and VAD thresholds are depicted in Figure 3. Comparison between the subgroups of rats that received saline first or last in the testing sequence revealed no differences in thresholds following saline treatment (aVTA: SMR, t(4) = 1.26, p = .28, VDS, t(4) = 1.26, p = .28, VAD, t(4) = .40, p = .71; pVTA: SMR, t(5) = 1.45, p = .21, VDS, t(5) = .41, p = .70, VAD, t(5) = 1.07, p = .33), indicating that repeated administration of carbachol did not alter baseline thresholds. Also, the comparison of SMR, VDS and VAD thresholds following saline treatment revealed that baseline thresholds of these responses did not differ in the pVTA group (F(2,18) = .61, p = .55). In aVTA, comparison of SMR, VDS, and VAD baseline thresholds indicated marginal differences in baseline responding (F(2,15) = 3.86, p = .045); however, post-hoc pairwise comparisons between responses failed to reveal any differences in baseline thresholds (Tukey HSD, all ps > .05).

Figure 3.

Dose-response effects of carbachol on pain thresholds. The effects of unilateral administrations of carbachol (C) or saline into the aVTA (A), pVTA (B), or midVTA and sites surrounding the VTA (other) (C) on the mean (±SEM) thresholds of spinal motor reflexes (SMR), vocalizations during shock (VDS), and vocalization afterdischarges (VAD). Asterisk (*) indicates thresholds significantly elevated compared to saline.

Administration of carbachol into the aVTA or pVTA differentially elevated response thresholds. Comparison of VAD, VDS, and SMR thresholds following carbachol and saline treatments (repeated measures MANOVA, Wilk's λ) revealed significant main effects of treatment (aVTA F(3, 20) = 34.01, p < .001; pVTA (F(3, 24) = 62.19, p < .001), and response (aVTA F(2, 19) = 45.40, p < .001; pVTA F(2, 23) = 232.97, p < .001), and a significant Treatment x Response interactions (aVTA F(6, 38) = 9.08, p < .001; pVTA F(6, 46) = 40.00, p < .001). The interactions indicate that carbachol microinjections within the aVTA (Fig. 3A) and pVTA (Fig. 3B) preferentially increased VAD threshold.

In the aVTA, comparisons of each response threshold across saline and carbachol treatments revealed that thresholds of all three responses were elevated by carbachol microinjections (SMR F(3,20) = 3.40, p < .05, VDS F(3,20) = 6.50, p < .01, and VAD F(3,20) = 29.06, p < .001). Post-hoc pairwise comparisons (Dunnett's test, all ps < .05) of thresholds following saline administration and each carbachol treatment revealed that the lowest dose of carbachol to significantly elevate thresholds was 4 μg for SMR, and 2 μg for VDS and VAD. Comparison of VDS and VAD thresholds revealed that VAD threshold was significantly elevated as compared with VDS threshold following administration of carbachol in doses of 1 μg (t(10) = 3.01, p < .05), 2 μg (t(10) = 3.73, p < .05), and 4 μg (t(10) = 9.32, p < .001).

In the pVTA, VDS and VAD thresholds also showed a significant elevation across treatments (VDS, F(3,24) = 4.89, p < .01, VAD, F(3,24) = 332.97, p < .001); however, the slight increase in SMR threshold observed following carbachol administration into the aVTA was not observed (F(3,24) = 1.62, p > .05). Post-hoc pairwise comparisons (Dunnett's test, all ps < .05) of thresholds following saline and carbachol treatments revealed that the lowest dose of carbachol to significantly elevate thresholds was 4 μg for VDS, and 2 μg for VAD. VAD threshold was significantly elevated as compared with VDS threshold following administration of carbachol in doses of 2 μg (t(11) = 4.48, p < .001) and 4 μg, t(11) = 4.05, p < .01. VAD thresholds did not differ following administration of the highest dose of carbachol (4 μg) into aVTA versus pVTA (t(11) = 1.63, p = .13).

Histological analysis of microinjection sites revealed an area of VTA between aVTA and pVTA where administration of carbachol failed to elevate response thresholds. We labeled this area as midVTA (see above). No differences in response thresholds were observed following saline versus 4 μg carbachol treatments into the midVTA (Fig. 3C, SMR, t(14) = 0.94, p > .80, VDS, t(14) = 1.48, p > .16, VAD, t(14) = 2.03, p > .06). Additionally, microinjection of 4 μg carbachol into areas dorsal and lateral of the aVTA or pVTA failed to raise response thresholds above those observed following saline treatment (Fig. 1C, SMR, t(12) = 0.62, p > .55, VDS, t(12) = 0.93, p > .37, VAD, t(12) = 1.81, p > .09). These injection sites were located in retroethmoid nucleus, rostral linear nucleus of the raphé, substania nigra (pars compacta and pars reticulata), red nucleus, pararubral nucleus, and interstitial nucleus of Cajal.

Experiment 2: Pharmacological Specificity of Analgesia

The effects of muscarinic and nicotinic receptor antagonism on carbachol-induced increases in response thresholds are depicted in Figure 4. Consistent with the results from the dose-response experiment, administration of 4 ug carbachol into either the aVTA or pVTA generated significant increases in VDS and VAD thresholds (VDS, ts > 3.22, ps < .01; VAD, ts > 8.59, ps < .001), but SMR thresholds were not affected, ts < 1.50, ps > .05. Furthermore, repeated drug administration into the aVTA or pVTA did not alter baseline thresholds. No differences in response thresholds after saline + saline treatments were observed in subgroups that were administered this treatment first or last in the testing sequence (aVTA: SMR, t(8) = .59, p = .57, VDS, t(8) = .59, p = .57, VAD, t(8) = 1.22, p = .26; pVTA: SMR, t(8) = 1.22, p =.26, VDS, t(8) = 1.85, p = .10, VAD, t(8) = 1.55, p = .16).

Figure 4.

Effects of cholinergic antagonists on carbachol-induced increases in pain thresholds. The effects of the co-administration of atropine (A) or mecamylamine (M) on increases in vocalization thresholds produced by injection of carbachol (C, 4 μg) into aVTA (A & C) or pVTA (B & D). Data are plotted as the mean (±SEM) threshold of spinal motor reflexes (SMRs), vocalizations during shock (VDSs), and vocalization afterdischarges (VADs). Asterisk (*) indicates thresholds significantly elevated above saline (sal) + saline treatment. Pound sign (#) indicates thresholds significantly reduced compared to saline + carbachol treatment.

Atropine

Figures 4A & B depict the effects of muscarinic receptor antagonism on increases in response thresholds generated by carbachol administered into VTA. In both regions of VTA, pre-treatment with atropine produced dose-dependent antagonism of carbachol-induced increases in thresholds for VDS (aVTA: F(2,20) = 4.52, p < .05, pVTA: F(2,20) = 4.30, p < .05) and VAD (aVTA: F(2,20) = 25.52, p < .001, pVTA: F(2,20) = 9.93, p < .001). In the aVTA and pVTA, post hoc analyses (Dunnett's test) indicated that 30 μg atropine reduced carbachol-induced increases in VAD thresholds (ps < .05), but thresholds remained significantly elevated compared to baseline thresholds, ts > 2.59, ps < .05. Similarly, carbachol-induced increases in VDS thresholds were reduced following injection of 30 μg atropine into aVTA (p < .05), and marginally reduced following its administration into the pVTA (p < .055). These VDS thresholds remained elevated compared to baseline, ts > 2.62, ps < .05. Following pretreatment of aVTA or pVTA with 60 μg atropine, carbachol-induced increases in VAD and VDS thresholds were also reduced (ps < .05), and no longer differed from baseline thresholds (ts < 1.82, ps > .05).

Atropine alone did not alter baseline response thresholds. In both aVTA and pVTA, comparisons of thresholds of each response after saline + saline and 60 μg atropine + saline revealed no significant differences (SMR, ts < 1.35, ps > .05, VDS, ts < 1.65, ps > .05, VAD, ts < 1.55, ps > .05).

Mecamylamine

The effects of nicotinic receptor antagonism on increases in response thresholds generated by carbachol administered into VTA are shown in Figures 4C & D. Mecamylamine administered into aVTA blocked the carbachol-induced increases in VAD threshold in a dose-dependent manner (F(2,19) = 6.36, p < .01). Post hoc analysis (Dunnett's test) revealed that the 15 μg dose of mecamylamine treatment had no effect on carbachol-induced increases of VAD threshold (p > .05). The 45 μg dose attenuated this increase of VAD threshold (p < .01) although it remained elevated compared to baseline (t(13) = 4.44, p < .001). Although mecamylamine did not attenuate the carbachol-induced increase in VDS threshold in a dose-dependent manner (owing to the slight increase in threshold following treatment with the 15 μg dose), the 45 μg dose produce a significant reduction in the carbachol-induced increase in VDS threshold (t(13) = 2.25, p < .05) returning it to baseline (t(13) = 1.71, p > .05).

In contrast, mecamylamine administration in the pVTA did not reduce the carbachol-induced increases in VAD or VDS thresholds (VAD, F(2,21) = .24, p = .79; VDS, F(2,21) = .05, p = .95). Both VAD and VDS thresholds remained significantly elevated compared to baseline following treatment with 4 ug carbachol in combination with either dose of mecamylamine (VAD, ts > 6.50, ps < .001; VDS, ts > 2.75, ps < .05).

Mecamylamine alone did not alter baseline response thresholds. Administration of saline + 45 μg mecamylamine did not alter the baseline thresholds of any response (SMR, ts < .74, ps > .05, VDS, ts < 1.66, ps > .05, VAD, ts < .90, ps > .05).

CPP Learning

Initial chamber preference

Rats were tested using an unbiased procedure in a two chamber CPP apparatus. Because there are reports that rats sometimes have an initial tendency to prefer one of the two chambers (for a discussion on this methodological issue, see Bardo and Bevins, 2000), a comparison of the amount of time spent in each chamber on the Habituation Day was conducted. Collapsing data across all groups, rats initially spent an equal amount of time in each chamber (Chamber A, Mean = 446 sec, SEM = 10.35; Chamber B, Mean = 454 sec, SEM = 10.30) indicating that an initial chamber preference did not confound the results of CPP training, t(56) = .39, p > .05.

Experiment 3: Carbachol-Induced CPP Learning

The capacity of 4 μg carbachol injected into the aVTA, midVTA or pVTA to support CPP learning is depicted in Figure 5A. Administration of 4 μg carbachol into either the aVTA or pVTA was effective in supporting CPP learning. The amount of time spent in the carbachol-paired chamber was directly compared before (H) and after (T) conditioning. Rats spent significantly more time in the carbachol-paired compartment after conditioning (pVTA: t(6) = 3.98, p < .01; aVTA: t(5) = 4.04, p < .01). Alternately, administration of 4 μg carbachol into the midVTA failed to support CPP learning. There was no significant difference between the time spent in the carbachol-paired compartment before and after conditioning (t(7) = .22, p > .05).

Figure 5.

Effects of cholinergic agents on CPP. A. Unilateral administration of 4 μg carbachol (C) promotes CPP learning in the aVTA and pVTA, but not midVTA. Asterisk (*) indicates significantly more time spent in the carbachol paired compartment after (Post, T = test day) versus before (Pre, H = habituation day) conditioning. B. Co-administration of either atropine (A, 60 μg) or mecamylamine (M, 45 μg) with 4 μg carbachol into the aVTA or pVTA prevents the formation of carbachol-induced CPP.

Experiment 4: Pharmacological Specificity of CPP Learning

The pharmacological specificity of carbachol-induced CPP was evaluated by pretreating the VTA with muscarinic (atropine) or nicotinic (mecamylamine) receptor antagonists prior to carbachol administration. Unilateral administration of 60 μg atropine or 45 μg mecamylamine prior to 4 μg carbachol injections into either the pVTA or aVTA prevented the development of carbachol-induced CPP learning (Fig. 5B). The amount of time spent in the drug paired compartment before and after CPP conditioning did not differ in groups administered atropine + carbachol or mecamylamine + carbachol into pVTA (atropine: t(5) = .22, p > .05, mecamylamine: t(5) = 1.77, p > .05), or atropine + carbachol or mecamylamine + carbachol into aVTA (atropine: t(8) = 2.15, p < .05, mecamylamine: t(6) = .99, p > .05). Preliminary data (not shown) demonstrated that atropine or mecamylamine administered alone into aVTA or pVTA did not alter compartment preference.

Comparison between Carbachol-Induced Affective Analgesia and Reward

Volume of the microinjections played no significant role in altering any of the responses in the pain experiments, as revealed by comparisons between groups that received 4 μg carbachol (.25 μl) and groups that received saline + 4 μg carbachol (.50 μl) in either the aVTA (SMR, t(14) = 1.03, p = .32, VDS, t(14) = .76, p = .46, VAD, t(14) = .42, p = .68), nor in the pVTA (SMR, t(16) = .97, p = .35, VDS, t(16) = .12, p = .90, VAD, F(16) = 1.67, p = .12). Thus, the data from rats that received 4 μg carbachol or saline + 4 μg carbachol were pooled (aVTA, n = 16; pVTA, n = 18) and their responses were compared with the midVTA group (n = 9).

As shown in Figure 6A, comparison of VAD thresholds following injection of 4 μg carbachol into the three VTA subregions revealed significant differences, F(2,40) = 46.4, p < .001. Planned pairwise comparisons revealed the mean threshold current intensity necessary to elicit VAD was significantly lower when carbachol was delivered into the midVTA when compared with aVTA (t(23) = 8.91, p < .001) and pVTA (t(25) = 10.08, p < .001). No difference in VAD thresholds was observed following injection of carbachol into aVTA versus pVTA (t(32) = .65, p = .52). VAD thresholds of each rat administered 4 μg carbachol into aVTA, midVTA, and pVTA are depicted in Figure 6E.

Figure 6.

Comparison of carbachol-induced affective analgesia and CPP across subregions of the VTA. Unilateral 4 μg carbachol injections support the development of CPP learning and affective analgesia in both the aVTA and pVTA, but not in the midVTA. A. Carbachol (4 μg) administered in midVTA (yellow) failed to elevate VAD threshold as compared with the same dose administered in aVTA and pVTA (red). Asterisk (*) indicates thresholds significantly elevated compared to midVTA. B.Carbachol (4 μg) administered aVTA and pVTA (red), but not in midVTA (yellow) supported the acquisition of CPP learning. Asterisk (*) indicates CPP score significantly elevated compared to midVTA. C, D. Photomicrographs of representative injections sites in the aVTA, midVTA, and pVTA (left to right) from the pain experiments (C) and reward experiments (D). E, F. The data obtained from individual rats that received carbachol in the pain experiments (E) or reward experiments (F) that were summarized in A and B. VAD thresholds and CPP scores are plotted as a function of millimeters posterior to bregma with designations for aVTA, midVTA, and pVTA. The insets represent sagittal VTA diagrams with the red-yellow color-coding corresponding roughly to the intensity of the effects observed with carbachol, with red being more effective and yellow being less effective.

Representative photomicrographs of injection sites within aVTA, midVTA, and pVTA are shown in Figure 6C.

Similarly, analysis of CPP learning following 4 μg carbachol injections into aVTA, midVTA, and pVTA revealed that the CPP score was significantly affected by the VTA subregion into which carbachol was delivered (Fig. 6B, F(2,18) = 7.32, p = .005). Direct planned comparisons revealed that rats spent significantly less time in the drug-paired chamber when carbachol was delivered into the midVTA compared with aVTA (t(12) = 3.36, p < .01) and pVTA (t(13) = 3.31, p < .01). Comparisons between 4 μg carbachol treatment in aVTA and pVTA indicated no difference in the CPP score (t(11) = .03, p = .98). CPP scores of each rat administered 4 μg carbachol into aVTA (n =6), midVTA (n = 8), and pVTA (n = 7) are depicted in Figure 6F. Representative photomicrographs of injection sites within aVTA, midVTA, and pVTA are shown in Figure 6D.

The regional effects within VTA of carbachol on affective analgesia and CPP and regional differences within VTA of acetylcholine receptor subtypes in supporting affective analgesia and CPP are summarized in Table 1. As described above, administration of carbachol into the aVTA and pVTA supported both affective analgesia and CPP; whereas, injection of the same dose of carbachol into midVTA failed to support either affective analgesia or CPP. The aVTA and pVTA differ in the acetylcholine receptor subtypes that contribute to affective analgesia and CPP. Within the aVTA, both muscarinic and nicotinic receptors contribute to affective analgesia and CPP. Muscarinic receptors with the pVTA also contribute to both affective analgesia and CPP. However, nicotinic receptors within the pVTA differentially contribute to affective analgesia and CPP. Whereas, nicotinic receptors within pVTA also contribute to development of CPP they do not contribute to production of affective analgesia. That is, nicotinic receptor antagonism within the pVTA failed to attenuate carbachol-induced affective analgesia, but eliminated carbachol-induced CPP.

Table 1.

Differential involvement of cholinergic receptors in aVTA, midVTA, and pVTA on affective analgesia and CPP generated by carbachol

	Affective Analgesia			CPP
	aVTA	midVTA	pVTA	aVTA	midVTA	pVTA
Carbachol	+	—	+	+	—	+
Atropine + Carbachol	—	NA	—	—	NA	—
Mecamylamine + Carbachol	—	NA	+	—	NA	—

Note: CPP = conditioned place preference, carbachol (4 μg), atropine (60 μg), mecamylamine (45 μg), plus sign = carbachol-induced affective analgesia or CPP, minus sign = no (or reduced) carbachol-induced affective analgesia or CPP, NA = not applicable, circled cells denote differential effects on affective analgesia and CPP

DISCUSSION

This study is the first to directly compare the extent of overlap between cholinergically mediated reward and affective analgesia within different regions of VTA. Development of CPP learning and increases in VAD threshold were used as measures of reward and affective analgesia, respectively. We tested Franklin's affective analgesia hypothesis that postulates that activation of the brain reward circuit preferentially suppress the affective dimension of pain (Franklin, 1989, 1998). Administration of the nonspecific cholinergic agonist carbachol (4 μg) into aVTA and pVTA supported both the development of CPP and affective analgesia; whereas, its injection into midVTA failed to support either CPP or affective analgesia. These findings support the affective analgesia hypothesis. However, the extent of overlap between the neural substrates underlying reward and affective analgesia is only partial, as different cholinergic receptors contribute to these effects in different subregions of VTA. Both nicotinic and muscarinic receptors mediate affective analgesia in aVTA, as shown by the ability of both atropine and mecamylamine to reduce carbachol-induced increases in VAD threshold. Alternately, only muscarinic receptors mediate affective analgesia in pVTA because atropine, but not mecamylamine, was effective in attenuating carbachol-induced increases in VAD threshold. On the other hand, the rewarding effects of carbachol are mediated by the activation of both nicotinic and muscarinic receptors in both aVTA and pVTA, as both atropine and mecamylamine prevented the development of CPP learning in both VTA subregions. That is, nicotinic receptor antagonism within pVTA failed to attenuate carbachol-induced increases in VAD threshold, but prevented carbachol-induced CPP.

This latter finding indicates a separation within pVTA of the neuropharmacology that underlies affective analgesia and reward. Combined administration of carbachol and mecamylamine into pVTA suppressed the affective response to pain and prevented engagement of the brain reward circuit. As activation of the brain reward circuit presumably mediates the addictive liability of strong analgesics, the present findings may provide insight into mechanisms that can dissociate analgesia from the potential of addiction.

It is important to note that administration of an antagonist prior to a non-specific agonist is not identical to administration of the other agonist alone. In other words, administration of mecamylamine + carbachol is not equivalent to administration of muscarine. Previous studies of VTA neurons demonstrated that activation or inhibition of particular receptors subtypes alters their response to activation of other receptor subtypes (Fiorillo and Williams, 2000; Paladini and Williams, 2004; Arencibia-Albite et al., 2007). Accordingly, our data suggests that there is a subpopulation of neurons in pVTA that contain both muscarinic and nicotinic receptors, and that activation of the muscarinic receptors coupled with inhibition of the nicotinic receptors contributes to affective analgesia, but not reward. Further studies are needed to characterize the potential analgesic and/or rewarding effects of direct nicotinic or muscarinic receptor activation within VTA by injecting nicotine or muscarine, with or without actively inhibiting the other receptor subtype.

The proposal that combined administration of an antagonist and non-specific agonist is different than administration of an agonist for the non-inhibited receptor can account for the CPP results in the present study. Pretreatment of aVTA or pVTA with atropine or mecamylamine abolished development of carbachol-induced CCP. However, both muscarinic and nicotinic receptors within VTA are directly involved in reward (Yeomans et al., 2000; Pons et al., 2008). Thus, pretreatment with atropine or mecamylamine may have altered the response of the non-inhibited receptor to carbachol, thereby preventing the non-inhibited receptor to engage the brain reward circuit and support CPP.

In the present study, administration of carbachol into aVTA and pVTA were equivalent in supporting development of CPP. The previous study (Ikemoto and Wise, 2002) that found that carbachol is not reinforcing in aVTA used a significantly lower dose of carbachol (0.09 μg) as compared with our doses that proved efficacious in inducing affective analgesia (2 and 4 μg). Our lowest dose (1 μg) of carbachol failed to induce affective analgesia in any VTA subregion. Therefore, it is possible that low doses of carbachol are not rewarding, but high doses are able to induce CPP in aVTA.

Carbachol-induced affective analgesia and reward is thought to be mediated by its binding to muscarinic and nicotinic receptors located on dopamine and GABA neurons within VTA resulting in direct activation and disinhibition of dopaminergic projections from the VTA, respectively (Omelchenko and Sesack, 2005, 2006; Zhang et al., 2005). Thus, carbachol administered into VTA mimics the endogenous activation of VTA dopamine neurons via cholinergic projections from the LTDg and PPTg, resulting in subsequent dopamine release into sites that receive afferents from VTA. Cholinergic activation of VTA following electrical stimulation of LTDg (Forster and Blaha, 2000; Yeomans et al., 2001; Forster et al., 2002; Lester et al., 2010) or intra-VTA administration of carbachol (Westerink et al., 1996), oxotremorine M (muscarinic agonist, Gronier et al., 2000) or nicotine (Blaha et al., 1996) results in increased efflux of dopamine in NAc. This cholinergically-mediated mesoaccumbal dopamine activation is associated with the rewarding effects of morphine (Rezayof et al., 2007), cocaine (You et al., 2008), and lateral hypothalamic self-stimulation (Rada et al., 2000). Whether this cholinergically-mediated release of mesoaccumbal dopamine supports affective analgesia has yet to be determined; however, dopamine efflux into NAc contributes to the suppression of paw-licking in the formalin test generated by injection of morphine into VTA (Altier and Stewart, 1998), and placebo analgesia in humans is correlated with increased accumbal dopamine neurotransmission (Scott et al., 2008). Regarding the results of the present study, it will be important to compare the capacity of carbachol to generate dopamine release in NAc following its administration into aVTA, midVTA, and pVTA. In particular, ascertaining the effect of mecamylamine + carbachol administered into pVTA on dopamine efflux in NAc is important given the capacity of this treatment to prevent carbachol-induced reward but not carbachol-induced affective analgesia. For example, muscarinic and nicotinic receptors in VTA differentially contribute to the dynamics of dopamine efflux in NAc (Forster and Blaha, 2000). It will be important to evaluate how the dynamics of dopamine release in NAc (Yeomans et al., 2001), and other brain sites that receive dopaminergic afferents from VTA (see below), contribute to affective analgesia and reward.

The dissociation of affective analgesia and reward in pVTA may reflect engagement of a subpopulation of muscarinically activated and nicotinically inhibited dopamine neurons that project to sites that contribute preferentially to affective analgesia versus reward. For example, the anterior cingulate cortex (aCC), insula, medial thalamus, and amygdala receive dopaminergic afferents from VTA (Moore and Bloom, 1978; Deniau et al., 1980; Swanson, 1982; Oades and Halliday, 1987), are implicated in processing the affective dimension of pain (Weigel and Krauss, 2004; Kulkarni et al., 2005; Spuz and Borszcz, 2012), and dopamine release into these sites generates antinociception (de la Mora et al., 2010; Shyu et al., 1992; Burkey et al., 1999; Lopez-Avila et al., 2004). The action of dopamine in these sites also supports reward (Baxter and Murray, 2002; Goeders and Smith, 1983, 1986; Hitchcott and Phillips, 1998; Volkow et al., 2005; Di Pietro et al., 2008), so it will be important to establish their relative contribution of affective analgesia versus reward. Alternately, there is no evidence of involvement of the ventrolateral periaqueductal gray (vPAG) in reward, which receives afferents from VTA (Kirouac et al., 2004) and is a nodal structure in the endogenous antinociceptive circuit (Basbaum and Fields, 1984; Borszcz, 1999). Projections from the vPAG to the medial thalamus and amygdala contribute to suppression of the affective dimension of pain (Borszcz, 1999; Harte et al., 2000) and projections from the medial thalamus (Munn et al., 2009), amygdala (Pavlovic and Bodnar, 1998), and aCC (LaBuda and Fuchs, 2005) to the vPAG also support affective analgesia. The role of cholinergically activated afferents from VTA to vPAG in engaging this endogenous antinociceptive circuit should also be evaluated.

The VTA also provides GABAergic and glutamatergic afferents to NAc, vPAG, amygdala, prefrontal cortex and other sites (Carr and Sesack, 2000; Hnasko et al., 2012; Kirouac and Mabrouk, 2004; Van Bockstaele and Pickel, 1995). Furthermore, a subset of dopaminergic neurons in VTA co-releases physiologically significant amounts of glutamate (Sulzer and Rayport, 2000; Trudeau, 2004). The relative contribution of these neurotransmitter systems in supporting affective analgesia and reward also warrants study. For example, GABAergic and glutamatergic neurons are not evenly distributed within VTA (Nair-Roberts et al., 2008) which could underlie the regional differences in the contribution of VTA to affective analgesia and reward. Furthermore, NAc medium spiny neurons provide reciprocal GABAergic input to VTA by targeting non-dopamine neurons some of which project back to NAc (Xi et al., 2011). These non-dopaminergic neurons have not been identified but they may also provide synaptic inputs that modulate the activation of dopaminergic projections from the VTA. As these GABAergic projections from NAc to VTA are inhibited by μ-opioids they may contribute to opiate-mediated analgesia and reward.

An additional unique, and unexpected, finding was identification of a behaviorally distinct subregion of VTA. Unlike aVTA and pVTA, injection of carbachol into midVTA failed to support either affective analgesia or reward. To the best of our knowledge, there is only one published study that reported an anterior-posterior bimodal activation of VTA. Marcangione and Rompré (2008) evaluated c-fos expression throughout the anterior-posterior extent of VTA elicited by self-stimulation of the posterior mesencephalon. Expression was greatest in aVTA and pVTA with midVTA exhibiting the fewest number of c-fos positive cells. More recently, relief from peripheral pain via peripheral nerve block was shown to generate negative reinforcement in support of CPP learning in rats (Navratilova et al., 2012). This effect was accompanied by an increase in the number of Fos-positive dopaminergic neurons in the pVTA but not in the midVTA (the aVTA was not evaluated), suggesting that activation of dopaminergic neurons in pVTA, but not midVTA, contribute the production of reward associated with pain relief. These results highlight that VTA is a heterogenous structure and encourages further evaluation of the anatomical, pharmacological and behavioral differences across VTA subregions.

Conclusion

The affective dimension of pain has a profound impact on human health. It motivates those in pain to seek health care, and underlies the development of emotional disturbances such as anxiety, fear, and depression that contribute to the suffering of patients in chronic pain (Loeser, 2000). The use of strong prescription analgesics that are effective in suppressing the affective dimension of pain has increased dramatically, as has the non-medical use, abuse, and dependence on these analgesics, and rate of overdose deaths associated with them (Becker et al., 2008; Bohnert et al., 2011). The abuse liability generated by these analgesics reflects their capacity to engage the brain reward circuit, and it is this capacity to engage the brain reward circuit that is believed to contribute to the affective analgesia generated by these treatments. The results of the present study indicate that the neuropharmacology within the brain reward circuit that contributes to addictive liability and analgesia are separable, and therefore provides a basis for the possible development of strong analgesics with limited addictive/abuse potential.

Research Highlights.

• Cholinergic activation of the ventral tegmental area (VTA) generates analgesia and reward.

• Analgesia and reward can be neuropharmacologically separated within the posterior VTA (pVTA).

• Nicotinic receptor antagonism within pVTA fails to attenuate analgesia, but prevents reward.

• A new subdivision of the VTA (midVTA) was identified.

• Cholinergic activation of the midVTA failed to support either analgesia or reward.

Abbreviations

VTA

ventral tegmental area

CPP

conditioned place preference

VAD

vocalization afterdischarge

VDS

vocalizations during shock

SMR

spinal motor reflex

NAc

nucleus accumbens

TH

tyrosine hydroxylase

LTDg

laterodorsal tegmental nucleus

PPTg

pedunculopontine tegmental nucleus