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Laboratory for Molecular Neurobiology, Clarke Institute of Psychiatry, and the Departments of Psychiatry and Pharmacology, and Institute of Medical Sciences, University of Toronto, Toronto, Ontario, Canada
Address all correspondence and requests for reprints to: Dr. H. H. M. Van Tol, Laboratory of Molecular Neurobiology, Clarke Institute of Psychiatry, 250 College Street, Toronto, Ontario, Canada M5T 1R8. E-mail hubert.van.tol{at}utoronto.ca
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Studies concerning dopaminergic control of PRL synthesis have centered around the hormonal control of PRL gene transcription, especially through promoter activity and regulation studies. Regions of the PRL promoter necessary for positive regulation of PRL gene transcription are clustered in two 5'-regions, one proximal (position -400 to -30) and the other distal (-1.8 to -1.5 kilobases) to the transcription initiation site (reviewed in Ref.5). Negative regulation of PRL gene expression by dopamine is thought to occur mainly through the proximal promoter region. It is generally thought that dopaminergic suppression of PRL gene transcription in pituitary lactotrophs is mediated by the coupling of D2 receptors to the inhibition of adenylyl cyclase (5). In support of this model is the observation that dopaminergic suppression of PRL gene transcription is reversed by the addition of cAMP analogs (7). Additionally, D2 antagonists that prevent the dopamine-dependent decrease in cAMP have been found to block PRL promoter regulation by dopamine in D2short-expressing GH4ZR7 cells (5). However, there is also considerable evidence for regulation of PRL transcription by cAMP-independent pathways as seen by extracellular K+ stimulation and [Ca2+]i changes (5, 8). It has further been shown that the D2 couples to cAMP inhibition and [Ca2+]i changes through distinct G protein subunits, and these seconds messengers, in turn, regulate PRL promoter activity through different pathways (9, 10).
Although the D2 receptors are the predominant dopamine receptors found in the anterior pituitary, there has also been evidence for D4 receptor messenger RNA (mRNA) in the anterior pituitary (11, 12). Moreover, there has been evidence of dopamine receptors other than D2 that might be involved in PRL secretion (13). D4 receptors are structurally, pharmacologically, and functionally very similar to the D2 receptors, and like D2 receptors, D4 receptors have been found to inhibit adenylyl cyclase activity. Therefore, it is possible that D4 receptors may also play a role in the regulation of PRL. In the current study we investigated a possible role of D4 in regulating PRL promoter activity and PRL secretion in GH4C1 cells. For this, we created and characterized the pharmacological profile of several GH4C1 cell lines stably transfected with D4 receptor variants containing two, four, or seven 48-bp repeat units in its putative third cytoplasmic loop. We determined the functional activity of the D4 receptor in these cell lines with respect to adenylyl cyclase inhibition, the ability to regulate cAMP response element (CRE)-directed lacZ expression (14), and the effect of D4 receptor stimulation on PRL promoter activity and VIP-stimulated PRL secretion. This work demonstrates that D4 activation can cause a decrease in cAMP levels in GH4C1 cells and can negatively regulate CRE-directed gene transcription, but does not affect PRL synthesis or secretion.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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200 fmol/mg protein or higher
were selected for our study. Stable
GH4C1 cell lines expressing
any one of the D4 receptor variants were grown under constant selection
pressure using medium supplemented with 200 µg/ml G418. A
GH4ZR7 cell line expressing the D2 receptor
(short isoform) was obtained from Dr. P. Albert (McGill University,
Montreal, Canada) and was cultured in a similar manner as the
D4-expressing cell lines.
Radioligand receptor binding
The transfected cells were homogenized (Polytron, Brinkmann
Instruments, Westbury, NY; setting 5, 5 sec) at 4 C in binding buffer
[50 mM Tris-HCl (pH 7.4), 5 mM EDTA, 1.5
mM CaCl2, 5 mM MgCl2, 5
mM KCl, and 120 mM NaCl]. The homogenates were
centrifuged for 15 min at 39,000 x g, and the pellets
were resuspended in binding buffer at a concentration of approximately
1 mg/ml. For saturation binding analysis, 250 µl homogenate were
incubated in duplicate with increasing concentrations (10–3000
pM) of [3H]spiperone (120 µCi/mmol).
Competition binding analysis was performed by coincubation of 200–250
pM [3H]spiperone and increasing
concentrations (10–3000 pM) of the competing ligand in
either the presence or absence of 200 µM
guanilyl-imido-diphosphate (Gpp(NH)p). Nonspecific binding was
determined by coincubation of [3H]spiperone with 30
µM dopamine. The samples were incubated in a final volume
of 1 ml for 2 h at room temperature and then filtered using a cell
harvester (Skatron Instruments, Lier, Norway). Radioligand bound to the
filters was detected by liquid scintillation counting (Packard 4660
scintillation spectrometer, Downers Grove, IL). The density of
[3H]ligand-binding sites (Bmax) and
dissociation constants of ligands (Kd) were determined by
Scatchard analysis. The ligand binding data were analyzed by the
nonlinear least squares curve-fitting program Ligand.
cAMP measurements
cAMP measurements were performed according to a previously
described protocol (17). Cells were plated onto six-well, 35-mm dishes
1–2 days before the experiment and grown to 60–80% confluency
(1.0 x 106 cell/plate). These cells were washed
with 1–2 ml HBBS buffer (118 mM NaCl, 4.6 mM
KCl, 1 mM CaCl2, 1 mM
MgCl2, 10 mM D-glucose, 20
mM HEPES, and 0.3 mM isobutyl-1-methylxanthine,
pH 7.2). To determine the potency of agonists, D4-expressing cell lines
were incubated with various concentrations of dopamine or quinpirole
ranging from 0.0001–10 µM in the presence or absence of
either 10 µM forskolin or 100–200 nM VIP.
Control groups received buffer alone or buffer containing only 1
µM dopamine (or quinpirole), 10 µM
forskolin, or 100–200 nM VIP. To determine the potency of
antagonists in blocking dopamine-mediated cAMP changes, cells were
treated with varying concentrations of antagonists in the presence of
dopamine or quinpirole (100 nM) and either of the cAMP
stimulators (10 µM forskolin or 100–200 nM
VIP). The cells were incubated for 30 min at 37 C in a final volume of
1 ml. At the end of the incubation period, the medium was removed by
aspiration, and the cells were harvested in 1 ml permeabilization
buffer [0.05% (vol/vol) Triton X-100 in HBBS buffer]. The samples
were vortexed and spun for 5 min at 13,000 rpm in a microcentrifuge.
The supernatant was collected and frozen at -80 C for cAMP measurement
by RIA.
RNA isolation from rat brain
Brain tissues (striatum, anterior pituitary, cortex, and
hippocampus) were obtained from male Sprague-Dawley rats for RNA
isolation according to a standard procedure outlined previously (18).
Briefly, the isolated brain tissues were separated and homogenized
(Polytron, setting 5, 5 sec) in 4 M filtered guanidium
thiocyanate and layered onto 5.7 M cesium chloride. The
samples were spun (Optima TL Ultracentrifuge, Beckman, Palo Alto, CA)
at 34,000 rpm (in SW41 Beckman rotors, Palo Alto, CA) or 36,000 rpm (in
SW60.1 Beckman rotors) for 12 h at 20 C. The supernatant was
discarded, and the RNA pellet was dissolved in water. An equal volume
of phenol-chloroform (1:1, vol/vol) was added to each sample, vortexed,
and spun at 13,000 rpm for 10 min at 4 C. The aqueous phase was
extracted, and two equivalent volumes of ice-cold 100% ethanol and 3
M sodium acetate, pH 5.2 (10%, vol/vol) were added to it.
The samples were mixed by inversion and chilled on ice for 25 min to
precipitate the RNA. Next, the samples were spun at 13,000 rpm at 4 C
for 25 min, and the ethanol was discarded. The pellets were washed with
70% ethanol, air-dried, dissolved in diethyl pyrocarbonate-treated
water, and stored at -80 C.
Reverse transcription-PCR (RT-PCR) of rat
D4 cDNA
One microgram (1 µg) of total RNA from each brain tissue was
used to reverse transcribe the polyadenylated mRNAs with an
oligo(deoxythymidine) primer according to the manufacturer’s protocol
(Superscript Preamplification System, Life Technologies, Grand Island,
NY). For amplification, two rat dopamine D4
receptor-specific primers were used (5'-GAGAGTCCTGCCGGTGGTAGTT-3' and
5'-TGGTG-TAGATGATGGGGTTGAG-3'; 0.5 µM each) (19).
Ten-microliter samples were denatured for 2 min at 94 C and then
amplified for 20 cycles at 94 C for 5 s, 65 C for 5 s, and 72
C for 15 s using an air thermocycler (1605 Air Thermo-Cycler,
Idaho Technology, Idaho Falls, ID). A final elongation step for 4 min
at 72 C completed the amplification process. From the amplified sample,
1 µl was diluted 10-fold with distilled, deionized water
(ddH2O) and 1 µl diluted sample was reamplified as
described above. The expected length of mRNA-derived rat D4
DNA fragment using these primers was 170 bp. For unprocessed RNA
(i.e. with unspliced introns) or genomic DNA contamination
of total RNA extracts, a 361-bp fragment was expected. Next, DNA from
each PCR reaction was electrophoresed on a 1.5% agarose gel and
transferred to a nylon hybridization filter (Hybond-N, Amersham,
Oakville, Canada). The filters were briefly washed with 2 x SSC
(standard saline citrate), and the DNA was fixed on the membrane by UV
cross-linkage (UV Stratalinker 1800, Stratagene, La Jolla, CA) at 1200
µJ. The filter was prehybridized for 2 h in 0.9 M
NaCl, 0.1% sodium citrate, 0.1% Ficoll (Mr,
400,000), 0.1% polyvinylpyrrolidone (Mr,
40,000), 0.1% BSA (fraction V), and 0.5% SDS at 70 C in a shaker (40
rpm). Next, we probed the filter with a primer that covered the splice
junction-site of intron 3. For this, the filter was removed briefly,
and a [
-32P]ATP end-labeled probe (a rat dopamine
D4 receptor-specific 24-mer oligonucleotide
(5'-TCAGGAAGGCCCCAACTACCACCG-3') was added to the prehybridization
mixture. The filters were then hybridized with this mixture overnight
at 70 C in a shaking incubator. After 12 h, the filter was washed
twice for 5 min each time with 2 x SSC and 0.5% SDS at room
temperature. A final wash of the filter with 1 x SSC and 0.1%
SDS was carried out at 70 C for 5 min. The filters were then subjected
to autoradiography at -70 C using x-ray film (Kodak X-Omat AR5,
Eastman Kodak, Rochester, NY). The amount of radioactivity of
[-32P]ATP-D4-specific probe bound to the
region in the filter that corresponded to a 170-bp D4 DNA
fragment (as detected by ethidium bromide staining of DNA and
autoradiography) was measured by excision of the band and liquid
scintillation counting (Beckman LS6000SC, Palo Alto, CA).
pCRE/lacZ expression studies
To study the effect of D2 or D4 activation on lacZ
expression, GH4C1,
GH4ZR7, D4.2–7, and
D4.4–1 cells (5 x 106) in 0.25 ml Ham’s F-10
medium were electroporated with pCRE/ß-gal plasmid DNA (14) (15 µg
in 20 µl PBS) at 4 C (BTX Electrocell Manipulator 600; 70 µF, 300
V). After electroporation, cuvettes were placed on ice for 5 min. Cells
were then resuspended in medium supplemented with 8% FBS and plated
onto 12 six-well, 35-mm dishes (1.5 ml/plate). Under these conditions,
the viability of cells was high, with more than 80% of the cells
attaching within 1–2 h. Twenty-four hours after transfection, medium
was aspirated, and the cells were incubated for 6 h at 37 C in
fresh medium containing no drugs, 1 µM quinpirole, 10
µM forskolin, 200 nM VIP, a combination of
forskolin and quinpirole, or VIP and quinpirole. After drug incubation,
medium was aspirated, and the cells were scraped into 250 µl Z-buffer
(60 mM NaHPO4, 40 mM
NaH2PO4, 10 mM KCl, and 1
mM MgSO4, pH 7.0), and the level of
lacZ expression was analyzed using a
4-methyl-umbelliferyl-ß-D-galactoside (MUG) fluorometric
assay, as described below.
PRL promoter activity studies
Transient transfection of cells with -422P/Luc construct
to assess PRL promoter-driven luciferase reporter activity was
performed with slight modifications to the protocol outlined previously
(5). Briefly, 5 x 106 cells in 0.25 ml Ham’s F-10
medium were cotransfected by electroporation of plasmid DNA [10 µg
pCMVß (CMV = cytomegalovirus) and 20 µg -422P/Luc in 20 µl
PBS] at 4 C (BTX Electrocell Manipulator 600; 70 µF, 300 V). After
electroporation, cuvettes were placed on ice for 5 min. Cells were then
resuspended in 12 ml medium supplemented with 8% FBS and were plated
onto six-well, 35-mm dishes (1.5 ml/plate). After 24 h, medium was
aspirated, and cells were treated with fresh medium with or without 1
µM quinpirole and/or 1 µM dopamine. After
5-h incubation at 37 C, the medium was aspirated, and cells were
harvested in 0.25 ml harvest buffer [50 mM Tris-MES
(2-[N-morpholino]ethanesulfonic acid), pH 7.8; 1
mM dithiothreitol; and 0.1% Triton X-100] for luciferase
assays and stored at -80 C. One 35-mm well of transfected cells was
harvested in 1 ml Z-buffer for MUG assay, whereas one 35-mm well was
used for ß-galactosidase histochemical staining as outlined below. To
measure luciferase activity, 200 µl cell lysate were combined with 15
µl luciferase cocktail (750 mM Tris-MES, pH 7.8; 15
mM magnesium acetate; and 40 mM ATP) in a
plastic cuvette and placed in the luminometer (BioOrbit 1250,
Pharmacia, Baie d’Urfé, Canada), where 200 µl 1 mM
luciferin dissolved in 5 mM potassium phosphate, pH 7.5,
were automatically dispensed. The peak luminosity was recorded on a
chart recorder attached to the luminometer. Each sample was measured in
duplicate, and appropriate dilutions of sample were made in harvest
buffer to detect luciferase activity in a range of 1–100 light units.
Background was recorded using nontransfected cells assayed in
parallel.
ß-Galactosidase and fluorometric MUG assays
To detect transfection efficiencies, cells were cotransfected
with the reporter vector pCMVß (Clontech Laboratories, Palo Alto,
CA). Transfected cells were rinsed with PBS and fixed with 2%
(vol/vol) formaldehyde and 0.2% (vol/vol) glutaraldehyde in 0.1
M sodium phosphate, pH 7.3, for 5 min at room temperature.
The fixative was aspirated, and cells were washed twice with PBS. The
cells were then overlayed with X-Gal stain (100 mM sodium
phosphate, pH 7.3; 1.3 mM MgCl2; 3
mM potassium ferricyanide; 3 mM potassium
ferrocyanide; and 1 mg/ml 5-bromo-4-chloro-3-indolyl
ß-D-galactoside substrate dissolved in
N',N-dimethylformamide). The fraction of stained
cells was measured after 24 h. Alternatively, cells cotransfected
with pCMVß and harvested in Z-buffer were pipetted in duplicate
(twice, 50 µl) into 96-well microtiter plates and lysed by incubation
with 20 µl 1% (vol/vol) Triton-X 100 for 5–10 min. Next, 30 µl 3
mM MUG substrate were added to the cell extract and
incubated at room temperature for 30 min. Finally, 50 µl STOP buffer
(300 mM glycine and 15 mM EDTA, pH 11.2) were
added. The fluorescence emitted from the sample, diluted to a final
volume of 1 ml in Z-buffer, was read using a fluorometer (Perkin-Elmer
Luminescence Spectrometer L550B, Norwalk, CT; excitation, 350 nm;
emission, 450 nm). A standard curve was generated by measuring the
fluorescence of various dilutions of a sample. A dilution that produced
fluorescence levels in the linear range of the emission curve (usually
between 50- and 500-fold dilution) was used to determine transfection
efficiency.
PRL secretion (RIA) measurement
The effect of D4 stimulation on PRL secretion was assessed
according to a previously described protocol (2). Cells were grown to
60–80% confluency in six-well, 35-mm dishes for 1–3 days before
experiments. Medium was aspirated, and cells were preincubated with
1–2 ml preheated (37 C) FAT (F-10 medium supplemented with 1
mM (-)-ascorbic acid and 20 mM Tris, pH 7.2)
for 5–10 min. This was followed by the addition of 1 ml/well FAT with
or without drugs, and incubation for an additional 30 min at 37 C. VIP
was used to stimulate PRL secretion. The effect of dopamine (1
µM) or quinpirole (1 µM) on basal and
stimulated levels of PRL secretion was assessed in the presence or
absence of VIP (200 nM). After incubation, media were
collected and stored at -80 C.
Rat PRL levels in the medium was determined using a RIA kit supplied by
NIDDK (Bethesda, MD). One hundred microliters of medium containing PRL
secreted from cells were transferred in duplicate to 4-ml Falcon test
tubes (Oxnard, CA). To each tube, 300 µl assay buffer [0.025
M phosphate buffer, pH 7.5; 0.15 M NaCl; 10
mM EDTA; 1% (wt/vol) BSA; 0.1% (wt/vol) NaN3;
and 0.01% (wt/vol) thimerosal] and 100 µl rabbit anti-rat PRL serum
(anti-rPRL-S-9, AFP-131581570; 1:10,000 dilution in assay buffer with
10 mM EDTA) were added. The tubes were vortexed and
incubated at room temperature for 2 h. Next, 100 µl
[125I]rat PRL obtained from Hazelton Corning (Washington,
DC) were added for competition with cold rat PRL present in the medium
(0.0045 µCi/sample). Tubes were vortexed and incubated at room
temperature for 24 h. To precipitate antibody-PRL complex, 200
µl 1% (vol/vol) normal rabbit serum in assay buffer without BSA, 100
µl 0.1 M EDTA (pH 7.5), and 100 µl secondary antibody
(goat anti-rabbit IgG whole molecule R0–881 (Sigma Chemical Co., St.
Louis, MO) were added. The secondary antibody was dissolved in 0.1
M phosphate buffer, pH 7.8, with 0.5% BSA and 0.1%
NaN3 and diluted 5-fold in the same buffer. The tubes were
vortexed and incubated at room temperature for 2 h. Then, each
tube received 750 µl 6% (wt/vol) polyethylene glycol
(Mr, 8000) and was centrifuged at 2000 x
g for 30 min. The supernatant was aspirated, and the amount
of radioactivity in the pellet was determined using a -counter
(Beckman Gamma 5500B, Palo Alto, CA) and converted to a corresponding
amount of rat PRL (rPRL-RP-3, AFP-4459B) from a standard curve with a
range of 0.08–10 ng PRL/100 µl sample volume assayed in parallel.
Total radioactivity was determined from samples containing no PRL (100
µl assay buffer), and background radioactivity was that detected in
samples with no PRL or rabbit anti-rPRL serum (200 µl assay buffer).
The intra- and interassay coefficients of variation were 2.2% and
9.7%, respectively.
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Results |
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Autoradiography revealed a 170-bp band in all samples, which
corresponds to the expected size of the amplified D4 DNA fragment (Fig. 1
). Relative levels of mRNA in the four brain regions,
as determined through scintillation counting (Beckman LS6000SC)
revealed the highest levels of D4 mRNA in the rat anterior pituitary
(279 dpm), followed by cortex (115 dpm), striatum (30 dpm), and
hippocampus (16 dpm). A 361-bp band that varied in intensity with the
levels of 170 bp D4 DNA fragment was also detected in all samples (Fig. 1). This could be due to genomic DNA contamination of total RNA
samples. However, genomic contamination is expected to be similar in
all samples and cannot explain the differences in intensity of the
361-bp band, which varied according to the level of the amplified
170-bp D4 DNA fragment. A more likely explanation would be the presence
of unspliced RNA that might have also been reverse transcribed and
amplified.
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Table 1 summarizes the affinity dissociation constants
(Ki) of various ligands for the D4 variants
expressed in GH4C1 cells.
The Ki values of dopamine alone or in the presence of
Gpp(NH)p for the D4 variants expressed in
GH4C1 cells were very
similar to those observed in Chinese hamster ovary (CHO) cells (17).
Most of the dopamine competition curves (four of six) displayed a
significantly better fit for the two-site model than the one-site model
(P < 0.05). In contrast, all of the dopamine curves
with Gpp(NH)p had a better fit for the one-site model. The
Ki values of antagonists (Table 1) were also comparable to
those previously observed in CHO cells (17) and COS-7 (15). The
pharmacological profile of the antagonists was identical for the three
D4 variants expressed in
GH4C1 (spiperone >
emonapride > haloperidol > clozapine >> raclopride).
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5 pmol/plate) were stimulated about 10-
to 40-fold by 100 nM VIP (Table 2). Incubation with dopamine (1
µM) reduced VIP stimulated cAMP levels by
25–70% in the various D4-expressing cell lines. In a control study,
VIP stimulated cAMP (10-fold) in both untransfected
GH4C1 cells and
D2short-expressing
GH4ZR7 cells, whereas
dopamine inhibited VIP-stimulated cAMP levels completely in
GH4ZR7, but had no effect
in GH4C1 cells (data not
shown). This is consistent with a previous report by Albert et
al. (2). Inhibition of 10 µM forskolin-stimulated
cAMP with dopamine was about 33% in the D4-expressing cell lines and
about 70% in GH4ZR7 cells
(data not shown).
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). No correlation
was detected between VIP stimulation of cAMP and the type of D4
variants studied, the basal level of cAMP, or the receptor density of
the individual cell lines. However, a correlation (r2 =
0.87) was observed between receptor density and the percentage of
dopamine-mediated inhibition of VIP-stimulated cAMP levels (data not
shown). That is, in cell lines with a higher density (Bmax)
of D4 receptors, dopamine was more efficacious in inhibiting
VIP-stimulated cAMP than in cell lines with a lower Bmax.
When the efficacy of dopamine-mediated inhibition of cAMP levels for
the individual cell lines was corrected for receptor density, no
difference was observed between the D4 variants.
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). A similar quinpirole-mediated inhibition of
lacZ gene expression was also observed in these cells after
stimulation of lacZ expression with VIP (200 nM;
data not shown). However, stimulation of lacZ expression
with 10 µM forskolin was typically between 5- to 8-fold
higher than basal levels, whereas VIP (200 nM) only
stimulated lacZ expression about 2- to 3-fold over basal.
Hence, the inhibition of forskolin-stimulated lacZ
expression with quinpirole was more pronounced and consistent than that
seen with VIP (10–100%). This study demonstrates that like D2
receptors, D4 receptors can also negatively regulate CRE-induced
expression of genes.
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). These results are consistent with those of a
previous study by Elsholtz et al. (5), where 72 ± 5%
inhibition on -422P/Luc was reported. We saw no effect of D2 or D4
activation on forskolin-stimulated PRL promoter activity (1.4–4 fold)
in any of the cell lines studied (data not shown).
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). Only the D4.2–7 cell line demonstrated a slight tendency to
inhibit PRL promoter activity (15.6 ± 3.5; n = 5), although
this response was not significantly different from the effects in
control GH4C1 cells. The
ability of D2short receptors to couple to the inhibition of
PRL promoter activity was significantly higher than that of all other
cell lines tested (by one-way ANOVA: F = 14.8; P
< 0.0005; by Student’s t test:
GH4ZR7 vs. any
other cell lines tested, P < 0.005; Fig. 3).
Quinpirole had no effect on 10 µM forskolin-stimulated
PRL promoter activity in any of the cell lines studied (data not
shown). The expression of dopamine receptors was confirmed by a concomitant [3H]spiperone binding assay that revealed a fairly high receptor density for each of the cell lines studied (GH4ZR7, 400 fmol/mg protein; D4.2–7, 548 fmol/mg protein; D4.4–1, 271 fmol/mg protein; D4.7–2, 630 fmol/mg protein). Thus, from this study it can be concluded that the D4 receptors do not significantly alter PRL transcription in GH4C1 cells.
Regulation of PRL secretion by D4 in
GH4C1 cells
To determine whether the D4 variants expressed in
GH4C1 cells have any
modulatory effect on PRL release, the levels of PRL in medium of cells
treated with 200 nM VIP in the presence and
absence of 1 µM quinpirole were measured by RIA
(Table 4). Parental
GH4C1 cells with no
endogenous dopamine receptor were used as a negative control, whereas
the GH4ZR7 cell line stably
expressing D2short served as a positive control. In
both GH4C1 and
GH4ZR7 cell lines, VIP
significantly (P < 0.05, by Student’s t
test) elevated basal PRL secretion [from 3.84 ± 0.97 to
5.87 ± 0.97 ng/plate in
GH4C1 cells (n = 12)
and from 20.9 ± 3.67 to 34.54 ± 2.59 ng/plate in
GH4ZR7 cell (n = 9);
mean ± SE]. In both cases, the VIP-mediated increase
in PRL secretion was approximately 50%, consistent with previous
observations (2). In GH4ZR7
cells, VIP-stimulated PRL secretion was significantly inhibited (36%;
P = 0.003, by Student’s t test) by 1
µM quinpirole, which was also consistent with results of
a previous study using the same cell line (2). However, quinpirole did
not significantly alter VIP-stimulated PRL levels in the parental
GH4C1 cells, confirming
that the effect seen in
GH4ZR7 cells was a specific
response of D2 receptor activation. Quinpirole by itself did not
significantly alter basal levels of PRL secretion in any of the cell
lines.
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) and
extracellular (data not shown) cAMP levels in response to 200
nM VIP. Basal PRL secretion in D4 cells was not
significantly increased in response to 200 nM VIP, although
there was a trend for a VIP-mediated PRL increase (from 4.22 ±
0.8 to 7.12 ± 1.2 ng/plate; n = 9; mean ±
SE) in D4.4–1 cells (P = 0.06, by
Student’s t test). Interestingly, basal levels of PRL
secretion appeared to be inversely correlated to the percentage of
VIP-stimulated PRL secretion in
GH4C1 and
GH4ZR7 cells, but not in
the D4-expressing cell lines. That is, lower basal secretion resulted
in higher VIP-stimulated PRL secretion, but as the basal level
increased, the percentage of VIP-mediated PRL release diminished. |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To date, much emphasis has been placed on investigating the role of the D4 receptor in disorders involving the dopaminergic mesocorticolimbic pathway, such as schizophrenia. This is in part due to studies that revealed a predominantly mesocorticolimbic distribution of this receptor with relatively high levels in the thalamus, hypothalamus, hippocampus, amygdala, nucleus accumbens, globus pallidus, and much of the cerebral cortex (21, 22, 23, 24). In addition, however, using the RT-PCR amplification approach, we found relatively high levels of D4 mRNA in the rat anterior pituitary. The presence of relatively high levels of human D4 mRNA in the pituitary has also been reported by others (11). These observations suggest a potential role for the D4 receptor in the tuberoinfundibular dopaminergic pathway.
It is presently unclear whether D4 receptors are indeed expressed in pituitary lactotrophs or any other pituitary cell type. Specific D4 antibodies or radioligands may shed some light on this issue in the future. If the D4 receptors are expressed in the pituitary lactotrophs, then dopamine released from the tuberoinfundibular dopamine neurons of the hypothalamus might act through these receptors to regulate endocrine functions of these cells, as with D2 receptors.
Our current study reveals that none of the D4 variants stably expressed in the somatomammotrophic GH4C1 cells had a significant effect in modulating PRL promoter (-422P/Luc) activity. Although the D4.2–7 cell line displayed about a 15–20% reduction in basal PRL promoter activity after 1 µM quinpirole treatment, the D2short receptors stably expressed in GH4ZR7 displayed over 50% inhibition of the -422P/Luc activity with 1 µM quinpirole. However, the D4-expressing cell lines could inhibit forskolin- and VIP-stimulated cAMP production by about 33% and 50%, respectively. Moreover, D4-mediated inhibition of cAMP production reversed forskolin- and VIP-induced ß-galactosidase expression driven from a VIP promoter containing five copies of CRE. These studies suggest that the degree to which cAMP is inhibited may be an important factor in determining whether PRL transcription can be repressed by dopamine. This point is further supported by the observation that quinpirole was ineffective in reducing forskolin-stimulated PRL promoter activity even in cell lines (GH4ZR7 and D4.2–7) in which it caused a reduction of basal levels of PRL promoter activity. This may be due to insufficient reduction of cAMP levels in the presence of forskolin to produce an effect at the PRL promoter level. Alternatively, the differential effect of D2 and D4 receptors on PRL transcription may be due to mechanisms other than cAMP inhibition (5).
It has been shown that binding of the pituitary cell-specific transcription factor, Pit-1, to specific 5'-regions of the PRL promoter confers positive regulation of PRL gene expression (reviewed in Ref.25). Negative regulation initiated by D2 receptor activation is thought to involve only the proximal sequences in the PRL promoter (5). It has further been shown that high affinity Pit-1 binding sites can selectively confer dopaminergic regulation, either in PRL-secreting GH4 cells or in a heterologous cell system transfected with a Pit-1 expression vector (10). Although minimal proximal PRL promoter regions containing high affinity Pit-1-binding sites were sufficient for negative dopaminergic regulation through D2short receptors expressed in GH4ZR7, maximal dopamine responsiveness was observed with the -422P/Luc constructs that contained 422 bp of the PRL promoter upstream from the transcription initiation site (5). This might be due to two additional putative regulatory regions found further upstream in the PRL promoter. One (position -97 to -84) is a CRE-like sequence containing 6 of the 8 bp of the TGACGTCA consensus sequence, and the other (position -78 to -71) is a G/C-rich motif resembling an activating protein-2 site (5). There is also evidence for D2 receptor-mediated negative regulation of the Pit-1 promoter itself, which contains two CRE sequences (5). Thus, a D2-mediated reduction in Pit-1 protein expression might be one of the ways in which dopamine regulates PRL synthesis. However, in our study, D4 receptors inhibited cAMP production as well as CRE-regulated transcription, suggesting that even if D4 could modify Pit-1 expression, this is not sufficient to affect Pit-1-dependent regulation of PRL promoter activity.
Normally in the pituitary, dopamine acts via the D2 receptors on lactotrophs to inhibit basal and hormone-stimulated secretion of PRL. Interactions of D2 with pertussis toxin-sensitive G proteins to reduce adenylyl cyclase activity and [Ca2+]i are implicated in the dopamine-mediated inhibition of PRL secretion (3). However, a study by Valerio et al. (13) suggested that PRL release might be regulated at least in part by a dopamine receptor distinct from D2. This was based on the observation that antisense oligonucleotides that knocked down D2 receptors from a primary culture of rat pituitary blocked the bromocriptine-mediated suppression of adenylyl cyclase activity and PRL gene transcription, but not PRL release. This as well as evidence for the presence of D4 mRNA in rat and human pituitary using RT-PCR prompted us to investigate whether human D4 variants could regulate PRL secretion in GH4C1 cells. However, as seen for PRL transcription, we observed no effect of D4 activation on PRL secretion in GH4C1 cells. Nevertheless, the fact that the GH4C1 cells might have lost crucial factors needed for D4-mediated inhibition of PRL secretion cannot be excluded.
Although D4 receptor activation leads to significant reduction of
VIP-stimulated cAMP levels, the efficacy of agonists such as dopamine
and quinpirole is about 2-fold reduced for D4 compared to D2 receptors
in GH4C1 cell lines. Thus,
it might also be possible that near-complete blockade of VIP-stimulated
cAMP levels is required for cAMP-dependent dopaminergic regulation of
PRL synthesis and secretion. However, there is also evidence for
regulation of PRL transcription by cAMP-independent pathways (5, 8).
Studies using antisense oligonucleotides to knock out specific G
subunits (9) or stable transfection of constitutively active mutants of
specific G
subunits (10) have demonstrated the involvement of
distinct G protein subunits in coupling of various second messenger
systems to D2. Alternatively, it cannot be excluded that the D4
receptor might activate additional pathways that would compensate for a
cAMP-mediated reduction of PRL promoter activity.
One of the side-effects of neuroleptic treatment, as for example in schizophrenia, is the occurrence of hyperprolactinemia. Classic neuroleptics such as haloperidol effectively block pituitary D2 receptors and, as a result, cause marked elevations in serum PRL levels. However, this effect is not seen with the so-called atypical neuroleptic clozapine, which displays a 10-fold higher affinity for D4 compared to D2 (26). Moreover, at therapeutic concentrations, clozapine occupies over 75% of the D4 receptors and only 30–60% of the D2 receptors (27, 28). Therefore, based on the presence of D4 receptors in the pituitary, it could be argued that clozapine does not mediate its antipsychotic effects through D2 receptors. However, the inability of D4 receptors to modulate PRL gene transcription and secretion in GH4C1 cells does not provide direct support for the above-mentioned contention.
In conclusion, this work provides support for the existence of the dopamine D4 receptor in the pituitary. However, based on the in vitro studies with the somatomammotrophic cell line GH4C1, the D4 receptor may only slightly modulate PRL synthesis, but not PRL secretion, despite its ability to block cAMP production and to negatively regulate CRE-mediated activity. This work provides evidence that the pituitary expresses D4 receptors that are not coupled to PRL regulation and may represent the D2-like sites reported by Valerio et al. (13)
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2 Career Scientist of the Ontario Ministry of Health.
Received September 6, 1996.
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2- and
Go-mediated signaling in the Pit-1-dependent inhibition of the
prolactin gene promoter. Control of transcription by dopamine D2
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represents one independent duplicate
measurement.