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The Journal of Neuroscience, April 1, 1998, 18(7):2342-2349
Nitric Oxide Depresses GABAA Receptor Function via
Coactivation of cGMP-Dependent Kinase and Phosphodiesterase
Eric M.
Wexler1, 2,
Patric K.
Stanton2, 3, and
Scott
Nawy1, 2
Departments of 1 Ophthalmology and Visual Science,
2 Neuroscience, and 3 Neurology, Albert
Einstein College of Medicine, Bronx, New York 10461
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ABSTRACT |
Nitric oxide (NO) is thought to play an essential role in neuronal
processing, but the downstream mechanisms of its action remain unclear.
We report here that NO analogs reduce GABA-gated currents in cultured
retinal amacrine cells via two distinct, but convergent, cGMP-dependent
pathways. Either extracellular application of the NO-mimetic
S-nitroso-N-acetyl-penicillamine (SNAP)
or intracellular perfusion with cGMP depressed GABA currents. This
depression was partially blocked by a pseudosubstrate peptide inhibitor
of cGMP-dependent protein kinase (PKG), suggesting both PKG-dependent
and independent actions of cGMP. cAMP-dependent protein kinase (PKA) is
known to enhance retinal GABA responses. 8-Bromoinosine 3',5'-cyclic
monophosphate (8Br-cIMP), which activates a type of cGMP-stimulated
phosphodiesterase that hydrolyzes cAMP, also significantly reduced GABA
currents. 1-Methyl-3-isobutylxanthine (IBMX), a nonspecific
phosphodiesterase (PDE) inhibitor, blocked both the action of 8Br-cIMP
and the portion of SNAP-induced depression that was not blocked by PKG
inhibition. Our results suggest that NO depresses retinal
GABAA receptor function by simultaneously upregulating PKG
and downregulating PKA.
Key words:
retina; amacrine; culture; nitric oxide; guanylate
cyclase; ODQ; SNAP; PKG inhibitory peptide
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INTRODUCTION |
Nitric oxide (NO) was first
identified as a factor released by endothelial cells that relaxes
vascular smooth muscle (Palmer et al., 1987 ). Subsequently, its
synthetic enzyme, nitric oxide synthase (NOS/NADPH diaphorase), has
been identified in almost every region of the CNS (Dawson et al.,
1991). As a gas, NO is able to cross cell membranes freely, giving rise
to the notion that NO might act as a retrograde messenger at synapses
from which it is released and possibly at neighboring synapses as
well.
One of the best-documented targets of NO is an intracellular soluble
guanylate cyclase (sGC), the enzyme that synthesizes cGMP (Schmidt et
al., 1993). cGMP can bind to at least three distinct classes of
proteins. First, cGMP can gate channels directly. Cyclic nucleotide-gated channels have been found in cells in both the CNS and
PNS (Fesenko et al., 1985; Nakamura and Gold, 1987; Nawy and Jahr,
1990; Goulding et al., 1992; Schmidt et al., 1993; Bourgeois and Rakic,
1996; Meissirel et al., 1997). Second, cGMP can activate cGMP-dependent
protein kinase (PKG) (Scott, 1991). Finally, two subtypes of
phosphodiesterases (PDE) are known to be regulated by cGMP. Type II PDE
(GS-PDE) is stimulated by binding cGMP, whereas type III PDE (GI-PDE)
is inactivated when cGMP is bound (Beavo et al., 1971a,b).
Cyclic nucleotides are well-established modulators of ion channels in
retinal neurons. Elevated levels of cAMP decrease electrotonic coupling
among both horizontal and amacrine cells (DeVries and Schwartz, 1989,
1992; Mills and Massey, 1995). Similarly, cGMP uncouples teleost
horizontal cells and reduces heterologous coupling between mammalian
bipolar and AII amacrine cells (DeVries and Schwartz, 1989, 1992; Mills
and Massey, 1995). Although both cAMP and cGMP reduce gap junction
coupling, they seem to act antagonistically in regulating the activity
of glutamate receptors. It has been shown that cAMP enhances
glutamate-evoked currents in horizontal cells (Knapp and Dowling, 1987;
Liman et al., 1989). In contrast, cGMP reduces such currents (McMahon
and Ponomareva, 1996).
In retinal neurons, cAMP also enhances GABA-gated chloride currents
(Feigenspan and Bormann, 1994; Veruki and Yeh, 1994). Despite the
recognized importance of NO and the observation that cAMP and cGMP act
on common targets, relatively little is known concerning cGMP-dependent
modulation of GABAA receptor function. However, studies in
the nucleus tractus solitarius (Glaum and Miller, 1993, 1995),
cerebellar granule cells (Robello et al., 1996), and hippocampus (Zarri
et al., 1994) suggest that cGMP may downregulate GABAA
receptor function. Therefore, the present study was undertaken to
examine how cGMP might modulate retinal GABAA receptor
function and to elucidate which enzymes mediate its effects.
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MATERIALS AND METHODS |
Cell culture. Retinas were removed from newborn
Long-Evans-hooded rats after cryoanesthesia and were incubated for
45-60 min at 37°C in DMEM with HEPES (Mediatech, Washington, DC),
supplemented with papain (Worthington, Freehold, NJ) at 6 units/ml and
cysteine at 0.2 mg/ml. Papain was inactivated by replacing the enzyme
solution with medium of the following composition: DMEM plus HEPES,
0.1% mito+ serum extender (Collaborative Research,
Bedford, MA), 5% heat-inactivated fetal calf serum (HIFCS), 0.75%
penicillin-streptomycin-glutamine mix (Life Technologies), and 7.5%
sterile water to lower osmolarity. Retinas were triturated through a
fire-polished Pasteur pipette, plated onto glass coverslips pretreated
with poly-D-lysine (0.1 mg/ml), and maintained in medium
supplemented with 15 mM KCl, 2 ng/ml basic fibroblast
growth factor (bFGF; Life Technologies), and 100 ng/ml brain-derived
neurotrophic factor (BDNF; Regeneron/Amgen). At 72 hr after plating,
cells were treated with the antimitotics 5-fluoro-2-deoxyuridine (0.01 mg/ml) and uridine (0.026 mg/ml) for 24 hr. Subsequently, every 3rd
day, 25% of the culture medium was exchanged for fresh medium. Cells
were used for recording after 10-17 d in vitro.
Immunocytochemistry. Coverslips containing primary cultured
neurons were washed three times with D-PBS, fixed in 4%
paraformaldehyde for 15 min at room temperature and with 100% methanol
at  10°C for 15 min, rinsed with Dulbecco's PBS (D-PBS), and then
incubated overnight at 8°C with primary antibody. Anti-HPC-1 (Sigma,
St. Louis, MO), anti-GABA (Chemicon, Temecula, CA), and
anti-neurofilament 145 (NF145) (Chemicon) were diluted 1:100 in D-PBS,
10% fetal calf serum, and 0.5% Triton X-100. After 12-24 hr,
coverslips were rinsed twice with D-PBS, incubated for 90 min in TRITC-
or FITC-conjugated secondary antibody (Chemicon) diluted 1:100 at room
temperature, rinsed twice in D-PBS and sterile water, and mounted onto
glass slides using Prolong AntiFade (Molecular Probes, Eugene, OR). For
labeling with anti-Thy1.1 IgM (gift of Dr. David Weinstein), live cells
were incubated with primary antibody diluted 1:50 in normal culture
medium for 60 min at 37°C, followed by rinsing twice in D-PBS and
fixing with 4% paraformaldehyde for 5 min at room temperature.
Labeling with secondary antibody was as detailed above. Cells were
viewed with a Zeiss Axiovert 135 microscope equipped with a 100 W
mercury lamp (HBO 100).
Electrophysiology. Recordings were made using an Axopatch
1-D amplifier, Digidata 1200 data acquisition board, and Axobasic software (Axon Instruments). Patch electrodes were pulled from standard
hematocrit tubing (VWR Scientific) using a Narishige PP-83 vertical
two-stage puller and were fire polished (Narishige MF-83) to a
resistance of 2-3 M . Recordings typically had series resistances of
10-20 M, and those that exceeded 20 M or varied by >10% during
the experiment were discarded.
The standard extracellular solution contained 160 mM NaCl,
2 mM CaCl2, and 5 mM HEPES.
The gluconate-based intracellular solution was as follows: 110 mM potassium gluconate, 40 mM KCl, 10 mM EGTA, 5 mM HEPES, 2 mM Mg-ATP,
and 250 µM Na-GTP, pH 7.3. Mannitol was added to adjust
the osmolarity. The phosphate-based intracellular solution contained
potassium phosphate monobasic (120-140 mM). 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one (ODQ)
was stored at 20°C as a 10 mM stock in DMSO.
8-Bromo-cGMP (8Br-cGMP), 8-bromoinosine 3',5'-cyclic monophosphate
(8Br-cIMP), and S-nitroso-N-acetyl-penicillamine (SNAP) were stored frozen (20°C) as a dry powder and were made up
each day in the standard extracellular solution. SNAP was prepared as
an 11% (w/v) stock in DMSO (i.e., 1.1 mg of SNAP/10 µl of DMSO). Sodium nitroprusside (SNP) was made every 3-5 hr. Final DMSO
concentrations never exceeded 0.1%. ATP, GTP, cAMP, cGMP, and
cGMP-dependent protein kinase inhibitory peptide (GKIP;
Arg-Lys-Arg-Ala-Arg-Lys-Glu; Peninsula Labs) were stored at 20°C as
1000× stocks in distilled water. 1-Methyl-3-isobutylxanthine (IBMX)
and microcystin (Research Biochemicals, Natick, MA) were stored at
20°C as 1000× stocks in dry DMSO. Drugs were delivered through a
pair of parallel, fused silica flow pipes (375 or 500 µm inner
diameter: Polymicro Technologies). Each flow pipe was supplied by six
separate reservoirs, each with its own control valve to feed fluid
through a six-to-one tubing manifold (Warner Instruments). The flow
pipe apparatus was under computer control and was driven by a piezo
bimorph actuator (Morgan Matroc). This apparatus is an adaptation of
that used by Lester et al. (1990). With the larger flow pipes, 10-90%
whole-cell solution exchanges were achieved in <2.5 msec.
In experiments in which either cGMP or microcystin was included in the
internal solution, the tip of the recording pipette was filled with
control solution, and the shank was filled with either the cGMP- or
microcystin-containing solution. This delayed the entrance of the test
compound into the cell for several minutes, during which time
ECl was observed to shift from 60 to 30 mV. GABA responses in cells held at 60 mV displayed a "run-up" in amplitude during this period of chloride shifting, a consequence of an
increased chloride driving force. This run-up period was succeeded by
1-3 min of stable, "baseline" GABA responses. Time 0 was defined
as the last point of this baseline period.
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RESULTS |
Identification of amacrine cells
Amacrine cells were distinguished from ganglion and horizontal
cells in culture using both morphological and physiological criteria.
Virtually all of the large multipolar cells >12 µm (280 ± 17 cells/mm2; n = 6 cultures) were
immunoreactive for HPC-1, an amacrine cell marker (Barnstable et al.,
1985). In contrast, <1% of all cells were labeled with medium weight
(145 kDa) neurofilament (NF145), a cytoskeletal protein expressed by
both horizontal cells and ganglion cell axons (Shaw et al., 1984).
Moreover, only 5% (15 ± 4; n = 6) of large (>12
µm) neurons expressed Thy1.1, an antigenic marker for ganglion and
displaced amacrine cells. Fewer than one fifth of these Thy1.1 positive
cells (i.e., 1% of all cells) exhibited a robust, nonpunctate staining
pattern typical of ganglion cells (Taschenberger and Grantyn, 1995).
Taken together, these data suggest that nearly all large cells were
amacrine and not ganglion or horizontal cells. This relative
homogeneity of cell type may have been the result of maintaining the
cultures in a medium of high osmolarity (340-385 mOsm), which has been
reported to prevent ganglion cell survival (Meyer-Franke et al.,
1995).
Figure 1A shows a
typical cell that was immunoreactive for GABA, the neurotransmitter
used by amacrine cells (Versaux-Botteri et al., 1989; Wu, 1992). Cells
with this morphology had membrane potentials of 30 to 40 mV and
rapidly accommodating action potentials, consistent with the
physiological properties of amacrine cells (Taschenberger and Grantyn,
1995). Under whole-cell voltage clamp, all cells of the type shown in
Figure 1A (7-14 d in culture) responded to GABA
(Fig. 1B, overhead bar) with a
desensitizing inward current that reversed at 30 mV, the predicted
ECl, and that was blocked by 100 µM bicuculline, suggesting that these cells expressed
primarily GABAA receptors.
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Figure 1.
Primary culture of postnatal day 0 rat retinal
amacrine cells 7-14 d in vitro. A, A
cultured neuron exhibiting the characteristic amacrine cell morphology
labeled with anti-GABA antiserum. Scale bar, 12 µm. B,
Whole-cell recording of GABA responses from a representative amacrine
cell held at 60 mV. The overhead bar shows the
duration of GABA (35 µM) application. As is
characteristic of GABAA responses, GABA elicited a rapidly
desensitizing inward current that was blocked by the coapplication of
bicuculline methiodide (100 µM). C,
Dose-response relation for the peak GABA currents elicited by 250 msec
applications of varying concentrations of GABA. The averaged data were
fit with the equation I = Imax × (1/[1 + (EC50/[GABA])n]), with an
observed EC50 of 40 ± 2 µM
(n = 8) and a Hill coefficient (N) of 1.8 ± 0.2.
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From the fit of the dose-response relation shown in Figure
1C, we obtained an EC50 of 40 ± 2 µM (n = 8) and a Hill coefficient of
1.8 ± 0.2. Subsequent experiments were performed using 50 µM GABA, a concentration near the EC50 for
these receptors. The Hill coefficient is in good agreement with the
value of 1.9 obtained for GABAA currents in amacrine cells
from rat retinal slice (Feigenspan and Bormann, 1994). The
EC50 for GABA in that study was found to be 72 µM, although agents that promote phosphorylation by
cAMP-dependent protein kinase (PKA) shifted the EC50 to 45 µM (Feigenspan and Bormann, 1994), close to the value
obtained in the present study.
NO agonists depress GABA currents
We examined the potential role of NO in regulating
GABAA receptor-gated currents using two NO analogs, SNP and
SNAP, which activate soluble guanylate cyclase (Bohme et al., 1984;
Schmidt et al., 1993). GABA was applied for 500 msec at intervals of 20 sec, a protocol that avoids cumulative desensitization associated with
longer or more frequent GABA applications. When the control solution
was switched to a solution containing 50 µM SNP, the GABA
response was reversibly reduced. In four cells, SNP reduced GABA
responses by 29 ± 7% (Fig.
2A).
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Figure 2.
Activators of soluble guanylate cyclase depress
GABA responses. A, Single-cell record of the peak inward
currents elicited by 250-500 msec applications of GABA (50 µM) delivered every 20 sec is shown. The overhead
bar indicates the timing of the application of 50 µM SNP. The insets are sample
traces recorded at the times indicated by the
numbers. The internal solution contained phosphate and
10 µM cAMP. SNP reversibly depressed the GABA response by 33% in this cell and by 29 ± 7% (n = 4) on
average. B, SNAP (500 µM) reversibly
depressed the GABA response by 38% in this cell and by 37 ± 2%
(n = 6) on average. C, 8Br-cGMP (1 mM) reversibly depressed the GABA response by 42% in this
cell and by 41 ± 9% (n = 8) on
average.
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SNAP also depressed GABA currents (Fig. 2B).
Application of 50 µM SNAP produced a depression of
24 ± 4%, whereas 100 µM SNAP depressed GABA
currents by an average of 34 ± 4%. One hundred micromolar SNAP
seemed to be near saturating as a fivefold higher concentration
produced little further increase in depression (37 ± 2%). DMSO
(0.1%) alone did not produce any consistent reduction of GABA
responses. Measurement of I-V relations indicated that depression of the GABA response by SNP or SNAP was not because of a
shift in the reversal potential of the response (data not shown). With
continuous application of either SNAP or SNP, recovery of the GABA
response was often observed. The reason for this recovery is not
clear.
If NO agonists depress GABA currents by activating sGC, then direct
application of cGMP should similarly depress them. GABA currents were
elicited as before, and cells were perfused with the membrane-permeant
cGMP analog 8Br-cGMP (Fig. 2C). In eight cells, 1 mM 8Br-cGMP decreased the amplitude of the GABA response by
an average of 41 ± 9%. Thus both cGMP itself and compounds that
elevate intracellular cGMP depressed GABA responses.
To test whether SNAP reduced GABA currents by activating sGC, we first
preincubated cells in 1 µM ODQ, a specific inhibitor of
sGC (Garthwaite et al., 1995). In the presence of ODQ, there was a
20 ± 2% enhancement of the GABA currents, possibly because of
inhibition of a basal guanylate cyclase activity and subsequent reduction in cGMP-mediated depression. Depression of GABA current by
8Br-cGMP was not significantly affected by ODQ, as would be expected if
raising intracellular levels of cGMP directly circumvents the need for
guanylate cyclase activity. In the presence of ODQ, 8Br-cGMP depressed
the GABA response by 35 ± 4% (n = 6; Fig.
3A). ODQ significantly reduced
the depression of GABA current by SNAP at all three concentrations of
SNAP that were tested (Fig. 3B). The efficacies of ODQ
against either a submaximal SNAP concentration (50 µM,
46%) or a near-saturating concentration (500 µM, 44%) were similar. ODQ is reported to block 75% of SNAP-induced sGC activity, measured biochemically, (Garthwaite et al., 1995) compared with the 45% efficacy obtained in this study. The difference suggests that there may be an additional component of SNAP-induced depression of
GABA currents in amacrine cells that is independent of sGC (Pozdnyakov
et al., 1993; Brune et al., 1994; Zoche and Koch, 1995).
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Figure 3.
SNAP depressed GABAA currents via
activation of soluble guanylate cyclase. A, ODQ does not
block cGMP-mediated depression of the GABA response. Cells were
internally dialyzed with control gluconate solution and externally
perfused with ODQ (1 µM). Application of ODQ potentiated
the GABAA currents by 20 ± 2% (n = 6). After a 5 min perfusion with ODQ alone, cells were perfused with
a combination of ODQ and 8Br-cGMP (1 mM) (overhead
bar). 8Br-cGMP depressed the GABA response by 35 ± 4%
from the new, elevated baseline (n = 6).
B, Summary of the dose-dependent depression of the peak GABA current by SNAP, an NO agonist, is shown. At the three
concentrations tested, SNAP (500, 100, and 50 µM)
depressed the GABA-evoked current by 37 ± 2%
(n = 6), 34 ± 4% (n = 8), and 24 ± 4% (n = 5), respectively, when
applied to a naive cell (control, solid bar; mean ± SEM). Immediately after a 10 min preincubation with the sGC
inhibitor ODQ (1 µM; hatched bar), SNAP
depressed the GABA-evoked currents by 16 ± 5%
(n = 3), 17 ± 4% (n = 8), and 11 ± 2% (n = 5), respectively (*p = 0.05, unpaired two-tailed Student's
t test, compared with control in SNAP alone).
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A component of cGMP-induced depression requires PKG
One target of cGMP is PKG. To determine whether PKG activation was
responsible for depression of GABA currents, we blocked the activation
of PKG in individual amacrine cells by including a pseudosubstrate
inhibitory peptide (GKIP) in the intracellular patch pipette solution
(Glass, 1983; Glass and Smith, 1983; McMahon and Ponomareva, 1996). The
average size of GABA currents in cells internally perfused with 50 µM GKIP for 10 min (867 ± 208 pA) was not
significantly different than that in untreated cells (835 ± 136 pA). However, SNAP produced only an 18 ± 2% mean depression in
six cells internally perfused with GKIP, approximately half of the
SNAP-induced depression observed in control cells (Fig. 4).
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Figure 4.
Inhibition of cGMP-dependent protein kinase
partially blocks SNAP-induced depression. A, Inward
currents elicited by 250 msec application of GABA (50 µM;
overhead bars) to either a control cell or one perfused
with a pseudosubstrate peptide inhibitor of PKG (GKIP; 50 µM). The three traces presented for each
cell were recorded before (Pre), during
(SNAP), or after (Recovery) application
of SNAP (500 µM). B, Time course of
inhibition of GABA currents by 500 µM SNAP (timing of
application indicated by overhead bar) in five cells
recorded with gluconate internal solution and in six cells recorded
with solution containing GKIP. GKIP blocked approximately half of the
SNAP-induced depression [GKIP, 18 ± 1% (n = 6); control, 37 ± 2% (n = 5)].
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Similar effects of GKIP were observed when cells were internally
dialyzed with cGMP (Fig. 5). Twenty-five
minutes after achieving whole-cell recording, GABA responses in cells
perfused with 1 mM cGMP were depressed by 58 ± 3%
(n = 8), compared with 10 ± 1% in the control
solution (n = 11). In the presence of GKIP, cGMP
decreased GABA-evoked currents by 35 ± 10% (n = 8) over the same period. Regardless of whether intracellular cGMP was
elevated directly by addition to the pipette or indirectly with an
analog of NO, inhibition of PKG blocked ~50% of the cGMP-induced
depression of GABA currents.
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Figure 5.
GKIP partially blocks cGMP-induced depression.
A, GABA-evoked currents immediately after establishment
of recording and after 20 min are shown in three cells perfused with
either control solution, 1 mM cGMP, or cGMP plus 50 µM GKIP. Overhead bars indicate timing of GABA
application. B, Inclusion of cGMP in the recording
pipette caused a 58 ± 3% (n = 8) decline in
the GABA response compared with a 10 ± 1% (n = 11) decline in control cells over the course of 25 min. When cells
were dialyzed with cGMP plus GKIP, GABA responses declined by 35 ± 10% (n = 8) over the same time period, about
one-half of the reduction observed in the presence of cGMP alone. For
each cell, the amplitude of the peak GABA response was normalized to
the initial amplitude. Normalized amplitudes were then averaged and
binned at 2 min intervals. The points plotted are
mean ± SEM of each bin.
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Depression of GABA currents was also observed when phosphatase activity
was inhibited either with a phosphate-based internal solution (Kennelly
et al., 1993; Thiebart-Fassy and Hervagault, 1993; Weiner et
al., 1993; Caselli et al., 1994; Gao and Fonda, 1994; Bernardi et al.,
1995) or with 1 µM microcystin-LR, an inhibitor of type 1 and 2A phosphatases (Honkanen et al., 1990). Data are summarized in
Figure 6A. Both
phosphate (39 ± 2%; n = 5) and microcystin (43 ± 5%; n = 12) produced a significant
depression of the GABA response compared with that seen in the control
(10 ± 1%; n = 11).
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Figure 6.
Phosphatase inhibition unmasks PKG-dependent
run-down of the GABA response. A, Summary of normalized
GABA currents recorded after 30 min in cells internally perfused with
control (gluconate), phosphate, or gluconate plus 1 µM
microcystin-LR. After 30 min, GABA responses recorded in control cells
were reduced by 10 ± 1% (n = 11) compared
with reductions of 39 ± 2% in high phosphate (n = 5) and 43 ± 5% (n = 12) in microcystin, consistent with the unmasking of a kinase-dependent
mechanism (*p = 0.05, one-way ANOVA, compared with
control). B, Averaged time course of six cells dialyzed
with GKIP (50 µM) plus high phosphate and five cells
dialyzed with high phosphate alone. GKIP prevented run-down of the GABA
response in high phosphate solution. GABA currents were reduced by
12 ± 2% in cells dialyzed with GKIP plus phosphate compared with
10 ± 1% in cells with gluconate.
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Run-down of GABA currents during phosphatase inhibition might be
expected if endogenous levels of cGMP are sufficiently high to activate
PKG or if basal PKG activity is sufficient to phosphorylate targets
when counteracting phosphatases are inhibited. Alternatively, run-down
may be caused by other factors not related to PKG. To distinguish
between these two possibilities, we included the PKG peptide inhibitor
in the phosphate-based internal solution. Run-down of GABA currents
because of high phosphate was almost completely blocked by inhibiting
PKG (Fig. 6B). The observation that inhibition of PKG
was sufficient to completely prevent depression of GABA currents in
response to basal, but not stimulated, levels of cGMP prompted us to
consider the possibility that cGMP could be acting on additional
targets besides PKG.
A second component of cGMP-induced depression of GABA currents
requires PDE activation
cGMP is known to activate a cGMP-stimulated cAMP-phosphodiesterase
(GS-PDE), an effect that is independent of PKG activation (Beavo et
al., 1994). Because cAMP and its analogs have been shown to potentiate
amacrine cell GABA responses (Feigenspan and Bormann, 1994), activation
of GS-PDE by cGMP could indirectly depress GABA currents by hydrolyzing
intracellular cAMP. cIMP is reported to be two orders of magnitude more
potent at activating GS-PDE than at activating PKG (Miller et al.,
1973). We therefore tested the possibility that selective activation of
GS-PDE with cIMP might depress GABA currents.
Application of a cell permeant analog of cIMP, 8Br-cIMP (250 µM), depressed GABA currents by an average of 18 ± 6% (Fig. 7B, hatched
bar; n = 6). The inhibition of GABA currents
by 8Br-cIMP in an individual cell is shown in Figure 7A.
When cells were internally perfused with the phosphate-based internal
solution and 10 µM cAMP to ensure maximal PKA-dependent
phosphorylation (Simmons and Hartzell, 1988), 8Br-cIMP depressed the
peak GABA current by 37 ± 3% (Fig. 7B, left
gray bar; n = 5). The tendency for the response to relax during prolonged 8Br-cIMP exposure has also been
observed for hippocampal calcium currents under similar experimental conditions (Doerner and Alger, 1988). The depression induced by 8Br-cIMP was blocked by internal perfusion of the PDE inhibitor IBMX
(Fig. 7B, right gray bar;
n = 3). In a separate group of cells dialyzed with IBMX
and the phosphate-based internal solution, the run-down of
GABAA currents over 20 min was 27 ± 7%
(n = 5). This difference was not statistically
significant when compared with cells perfused with phosphate-based
solution alone (28 ± 4%; n = 5). Our results
suggest that amacrine cells may contain a PDE that is inhibited by IBMX
and stimulated by cIMP, resulting in a depression of GABA currents.
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Figure 7.
Activation of phosphodiesterase also suppresses
GABA currents. A, Application of 8Br-cIMP
(overhead bar) to a cell perfused with phosphate plus 10 µM cAMP produced a rapid decline in the GABA-evoked
current. All currents were normalized to the amplitude of the current
just before the application of 8Br-cIMP. The insets are
currents evoked by 250 msec applications of 50 µM GABA to a representative cell before, during, and after the application of
8Br-cIMP, as indicated by the numbers. B,
Application of the GS-PDE activator 8Br-cIMP (500 µM)
depressed GABA-evoked currents by 18 ± 6% in cells perfused with
gluconate (n = 6; hatched bar) and
by 37 ± 3% in cells perfused with phosphate plus cAMP
(n = 5; left gray bar). In contrast,
8Br-cIMP did not diminish the GABA current in cells perfused with IBMX,
consistent with the idea that 8Br-cIMP activated PDE
(n = 3; right gray bar;
p = 0.05, unpaired two-tailed Student's
t test).
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Inhibition of both PKG and PDE blocked most of the SNAP-induced
depression that was not blocked by PKG inhibition alone. We compared
the depressant effects of 100 µM SNAP when GKIP alone was
added to the internal solution with that observed when 1 mM IBMX was added together with GKIP. We used twice the concentration of
GKIP (100 µM) used in previous experiments to help insure
that PKG was maximally blocked. The averaged time course and peak
inhibition of SNAP under each condition are shown in Figure
8. SNAP depressed the GABA current to a
lesser degree (21 ± 3%; n = 15) in cells perfused with GKIP than in control cells (34 ± 4%;
n = 10). Moreover, internal perfusion with a
combination of GKIP and IBMX in 14 cells further limited the
SNAP-induced depression of the GABA response to 11 ± 2%.
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Figure 8.
Inhibition of phosphodiesterase attenuates
PKG-independent depression. A, Averaged time course of
SNAP-induced depression of GABA currents in cells dialyzed with control
(gluconate) internal solution, GKIP (100 µM), or GKIP
plus 1 mM IBMX. Application of 100 µM SNAP is
indicated by the overhead bar.
B, Summary of results from A. SNAP
depressed GABA currents by 34 ± 4% in control cells (n = 10), by 21 ± 1% in cells dialyzed with
GKIP (n = 15), and by 11 ± 1% in cells
dialyzed with GKIP plus IBMX (n = 14). The single asterisk indicates p = 0.05 compared with control. The double asterisks indicate
p = 0.05 compared with both control plus GKIP;
p = 0.05; one-way ANOVA.
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DISCUSSION |
Our findings support the following NO-mediated cascade regulating
GABAA receptors on retinal amacrine cells. NO stimulates soluble guanylate cyclase, which raises the intracellular concentration of cGMP. cGMP increases PKG-mediated phosphorylation and concomitantly decreases PKA phosphorylation, via stimulation of GS-PDE. These events
act synergistically to depress GABAA receptor-gated
currents. With continuous application of NO agonists, recovery of the
GABA response was often observed.
Of the seven known forms of PDE, two are regulated by cGMP. Although
both catalyze the hydrolysis of cAMP, the type III form is inhibited by
cGMP, whereas the type II form (GS-PDE) is stimulated by cGMP (Beavo et
al., 1971a,b). There is precedence for a GS-PDE indirectly modulating
ion channel function. Calcium currents
(ICa) in cardiac myocytes (Hartzell and
Fischmeister, 1986) and hippocampal CA1 pyramidal cells (Doerner and
Alger, 1988) are potentiated by PKA phosphorylation, yet they are
depressed by cGMP via PKG-independent activation of a GS-PDE. In these
cells, cGMP stimulation of GS-PDE lowers intracellular cAMP,
subsequently reducing PKA activity.
Two lines of evidence support our contention that regulation of
amacrine cell GABA-gated currents by GS-PDE may be analogous to
regulation of calcium currents in hippocampal pyramidal cells and
cardiac myocytes. First, the application of 8Br-cIMP, a selective activator of GS-PDE, produced a marked depression of GABA currents. This effect was blocked by the broad-spectrum PDE inhibitor IBMX, consistent with an action on GS-PDE (Fig. 7). Second, intracellular dialysis with IBMX also blocked a substantial fraction of the SNAP-induced depression that was not blocked by PKG inhibition alone
(Fig. 8). However, because neither inhibition of sGC nor the combined
inhibition of PKG and PDE completely blocked SNAP-induced depression of
GABA currents, it is possible that SNAP might also inhibit GABA
currents via a third mechanism that is completely independent of cGMP,
perhaps by acting directly on the channel, as has been demonstrated for
olfactory cAMP-gated channels (Broillet and Firestein, 1996).
When the phosphorylation-dependent component of the depression was
pharmacologically isolated by inhibiting phosphatases, GABA currents
ran down with time. Because there was no source of exogenous cGMP in
this experiment, there must have been sufficient active PKG to produce
a net increase in phosphorylation while phosphatases were inhibited. In
phosphate, run-down of IGABA was slow, ~1.5
pA/min, and this may reflect a very low level of basal PKG activity.
Our data also suggest that, in the absence of cGMP stimulation, GS-PDE
is inactive, because inhibition of PKG by GKIP was sufficient to block
nearly all of the run-down. One possible explanation for this finding
is that higher concentrations of cGMP are needed to activate GS-PDE
than PKG. Biochemical studies have suggested that PKG can be activated
by concentrations of cGMP that are between 10 and 100 times lower than
those that are required to activate GS-PDE (Martins et al., 1982; for
PDE review, see Butt et al., 1993). Addition of extracellular cGMP
(Fig. 4) or SNAP (Fig. 4) produced a depression of GABA currents that
was only partially blocked by inhibition of PKG, presumably because the
intracellular levels of cGMP were increased sufficiently to activate
both GS-PDE and PKG. It seems unlikely that the partial block of
SNAP-induced IGABA is attributable to incomplete
PKG inhibition by GKIP, because doubling the concentration of the peptide inhibitor from 50 to 100 µM did not decrease the
suppression of GABA currents by SNAP. Second, GKIP is a pseudosubstrate
inhibitor, acting at the PKG catalytic site rather than at the
regulatory site. Thus, it is not competitively antagonized by
increasing intracellular cGMP.
Several reports have defined a mechanism by which activators of
adenylate cyclase, such as dopamine, enhance GABAA currents in mammalian retina (Veruki and Yeh, 1992, 1994; Feigenspan and Bormann, 1994). The present study suggests that activators of guanylate
cyclase, such as NO, do just the opposite. The enzymes that synthesize
dopamine and NO are both located in subpopulations of GABAergic
amacrine cells (Wassle and Chun, 1988; Vaney and Young, 1988; Darius et
al., 1995). Thus, GABAergic transmission in the inner retina may be
modulated in a push-pull manner by activators of adenylate cyclase
such as dopamine or vasoactive intestinal peptide on the one hand and
by NO on the other.
| |
FOOTNOTES |
Received Aug. 26, 1997; revised Jan. 12, 1998; accepted Jan. 12, 1998.
This work was supported by National Institutes of Health Grant EY10254
(S.N.), by a Medical Scientist Training Program grant (E.M.W.), by
Alcon Laboratories, and by an unrestricted grant from Research to
Prevent Blindness, Inc. (S.N.). We thank Regeneron Pharmaceuticals for
the gift of BDNF and Alex Peinado for helpful comments on this
manuscript.
Correspondence should be addressed to Dr. Eric M. Wexler, Kennedy
Center Room 525, 1410 Pelham Parkway South, Bronx, NY 10461.
| |
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Abstract
Introduction
