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Dopamine, acting at the D1 family receptors (D1R) is critical for the functioning of the amygdala, including fear conditioning and cue-induced reinstatement of drug self administration. However, little is known about the different contributions of the two D1R subtypes, D1 and D5. W identified D1-immunoreactive patches in the primate that appear similar to the intercalated cell masses reported in the rodent; however, both receptors were present across the subdivisions of the primate amygdala including the basolateral amygdala (BLA). Using immunoelectron microscopy, we established that both receptors have widespread distributions in BLA. The D1R subtypes colocalize in dendritic spines and terminals, with D1 predominant in spines and D5 in terminals. Single cell PCR confirmed that individual BLA projection neurons express both D1 and D5 mRNA. The responses of primate BLA neurons to dopamine and D1R drugs were studied using in vitro slices. We found that responses were similar to those previously reported in rat BLA neurons and included a mixture of postsynaptic and presynaptic actions. Given this we investigated the distribution of D1R in the rat BLA and found that there were similarities between the species, such as more prominent D5 localization to presynaptic structures. The higher affinity of D5 for dopamine suggests that presynaptic actions may predominate in the BLA at low levels of dopamine, while postsynaptic effects increase and dominate as dopaminergic drive increases. The results presented here suggest a complex action of dopamine on BLA circuitry that may evolve with different degrees of dopaminergic stimulation.
Dopaminergic neurotransmission at D1 family receptors (D1R) is critical for the functioning of the amygdala. Hence, D1R activation in the basolateral amygdala (BLA) is necessary for the formation of cross-modal associations between conditioned stimuli and unconditioned stimuli in fear conditioning paradigms (Lamont and Kokkinidis, 1998; Nader and LeDoux, 1999; Guarraci et al., 1999; Greba and Kokkinidis, 2000; Macedo et al., 2007), and injections of D1R antagonists into the amygdala have anxiolytic effects (de la Mora et al., 2005). Moreover, D1R activation in the BLA is also essential for cue-induced reinstatement of drug self administration (Ciccocioppo et al., 2001; See et al., 2001; See et al., 2003; Berglind et al., 2006).
Recently, the cellular effects of dopamine activation of D1R in the amygdala have begun to be explored. D1R activation has been shown to increase the excitability of projection neurons (Kroner et al., 2005; Pickel et al., 2006; Yamamoto et al., 2007), and inhibitory interneurons in the basolateral complex (Loretan et al., 2004; Kroner et al., 2005), although GABAergic neurons in the intercalated cell groups are inhibited by D1R activation (Marowsky et al., 2005). Furthermore, activation of D1R attenuates excitatory glutamatergic input onto BLA projection neurons from the prefrontal cortex (Rosenkranz and Grace, 2002), perhaps via suppressing NMDA receptor mediated currents associated with these inputs (Pickel et al., 2006). Clearly D1R activation plays a complex role in regulating the input-output relationship of the BLA.
Despite the established importance of D1R in the normal function of the BLA, the relationship of D1R to BLA circuitry is unclear. Critically, the D1 family of dopamine receptors consists of two separate receptors, the D1 and D5 receptors (Civelli et al., 1993; Gingrich and Caron, 1993). While available pharmacological tools do not differentiate between these receptors, a growing body of evidence points to important functional differences (Sunahara et al., 1991; Tiberi et al., 1991; Hersi et al., 2000; Liu et al., 2000; Lee et al., 2002; Centonze et al., 2003; Laplante et al., 2004).
Consistent with results from D1R binding (Scibilia et al., 1992), in the rat, low levels of D1 mRNA have been reported in the BLA (Mansour et al., 1991) with higher levels of D1 mRNA (Maltais et al., 2000) and immunolabeling (Fuxe et al., 2003; Jacobsen et al., 2006) seen in the intercalated cell groups of the amygdala. Two studies have examined the subcellular distribution of D1 in the amygdala but with contrasting findings. In the first study of the BLA, D1-immunoreactivity (-IR) was reported mainly in cell bodies and dendrites, with spines and axon terminals more rarely seen (Pickel et al., 2006). The second study in the BLA and intercalated cell groups reported that D1-IR was most commonly seen in dendritic spines, with dendritic shafts, axonal elements and glia less commonly seen (Pinto and Sesack, 2008). Levels of D5 in the amygdala have not been examined directly, but low levels were inferred from studies of D1R binding in D1-knockout mice (Montague et al., 2001). To date, no study has examined the subcellular distribution of D5 receptors in the BLA.
The goal of this study was to establish the pattern of distribution of D1 receptors in the BLA with a larger, quantitative analysis and compare that directly with the D5 receptor distribution. We focused our work on macaque monkey tissue to better relate to our previous work on D1 and D5 in monkey prefrontal cortex. In addition, we used in vitro slice electrophysiology to examine some of the actions of D1R stimulation and examine whether previous findings in rat amygdala could be replicated in a non-human primate model. Finally, we quantified the distribution of D1 and D5 in rat BLA to compare to our monkey findings.
Two antibodies were used in this study. The rat anti- D1 antiserum (Sigma-Aldrich, St. Louis, MO, #D187) was prepared against a 97 amino acid synthetic peptide corresponding to the C-terminus of the human D1 receptor. The antiserum stains 2 major bands at 40–45 and 65–75 kD (Hersch et al., 1995), and all staining at the light and electron microscopic levels was abolished when the antiserum was preincubated with 0.5 mg/ml of D1-GST fusion protein (Smiley et al., 1994). The D5 antiserum was a rabbit polyclonal antiserum raised against residues 428–438 of the D5 receptor. This sequence is common to both rat and human D5 receptors. This antiserum reacts to D5-expressing recombinant Sf9 cells but not Sf9 cells expressing D1, D2, D3 or D4 (Khan et al., 2000). Western blot in macaque PFC, striatum and hippocampus labeled a single band with a molecular weight of approximately 53–54 kD (Bordelon-Glausier et al., 2008), in line with the predicted molecular weight of the D5 receptor of approximately 53 kD (Sunahara et al., 1991; Tiberi et al., 1991). Immunohistochemical staining of macaque brain was abolished when the antiserum was pre-incubated with the cognate peptide (Bordelon-Glausier et al., 2008).
Tissue from eight Macaca mulatta monkeys and 5 adult male rats were used for this study. The care of the animals and all anesthesia and sacrifice procedures in this study were performed according to the National Institutes for Health Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee of Emory University. The animals were sacrificed with an overdose of pentobarbital (100mg/kg) and then perfused with a flush of Tyrode’s solution. The flush was followed by 4 liters of fixative solution of 4% paraformaldehyde/0.2% glutaraldehyde/0–0.2% picric acid in phosphate buffer (0.1M, pH 7.4; PB). The brain was blocked and post-fixed in 4% paraformaldehyde for 2–12 hours. Coronal, 50 µm thick vibratome sections of the medial temporal lobe were cut and stored frozen at −80°C in 15% sucrose until immunohistochemical experiments were performed.
Single-label immunoperoxidase labeling was performed using rat anti-D1 at a 1:500 dilution or rabbit anti-D5 antisera at 1:500. The single-label immunoperoxidase labeling for D1 and D5 was performed as described previously (Muly et al., 2003). Briefly, sections were thawed, incubated in blocking serum (3% normal goat serum, 1% bovine serum albumin, 0.1% glycine, 0.1% lysine) in 0.01 M phosphate buffered saline, (PBS; pH 7.4) for 1 hour and then placed in primary antiserum diluted in blocking serum. After 36 hours at 4°C, the sections were rinsed and placed in a 1:200 dilution of biotinylated donkey anti-rat IgG (Jackson Immuno Research, West Grove, PA) for D1 or goat anti-rabbit IgG (Vector, Burlingame, CA) for D5 for 1 hour at room temperature. The sections were then rinsed, placed in avidin-biotinylated peroxidase complex (ABC) (ABC Elite, Vector, Burlingame, CA) for 1 hour at room temperature, and then processed to reveal peroxidase using 3,3'-diaminobenzidine (DAB) as the chromagen. Sections were then post-fixed in osmium tetroxide, stained en bloc with uranyl acetate, dehydrated, and embedded in Durcupan resin (Electron Microscopy Sciences, Fort Washington, PA). Selected regions were mounted on blocks, and ultrathin sections were collected onto pioloform-coated slot grids and counterstained with lead citrate. Control sections processed as above except for the omission of the primary immunoreagent, did not contain DAB label upon electron microscopic examination.
To examine the possibility of co-localization of the two D1R subtypes, we performed double-label immunogold/DAB experiments. In one condition D1 was labeled with immunogold, and D5 was labeled with DAB. In a second condition the labels were reversed. Amygdala tissue sections were thawed and rinsed in PBS. They were incubated in blocking serum (3% normal goat serum, 1% bovine serum albumin, 0.1% glycine, 0.1% lysine and 0.5% fish gelatin) in PBS for one hour and then placed in the primary antiserum diluted at the same concentrations as the single label experiments overnight at 4°C. The sections were removed from the primary antiserum, rinsed in PBS and placed in secondary antiserum (1nm gold conjugated goat anti-rat, used at 1:100, Nanoprobes, Yaphank, NY and biotinylated goat anti-rabbit, used at 1:200, Jackson Immuno Research, West Grove, PA; 1nm gold conjugated goat anti-rabbit, used at 1:100, Nanoprobes, Yaphank, NY and biotinylated donkey anti-rat, used at 1:200, Jackson Immuno Research, West Grove, PA) overnight at 4°C. The tissue was then rinsed in PBS, placed in 2% glutaraldehyde for 20 minutes, rinsed in PBS, rinsed in 2% acetate buffer, silver-intensified for four minutes (HQ silver, Nanoprobes, Yaphank, NY), then rinsed in acetate buffer and in PBS. The sections were incubated in ABC overnight at 4°C and reacted in the same manner as the single-label material. Control sections were processed in which primary antisera were omitted or in which one primary antisera was used and matched with secondary antisera raised to the other species used in our experiments. These sections showed no evidence for nonspecific DAB or immunogold labeling or for cross reactivity of our secondary’s for the other primary antisera.
In order to quantify the extent of co-localization of D1 and D5, tissue sections were incubated in a cocktail of the primary immunoreagents rat anti-D1 and rabbit anti-D5 at a dilution of 1:500 and compared to tissue sections that were incubated with rat anti-D1 alone or rabbit anti-D5 alone at a dilution of 1:500. This cocktail procedure has been described in detail (Muly et al., 2001), and has been used in subsequent studies (Lei et al., 2004; Mitrano and Smith, 2007; Bordelon-Glausier et al., 2008). Briefly, the tissue sections in the cocktail condition were incubated with both primary antisera, then in a cocktail of biotinylated secondary IgGs and in ABC to reveal D1 and D5. DAB was used as the chromagen for both D1 and D5. The D1 alone and D5 alone conditions were processed as described above.
For our quantitative analyses in monkey BLA, four animals were used: three female, one male, ranging in age from 2–5 years. No differences in receptor localization related to monkey age were observed. The single-label DAB material was analyzed as previously described (Muly et al., 2003). Blocks of tissue from the BLA were made and cut in ultrathin sections that were examined using a Zeiss EM10C electron microscope. Regions of the grids containing neuropil were selected based on the presence of label and adequate ultrastructural preservation. Fields of immunoreactive elements in the neuropil were randomly selected, and images were collected at a magnification of 31,500 using a Dualvision cooled CCD camera (1300 × 1030 pixels) and Digital Micrograph software (version 3.7.4, Gatan, Inc., Pleasanton, CA). Images selected for publication were saved in TIFF format and imported into an image processing program (Canvas 8; Deneba Software, Miami, FL). The contrast was adjusted, and the images were cropped to meet size requirements. For D1 a total of 617 micrographs representing 3,767 µm2 of BLA were analyzed across four monkeys and a total of 1014 labeled profiles were counted. For D5 a total of 544 micrographs representing 3,321 µm2 of BLA were analyzed across four monkeys and a total of 994 labeled profiles were counted. For our rat studies we examined tissue from 5 adult male rats. In rat BLA, we examined 293 micrographs of tissue labeled for D1 receptor, representing 1788 um2 of BLA neuropil, and counted a total of 565 labeled profiles. For D5 in the rat, we examined 244 micrographs, representing 1489 um2 of BLA neuropil, and counted 479 labeled profiles.
On each micrograph, DAB-labeled profiles were identified and classified as spines, dendrites, terminals, axons, glia or unknown based on ultrastructural criteria (Peters et al., 1991) as previously described (Muly et al., 2003). Profiles that could not be clearly characterized based on these criteria were considered unknown profiles. The number of immunoreactive profiles was tabulated and the distributions (excluding the unknown profiles) compared with a Chi-square analysis.
For the analysis of the immunogold/DAB, we examined ultrathin sections from the surface of each block where both immunoperoxidase label and immunogold label were visible. Because immunogold label can be noisy, we sought to avoid false-positive labeling by avoiding the very surface of each block, where non-specific gold particles tend to accumulate. We compared the immunogold label in a given structure to the surrounding background level of immunogold labeling, as well as the size of the silver intensified gold particles. If the profile qualitatively contained more immunogold label than the background level, we deemed that acceptable immunogold labeling.
For the cocktail double-label experiments in monkey, blocks of tissue from BLA were made for the D1/D5 cocktail condition, D1 alone and D5 alone conditions. Fields of the neuropil were randomly selected, and images were collected at a magnification of 20,000. An ANOVA sample size analysis (SigmaStat, Version 2.03, SPSS Inc.) indicated that the minimum sample size required to have a statistical power of 80% and a minimum detectable difference in group means of five was 474 images; therefore, we analyzed 629 images in the D1 alone condition, 489 images in the D5 alone condition and 572 images in the D1/D5 cocktail condition. In each experimental condition, the number of images analyzed from each monkey was similar. The images were coded and the analysis of these images was performed by an experimenter blind to the identity of the material until all analysis was complete. On each micrograph, spines and axon terminals were identified using the previously described ultrastructural criteria (Peters et al., 1991), then classified as immunopositive or immunonegative, and the percentage of identified spines or terminals that were immunopositive was calculated. The mean percentage of immunopositive spines and axon terminals were tabulated for each condition and compared across antigen conditions using an ANOVA. The results are reported as mean ± standard error.
The methods for obtaining in vitro brain slices were similar to those described previously in rodents (Rainnie, 1999; Rainnie et al., 2006). Briefly, blocks of unfixed amygdala tissue were placed into oxygenated ice cold “cutting” aCSF (130 mM NaCl, 30 mM NaHCO3, 3.5 mM KCl, 1.1 mM KH2PO4, 6 mM MgCl2, 1 mM CaCl2, 10 mM glucose) supplemented with 5 mM kynurenic acid to minimize glutamate-induced neurotoxicity. Tissue blocks were then glued to the stage of an OTS 4000 vibratome (Electron Microscopy Instruments) and 350 µm coronal slices of monkey amygdala were sectioned and placed in oxygenated “cutting” aCSF at 32°C for 1 hour to recover from the sectioning procedure prior to experimentation. Slices were then transferred to control aCSF of the following composition: NaCL 130 mM, 30 mM NaHCO3, 3.5 mM KCl, 1.1 mM KH2PO4, 1.3 mM MgCl2, 2.5 mM CaCl2, 10 mM glucose for at least 30 minutes prior to the onset of whole-cell patch clamp recording.
Individual slices containing the BLA were placed in a Warner Series 20 recording chamber (Warner Instruments, Hamden, CT) mounted on the fixed stage of a Leica DM-LFS microscope (Leica Microsystems). Slices were fully submerged and continuously perfused at a rate of 1–2 ml per min with heated (32°C) and oxygenated control ACSF. Neurons were visualized using infrared (IR) illumination and a 40X water immersion objective (Leica Microsystems). Images were captured using an IR sensitive charge-coupled device (CCD) digital camera (Orca ER, Hamamatsu, Tokyo Japan) coupled to a Meteor-II video frame grabber (Matrox Electronic Systems Ltd, Dorval, Canada), and displayed on a computer monitor using Simple PCI 6.11 software (Compix Inc., Cranberry Township, PA).
For whole-cell patch-clamp recording, thin walled borosilicate glass patch electrodes (WPI, Sarasota, Fl.) were pulled on a Flaming/Brown micropipette puller (Model P-97, Sutter Instruments). Patch electrodes had resistances ranging from 4–8 MΩ, when filled with a standard patch solution that contained (in mM): K-Gluconate (138), KCl (2), MgCl2 (3), phosphocreatine (5), K-ATP (2), NaGTP (0.2), HEPES (10). The patch recording solution was adjusted to a pH of 7.3 with KOH and had a final osmolarity of 280 mOsm. Whole-cell patch clamp recordings were obtained as previously described (Rainnie et al. 2004), using an Axopatch-1D amplifier (Molecular Devices, Sunnyvale, CA), a Digidata 1320A A–D interface, and pClamp 9.2 software (Molecular Devices). In cell-attached mode, patch electrode seal resistance was considered acceptable if it was >1.5 GΩ. Whole-cell patch-clamp configuration was established in current-clamp mode and neurons were excluded from analysis if they showed a resting membrane potential (Vm) more positive than −55 mV, and/or had an action potential that did not overshoot > +5 mV. Data from current-clamp recordings were sampled at 10 kHz and filtered at 5 kHz.
To examine the current-voltage relationship of the response to drug application, voltage ramps were applied from −100 to 0 mV at a rate of 10 mV/s, before, during, and after drug application. During drug application, voltage ramps were applied only after a steady-state drug response had been achieved. Steady-state whole-cell currents were elicited from a holding potential of −60 mV unless otherwise stated. Projection neurons were tested for their response to bath application of dopamine (DA, 50 µM), and the D1 receptor selective agonist, (1R,3S)-1-(aminomethyl)-3,4-dihydro-5,6-dihydroxy-3-phenyl-1H-2-benzopyran (A68930, 50 µM).
To examine the effects of DA on stimulus-evoked postsynaptic potentials, a concentric stimulating electrode (FHC, Bowdoinham, ME) was placed in the white matter tract immediately adjacent to the recorded neuron, and close to the fiber tract of the external capsule immediately adjacent to the BLA. Postsynaptic currents were elicited orthodromically using 5 low intensity 150 µs square-wave pulses administered once every 5 s. Unless stated otherwise, all synaptic data shown are the average of 5 consecutive stimuli.
Miniature spontaneous excitatory postsynaptic currents (mEPSCs) were recorded at a holding potential of −60 mV in the presence of tetrodotoxin (TTX, 1 µM) and 6-imino-3-(4-methoxyphenyl)-1(6H)-pyridazinebutanoic acid hydrobromide (SR 95531; Gabazine, 5 µM) to block fast GABAA receptor-mediated inhibitory transmission. All drugs were purchased from Sigma-Aldrich (St. Louis, MO), except for A68930 that was purchased from Tocris Cookson Inc, (Ellisville, MO). Drugs were applied by gravity perfusion at the required concentration in the circulating ACSF.
After obtaining the basic electrophysiological properties of recorded neurons, the cell cytoplasm was aspirated into the recording pipette under visual control, by applying gentle negative pressure. The capillary glass used for making the pipette was previously autoclaved and the pipette contained RNase free patch solution. After aspiration of the neuron the electrode tip was broken off and the contents were expelled into an eppendorf tube containing 1.5 µl of cell lysis buffer [1XPCR buffer II (Applied Biosystems, Foster City, CA), 1.5 mM MgCl2 (Applied Biosystems), 0.5% NP40 (USB corporation, Cleveland, OH), 5 mM DTT (Invitrogen, Carlsbad, CA), 50U/µl RNase Inhibitor (Invitrogen), 10 ng/µl primer V1(dT)24 and 2.5 mM dNTP mix (Invitrogen)] by applying positive pressure. After 15s centrifugation, the tubes was incubated in water bath at 70°C for 90s, and then immediately put on ice for 1 min. A 0.3µl volume of Reverse Transcriptase (RT) mixture [200 U/µl SuperScript III (Invitrogen), 50 U/µl RNase Inhibitor and 6 µg/µl T4 gene 32 protein (Roche, Basel, Switzerland)] was added to each reaction tube, which were then incubated at 50°C for 5 min, followed by heat-inactivation at 70°C for 10 min. The tubes were immediately put on ice for 1 min, and after 15 s centrifugation, 1 µl of Exonuclease I mixture [1X Exonuclease I buffer and 0.5 U/µl Exonuclease I (NewEngland BioLabs, Ipswich, MA)] was added to each tube. The reaction mixture was then incubated at 37°C for 30 min, heat-inactivated at 80°C for 25 min, and then placed on ice for 1 min. Next, 6 µl of terminal deoxynucleotidyl transferase (TdT) mixture [1X PCR buffer II, 25 mM MgCl2 (Qiagen Germantown, MD), 100 mM dATP (Invitrogen), 2 U/µl RNaseH (Invitrogen) and 15 U/µl TdT (Invitrogen)] was added to each tube, and the mixture was incubated at 37°C for 15 min followed by heat-inactivation at 70°C for 10 min. The RT product was stored at −20°C before further processing. The RT product in each tube (~13 µl) was divided into four 0.2 ml thin walled PCR tubes (3 µl each). Then, 19 µl of PCR mixture I [1X Buffer (Qiagen), 10 mM dNTP mix, 1 µg/µl primer V3 (dT)24 and 0.05 U/µl Taq DNA Polymerase (Qiagen)] was added to each tube for the first round of PCR: 95 °C for 3 min, 50°C for 2 min and 72°C for 3 min. The tubes were immediately put on ice for 1 min, and 19 µl of PCR mixture II was added, with a composition similar to that of PCR buffer I except that primer V3 (dT)24 was replaced with the primer V1 (dT)24. The sequence of V1 (dT)24 and V3(dT)24 primer was 5’ATA TGG ATC CGG CGC GCC GTC GAC TTT TTT TTT TTT TTT TTT TTT TTT-3’ and 5’-ATA TCT CGA GGG CGC GCC GGA TCC TTT TTT TTT TTT TTT TTT TTT TTT-3’ (Kurimoto et al., 2006). A 20-cycle PCR amplification was then performed 95 °C for 30 s, 67°C for 1 min and 72°C for 3 min with a 6 s extension per cycle using PTC-200 Peltier thermal cycler (MJ Research). The amplified cDNA from each cell was screened for the expression of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as a positive control. The amplified cDNA from GAPDH positive cells was subjected to another amplification step using 2 µl of cDNA from each cell as a template and 100 nM of each of the primers. All the primers described in this paper were purchased from GenBank sequences with commercially available Oligo software (IDT Tools, Coralville, IA). The following primers were used: GAPDH, 5’-GCC ATC AAC GAC CCC TTC AT-3’and 5’-TTC ACA CCC ATC ACA AAC AT-3’ (315 bp product; GenBank accession no M17701); D1R-5’-CAG TCC ATG CCA AGA ATT GCC AGA-3’ and 5’ – AAT CGA TGC AGA ATG GCT GGG TCT-3’ (225bp product; GenBank accession no M35077); D5R-5’-AGT CGT GGA GCC TAT GAA CCT GAC-3’ and 5’- GCG TCG TTG GAG AGA TTT GAG ACA-3’ (517bp product; GenBank accession no M69118). The PCR master mix was comprised of 10X PCR Buffer, 3 mM MgCl2, 10 mM dNTPs, 2.5 U of Taq DNA Polymerase in a final volume of 20 µl. PCR was performed using a 10 min hot start at 95°C followed by a 40 cycle program for GAPDH and D1 (94 °C for 40 s, 56°C for 40 s and 72°C for 1 min), for D5 (94 °C for 1 min, 58°C for 1 min and 72°C for 1.5 min). PCR products were visualized by staining with ethidium bromide and separated by electrophoresis in 1% agarose gel.
D1-immunoreactivity (-IR) is present throughout the amygdala but is heterogeneously distributed (Fig. 1A and B). D1-IR is robust in the BLA and lateral subdivisions of the amygdala, is found at moderate levels in the basomedial subdivision and is relatively weak in the central nucleus. Of particular note, there are densely stained patches of D1-IR found between the subdivisions of the amygdala, especially between the central nucleus and the more ventral regions. These appear to be analogous to the intercalated cell groups identified in rat amygdala and which stain intensely for D1 (Fuxe et al., 2003). D5-IR is also present throughout the amygdala but appears more homogeneous than D1, though the central nucleus is more lightly labeled than the rest of the structure (Fig. 1C and D). When examined at higher magnification, D1-IR in the BLA is most intense in a perinuclear reticular network in cell bodies (Fig. 2A), which has previously been associated with labeling in the Golgi apparatus (Muly et al., 1998; Bordelon-Glausier et al., 2008). In addition, D1-IR can also be identified within the soma outside of this reticular network as well as in the neuropil where a light, granular or punctate pattern can be identified. Within the dense patches of D1-IR interposed between the central nucleus and ventral amygdala, neuropil staining is more intense and immunoreactive fibers can be identified (Fig. 2B). D5-IR in the BLA is found in cell bodies and in the neuropil with both puncta and processes labeled (Fig. 2C).
We next examined the localization of D1Rs in the primate BLA in material prepared for electron microscopy. As anticipated from the light microscopy examination, both D1- and D5-IR are found in neuronal cell bodies in the BLA. As previously reported in the prefrontal cortex, somatic D1-IR is concentrated in the Golgi apparatus of neurons (Fig. 3A); however, unlike cortical neurons, D1-IR is also found associated with other internal membranes in BLA neurons (Fig. 3B). In contrast with D1-, D5-IR is not localized to Golgi complexes, but rather is found associated with other internal membranes (Fig. 3C). D5-IR can also be observed in proximity to the plasma membrane, occasionally adjacent to subsurface membrane structures (Fig. 3C), analogous to previous reports in the prefrontal cortex (Paspalas and Goldman-Rakic, 2004).
The bulk of immunoreactivity for both D1R subtypes is found in the neuropil and both were widely distributed. Both D1- and D5-IR were commonly seen in dendritic structures, including spines (Fig. 4A and B) and dendritic shafts (Fig. 4C–E). These immunoreactive shafts included dendrites that received asymmetric (Fig. 4D) and symmetric synapses (Fig. 4E). Glial processes were also observed with label for both D1 and D5 (Fig. 4F). In addition, both D1- and D5-IR were seen in preterminal axons and axon terminals (Fig. 5). Both D1Rs were found in a diverse group of terminals, including those making asymmetric (Fig. 5A and B) and symmetric synapses (Fig. 5C) as well as those containing prominent dense core vesicles (Fig. 5B). In addition, D1-IR terminals were occasionally observed making asymmetric synapses onto D1-IR dendritic spines (Fig. 5D and E). In order to determine if the patterns of neuropil distribution differed for the two D1R receptors, we quantified their distribution in the BLA. We examined randomly collected images of neuropil labeling for each receptor and identified 918 D1-IR profiles and 903 D5-IR profiles. The patterns of D1 and D5 labeling confirmed our impression of widespread distributions and differed significantly (Fig. 6; χ2= 79.679; p<0.0001). Post-hoc testing revealed that D1 immunoreactivity was more commonly found in dendritic shafts, while D5 immunoreactivity was more commonly found in axon terminals and preterminal axons.
Our previous work in rhesus monkey prefrontal cortex has indicated an extensive overlap of D1 and D5 receptors in populations of dendritic spines and axon terminals (Bordelon-Glausier et al., 2008). Because both receptors are found commonly in these structures in the BLA as well, we used double-label pre-embedding immunogold and immunoperoxidase methods to determine if the two receptors colocalized in the BLA. While the signal obtained with the pre-embedding immunogold method is weaker than that seen with immunoperoxidase, we did achieve specific labeling, as evidenced by D1 gold labeling in Golgi complexes (Fig. 7A). Double labeled dendrites were frequently observed, regardless of which receptor was labeled with gold (Fig. 7B and C). Gold signal in spines and terminals was weaker and sometimes difficult to distinguish from background staining; however, examples of D1/D5 double labeled spines and terminals were observed (Fig. 7D–G).
In order to quantify the degree to which the two receptors co-localize in spines and axon terminals, we used a cocktail labeling approach which we have previously used to identify overlapping distributions of proteins (Muly et al., 2001; Bordelon-Glausier et al., 2008). The advantage of this method is that the labeling method used for each receptor has the same sensitivity and penetration into tissue, thus minimizing the effect of false negative labeling which is problematic for pre-embedding immunogold labeling. We randomly imaged material labeled with antiserum to D1, D5 or a cocktail of antisera to both receptors and calculated the percentage of spines and terminals labeled for each receptor individually as well as for the two receptors combined. We found that 24.1% +/−1.3% of spines in the BLA contained D1-IR, while 18.0% +/− 1.4% contained D5-IR. The percentage of spines in the BLA labeled individually for the two receptors or the cocktail of both antisera differed significantly (Fig. 8A; F2,1687 = 5.729, p = .0033), and post-hoc Scheffe tests confirmed that the percentage of spines labeled for D5 was significantly less than for D1 (p = .0069) and the cocktail (p = .0226). However, there was no significant difference between the percentage of spines labeled for D1 or the cocktail of D1 and D5 (p = .9377). The finding that the percentage of spines labeled by D1 and a cocktail of D1 and D5 is not significantly different demonstrates that the D5 receptor is found in a subpopulation of the D1 positive spines. If D1 and D5 labeled separate populations of spines, the cocktail condition would label a higher percentage of spines than D1 alone. This data indicates that both D1 and D5 are found together in approximately 18% of dendritic spines, and that D1 is found in an additional 6% of BLA spines.
The extensive overlap of D1 and D5 in dendritic spines suggests that individual BLA projection neurons must co-express these receptors. We tested this hypothesis using single-cell RT-PCR on samples obtained from primate BLA projection neurons recorded in in vitro slices (Fig. 9). Here, we looked for the presence of D1 and D5 mRNA in a representative sample of 19 primate projection neurons using available primers based on the rat sequence for each gene. We were able to identify the mRNA for a housekeeping gene, glyceraldehydes-3-phosphate dehydrogenase (GAPDH), in 15/19 neurons and only these neurons were then tested for their expression of D1 and D5 mRNA. An mRNA signal for either D1 or D5 was detected in 13/15 GAPDH positive samples. An mRNA signal was detected in nine neurons each for D1 and D5. Significantly, 5/9 neurons had mRNA transcripts for both D1 and D5 receptors. These results strongly support our hypothesis that individual BLA projection neurons express both D1 and D5 receptors. Moreover, given the variable sensitivity of RT-PCR and our use of primers from rat, the degree of co-expression observed with this method is likely an underestimate of D1 and D5 co-expression.
We also performed a double-label cocktail analysis for axon terminals in the BLA, as we had for dendritic spines. We found that 3.6% +/− 0.3% of terminals in the BLA contained D1-IR, while 6.4% +/− 0.4% contained D5-IR. As seen for dendritic spine labeling, the percentage of axon terminals labeled for D1, D5 or the cocktail differed significantly (Fig. 8B; F2,2021 = 15.683, p<.0001), and post-hoc Scheffe tests confirmed that the percentage D1 labeled terminals was significantly less than D5 (p < .0001) and the cocktail (p < .0001); however, there was no significant difference between D5 and the cocktail (p = .8220). Thus, as in spines there is extensive overlap between the populations of terminals that contain the two receptors. Unlike spines, in terminals, D5 is found more frequently and our results indicate that both D1 and D5 are found together in approximately 3.5% of axon terminals, and that D5 is found in an additional 3% of BLA terminals
We next examined the synaptic type and postsynaptic structures of D1R-labeled axon terminals. Of the 26 D1 positive axon terminals which made identifiable synapses, 22 were asymmetric and four were symmetric. Sixteen of the asymmetric synapses were onto spines, while the remaining six were onto unlabeled dendrites. Three of the symmetric synapses were onto dendrites and the remaining D1 positive axon terminal formed a symmetric synapse onto an unlabeled spine. Of the 29 D5 positive axon terminals which made identifiable synapses, 22 were asymmetric and seven were symmetric. Twenty of the asymmetric synapses were onto spines, while the remaining two were onto dendrites. The symmetric synapses were all onto dendrites. In addition, dense core vesicles were frequently observed both in D1-IR (29.9%) and D5-IR (27.3%) terminals. These data suggest that D1R are positioned to presynaptically modulate a wide variety of inputs, in particular excitatory axospinous inputs.
To determine how D1 family receptor activation might modulate neuronal activity in the primate basolateral amygdala (BLA) we used in vitro whole cell patch-clamp recording from BLA projection neurons to examine the response to exogenous application of either dopamine or the D1R–selective agonist, A68930. To date no study has reported on the basic membrane properties of neurons in the primate BLA. In order to make a direct comparison between the dopamine-evoked response in rodent and primate projection neurons we first characterized some of the basic membrane properties of projection neurons in the primate BLA, a more detailed analysis of the membrane properties of these neurons is in preparation. As illustrated in Figure 10, the voltage response (upper trace) of primate BLA projection neurons to graded injection of either hyperpolarizing or depolarizing current (lower trace) is qualitatively no different from that previously reported for rodent BLA projection neurons (Rainnie et al., 1993; Sah et al., 2003; McDonald et al., 2005). In response to depolarizing current injection, primate projection neurons often fired an initial doublet of action potentials and then settled down to a relatively rhythmic firing pattern. In the hyperpolarizing direction, voltage deflections of increasing amplitude were characterized by the presence of a time-dependent depolarizing sag that increased in amplitude as the membrane potential became more negative. This depolarizing sag is characteristic of the voltage response seen in rodent BLA projection neurons following activation of the hyperpolarization-activated non-selective cation current, Ih.
Note the occurrence of spontaneous IPSPs in the voltage traces above −60 mV (Fig. 10), which reverse polarity at more negative membrane potentials. Similar to the high amplitude and long duration spontaneous IPSPs observed in rodent projections neurons that are mediated by GABAA receptor activation (Rainnie, 1999), the spontaneous IPSPs in primate projection neurons occur at a frequency of ~ 1 Hz and were blocked by prior application of the selective GABAA receptor antagonist, bicuculline methiodide (30 µM, data not shown).
An examination of the resting membrane properties and spike characteristics of revealed significant differences in some, but not all, of the properties of primate BLA projection neurons compared to those of a representative sample of rodent projection neurons (Table 1). Primate projection neurons had a significantly lower membrane input resistance (Rm; Primate = 39±5 MΩ: Rat = 71±3 MΩ; p < 0.01), which is consistent with them having a greater somatic surface area and a more extensive dendritic arbor. No significant difference was observed in the threshold for action potential generation, and either the rise-time or amplitude of single action potentials. However, the primate action potential half-width was significantly longer than that of the rodent (Primate = 1.1 ± 0.1 ms; Rat = 0.89 ± 0.1 ms; p < 0.01), which is consistent with the significantly slower decay time seen in primate projection neurons (Primate = 1.5 ± 0.1 ms: Rat = 0.92 ± 0.1 ms; p < 0.01). Taken together, these data suggested that primate projection neurons expressed many of the same intrinsic membrane currents that are found in projection neurons from the rat BLA, with some differences observed that are likely attributable to the larger size of primate neurons.
Given the similarities in membrane response properties between primate and rat BLA neurons, we predicted that exogenous dopamine would affect the excitability of primate BLA projection neurons in much the same way as it affects rat projection neurons. Consistent with our prediction, application of dopamine (50 µM) caused a reversible 3.8±0.2 mV depolarization of the resting membrane potential of primate projection neurons that was associated with a significant reduction (21%) in the membrane input resistance (Fig. 11 A; Control: 35±7 MΩ; Dopamine 27±7 MΩ, n = 10, p < 0.05). In voltage clamp mode (Fig. 11B), exogenous dopamine application was seen to elicit an inward current (59 ± 5.6 pA, n=17) that persisted in the presence of tetrodotoxin (TTX, 1 µM), and which had an apparent reversal potential of −39 ± 2.1 mV (n=17). These data are consistent with previous studies in the rat BLA in which a similar dopamine-evoked inward current observed in projection neurons was mimicked by the D1R selective agonist, dihydrexidine, and blocked by the D1R selective antagonist SCH 23390 (Levita et al., 2003; Pickel et al., 2006).
Moreover, in control aCSF application of dopamine (50 µM) routinely caused a doubling in the frequency of the spontaneous IPSPs in primate projections neurons (Fig. 12). Hence, in aCSF spontaneous GABAA receptor-mediated IPSPs occurred at a frequency of 0.85±0.2 Hz (Fig. 12A, n = 8). In the presence of dopamine (50 µM) the frequency of spontaneous IPSPs increased to 1.65± 0.3 Hz (Fig. 12 B, n = 8, p<0.01), and returned back to control values (0.7 ± 0. 4 Hz) following washout with aCSF (Fig 12C, n = 8). A similar increase in spontaneous IPSP frequency has been reported in projection neurons of the rat BLA (Levita et al., 2003; Kroner et al., 2005) and has been shown to result from a direct D1R–mediated excitation of local circuit interneurons.
Our localization studies found both D1 and D5 receptors present in axon terminals making asymmetric contacts with the spines of primate BLA projection neurons, similar to observations of the D1 receptor in the rat BLA (Pickel et al., 2006). Consequently, we next examined the effects of exogenous application of the D1R selective agonist, A68930 (50 µM), on the amplitude and frequency of miniature excitatory postsynaptic currents (mEPSCs) recorded in primate projection neurons in the presence of TTX and GABA receptor antagonists. As illustrated in Figure 13, application of A68930 increased the frequency (control 4.8±1.1 Hz; A68930 6.8±1.7 Hz; n=6, p<0.05), but not the amplitude (control 14.6±0.6 pA; A68930 14.3±0.9 pA; n=6, p>0.05), of mEPSCs in primate projection neurons, suggesting a presynaptic site of D1R action. Prior application of the non-selective D1R antagonist (SCH23390, 20 µM), which has no effect on either the amplitude or the frequency of mEPSCs by itself, fully blocked the A69830 induced increase in mEPSC frequency (SCH 23390 3.7±0.7 Hz; A68930 in SCH23390 3.3±0.6 Hz, n=6, p>0.05; Fig. 14).
Because of the similarities between what we observed of the actions of dopamine and D1R stimulation in primate BLA and what has previously been reported in rat BLA, we wanted to determine how the distribution of D1R in rat BLA compared to primate. We performed immunohistochemistry for both D1 and D5 in rat brain sections and quantified the distribution of these receptors in BLA. We examined randomly collected images of neuropil labeling for each receptor and identified 526 D1-IR profiles and 451 D5-IR profiles. Both D1 and D5 were widely distributed within rat BLA (Fig. 15), as we found in primate BLA. The distributions of the two receptors differed in rat BLA (χ2= 72.371; p<.0001). Post-hoc testing revealed that D1 immunoreactivity was more commonly found in dendritic spines, while D5 immunoreactivity was more commonly found in axon terminals and glia. When we contrast the distribution of each receptor between the two species, in the rat D1 is found more frequently in spines and preterminal axons and less commonly in dendritic shafts than we observed in primate while D5 is found more frequently in dendrites and glia in rat and less frequently in spines compared to primate. On the other hand, the distribution of D1R in the BLA of the two species shares some significant similarities, including relatively more presynaptic localization of the D5 receptor and relatively more postsynaptic localization of the D1 receptor.
Dopamine activation of D1 family dopamine receptors plays a critical role in amygdala functioning. Dopamine levels are increased in the amygdala by conditioned fear stress (Yokoyama et al., 2005) and D1R activation in the BLA is necessary for the development of conditioned fear (Lamont and Kokkinidis, 1998; Nader and LeDoux, 1999; Guarraci et al., 1999; Greba and Kokkinidis, 2000; Macedo et al., 2007). The data presented here provide the first detailed description of the distribution of D1 family dopamine receptors in the primate BLA. In addition, we present physiological evidence for dopamine actions in the primate BLA that are similar to those reported in the rat. Finally, we contrast the distribution of D1 and D5 in the rat and monkey.
Our results demonstrate a complex and widespread distribution of both D1 and D5 receptors in primate BLA with both receptors found both postsynaptically in the dendritic arbor of BLA neurons and presynaptically in axon terminals and preterminal axons. Our electrophysiological data confirmed that dopamine and D1R stimulation can directly depolarize and excite both BLA projection neurons and interneurons, as well as act presynaptically in increase neurotransmitter release from glutamatergic terminals. Furthermore, while previous studies have highlighted D1 localization in intercalated cell masses, our finding that one quarter of dendritic spines in the BLA contain either D1 or D1 and D5 show that dopamine acting on these receptors will have powerful effects by directly modulating the excitability of BLA projection neurons. Thus, dopamine’s actions in primate amygdala are a complex mix of post- and presynaptic actions at several sites in intra-amygdala circuits.
The finding that the D5 receptor is found in significant numbers of BLA spines, though the D1 receptor is more abundant is significant given the observation that D1R agonists do not appear to couple to adenylate cyclase the amygdala (Leonard et al., 2003), but rather couple to phosphoinositol hydrolysis (Undie and Friedman, 1990). Studies from transgenic mice have suggested that D1R coupling to phospholipase C is mediated by the D5, but not the D1 receptor (Friedman et al., 1997; Sahu et al., 2008). Our study shows significant D1 receptor localization in both monkey and rat BLA, thus the alternate coupling observed in the amygdala is not solely due to a different D1R subtype present there. It remains to be determined if the different D1R signaling in the amygdala is due to different availability of signal transduction proteins there or possible heterodimerization of D1R subtypes with each other or another receptor.
Our comparison of the distributions of the D1 and D5 receptors in monkey BLA shows patterns that are in general similar but with some significant differences. D1 is found more frequently in dendrites than D5, while D5 is more frequently found in presynaptic compartments. The two D1R subtypes are co-expressed in BLA projection neurons and overlap extensively in spines and terminals but with D1 predominant in spines while D5 is predominant in terminals. The significance of this finding depends on the functional differences between the two D1R subtypes. Efforts to determine these differences have been hampered by a lack of pharmacological tools that adequately differentiate the two receptors. Despite this, a growing body of evidence points to important differences between the two. Investigators using transgenic mice or antisense oligonucleotide injections have found that D5, but not D1 receptors, are necessary for cocaine discrimination as well as dopamine-evoked acetylcholine release in the hippocampus (Filip et al., 2000; Hersi et al., 2000; Laplante et al., 2004). Furthermore, D1 directly interacts with the NMDA receptor (Lee et al., 2002), while D5 directly interacts with the GABAA receptor (Liu et al., 2000). Intriguingly, the two D1R subtypes appear to play differential roles in modulating synaptic plasticity. Corticostriatal long-term potentiation is disrupted in D1 knockout mice, while blockade of the remaining D5 receptors prevents induction of long-term depression (Centonze et al., 2003). In particular, the D5 receptor has a 10 fold higher affinity for dopamine, the endogenous ligand (Sunahara et al., 1991; Weinshank et al., 1991). This observation, coupled with the D5 receptor’s relatively preferential localization in presynaptic compartments suggests that in the BLA presynaptic D1R effects might dominate at low levels of dopamine release and that as the activity of dopaminergic inputs increased, postsynaptic actions might become more prominent.
This study also allowed us to compare the distributions of D1R subtypes between two species, the rhesus monkey and rat. Previously, only the localization of the D1 subtype in rat amygdala has been examined qualitatively and with somewhat divergent results. Pickel and colleagues (2006) reported D1-IR mainly in cell bodies and dendrites, with spines and axon terminals seen more rarely. Alternately, Pinto and Sesack (2008) reported D1-IR was most commonly seen in dendritic spines, with dendritic shafts, axonal elements and glia less commonly seen. Our data confirmed that label is more common in the dendritic arbor than the axon arbor with both spines and dendrites being commonly labeled, with presynaptic and glial localization relatively less common. In comparing the localization profiles we observed in monkey and rat, there were significant similarities noted. Both receptors were widely distributed in both species. Labeling was more common in the dendritic arbor for D1 in both species and in the axonal arbor for D5 in both species. As discussed above, D5 has a higher affinity for dopamine than D1 and our results in rat and monkey suggest that presynaptic action at a low concentration of dopamine is an important and conserved aspect of dopaminergic regulation of the BLA. There were some differences between the two species, in particular D1 was found more frequently in spines and less so in dendrites in rats, while D5 was found less frequently in spines and more so in dendrites in rats compared to monkeys. Our physiological data in monkey was notable for its similarities with previous studies in rat. In future studies it will be interesting to examine whether there are any major differences in the postsynaptic responses to dopamine between the two species.
This work was supported by a Merit Award from the Office of Research and Development, Department of Veterans Affairs to ECM; MH069852 from the National Institutes of Health to DGR and BFU 2006-00306 from the Ministerio de Educacion y Ciencia to ZUK; and the Center for Behavioral Neuroscience (NSF agreement IBN-9876754), and by an NIH/NCRR base grant (P51RR000165) to Yerkes National Primate Research Center.