From the * Howe Laboratory of Ophthalmology, Harvard Medical School and the Massachusetts Eye and Ear Infirmary, Boston, Massachusetts 02114; and Laboratory of Molecular Biology, University of Wisconsin, Madison, Wisconsin 53706
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ABSTRACT |
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Light adaptation in vertebrate photoreceptors is thought to be mediated through a number of biochemical feedback reactions that reduce the sensitivity of the photoreceptor and accelerate the kinetics of the photoresponse. Ca2+ plays a major role in this process by regulating several components of the phototransduction cascade. Guanylate cyclase and rhodopsin kinase are suggested to be the major sites regulated by Ca2+. Recently, it was proposed that cGMP may be another messenger of light adaptation since it is able to regulate the rate of transducin GTPase and thus the lifetime of activated cGMP phosphodiesterase. Here we report measurements of the rates at which the changes in Ca2+ and cGMP are followed by the changes in the rates of corresponding enzymatic reactions in frog rod outer segments. Our data indicate that there is a temporal hierarchy among reactions that underlie light adaptation. Guanylate cyclase activity and rhodopsin phosphorylation respond to changes in Ca2+ very rapidly, on a subsecond time scale. This enables them to accelerate the falling phase of the flash response and to modulate flash sensitivity during continuous illumination. To the contrary, the acceleration of transducin GTPase, even after significant reduction in cGMP, occurs over several tens of seconds. It is substantially delayed by the slow dissociation of cGMP from the noncatalytic sites for cGMP binding located on cGMP phosphodiesterase. Therefore, cGMP-dependent regulation of transducin GTPase is likely to occur only during prolonged bright illumination.
Key words: light adaptation; guanylate cyclase; phosphodiesterase; rhodopsin kinase; Ca2+ ![]() |
INTRODUCTION |
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Photoresponses in vertebrate photoreceptors begin when
light activates an enzymatic cascade including rhodopsin, transducin, and phosphodiesterase (PDE).1 The resulting decrease in cGMP causes closure of cationic
channels in the plasma membrane of the photoreceptor outer segment (reviewed in Chabre and Deterre,
1989; Pugh and Lamb, 1990
). Photoreceptors adapt to
ambient light through feedback reactions that regulate either the catalytic activity or the catalytic lifetime of
individual components of the phototransduction cascade. It is well established that Ca2+ plays a significant
role in this process (reviewed by Lagnado and Baylor,
1992
; Koutalos and Yau, 1993
; Bownds and Arshavsky, 1995
). Ca2+ declines in response to light because it can
no longer enter through the cationic channels while it
continues to be extruded through the Na/Ca,K exchanger. There are at least three sites of Ca2+ regulation in the cascade. The first is the regulation of the lifetime of light-activated rhodopsin. Rhodopsin is turned
off when it is phosphorylated by rhodopsin kinase followed by the binding of arrestin. Rhodopsin phosphorylation is inhibited when rhodopsin kinase forms a
complex with the Ca2+-binding protein, recoverin. The
light-dependent decrease in cytoplasmic Ca2+ is thought
to cause the dissociation of this complex, increasing the
rate of rhodopsin phosphorylation. Another site of Ca2+
regulation is guanylate cyclase, the enzyme responsible
for cGMP synthesis. This regulation is conferred through
the Ca2+ binding proteins known as guanylate cyclase
activating proteins (GCAPs) (Palczewski et al., 1994
;
Dizhoon et al., 1995). Cyclase activity is low in darkness
and increases when Ca2+ levels drop in response to
light. The third site is the regulation of the sensitivity of
the cationic channel to cGMP. Lowering Ca2+ causes
the channel to become more sensitive to cGMP due to
the dissociation of calmodulin (CaM) or a closely related Ca2+ binding protein. This might result in the accelerated recovery of the photoresponse by facilitating
the reopening of channels. In addition, the gain of the
cascade may also be regulated by Ca2+ independently
of changes in inactivation and recovery (Lagnado and
Baylor, 1994
). This hypothesis is based on the observation that the rate of the photoresponse rising phase in
truncated rods is reduced when Ca2+ concentration is
lowered.
cGMP might also serve as a messenger of adaptation
by regulating the duration of PDE activation by transducin (Arshavsky et al., 1991, 1992
; Arshavsky and Bownds,
1992
; Cote et al., 1994
). cGMP binding to noncatalytic
sites on the PDE molecule modulates the rate of transducin GTPase, the reaction responsible for PDE shut
off. When these sites are occupied by cGMP, the rate of
GTP hydrolysis is relatively slow and thus PDE stays active for a relatively long time. When light causes the decline of free cGMP, followed by cGMP dissociation
from the noncatalytic sites, the rate of GTP hydrolysis is
accelerated by severalfold and the duration of PDE activation is reduced. This mechanism could accelerate the
recovery of the photoresponse.
To understand the time course of the onset of the
feedback controls discussed above, three issues should
be addressed: (a) the kinetic parameters for the regulation of the enzymes targeted by feedback messengers,
(b) the rate of reduction of free feedback messengers
in the photoreceptor cytoplasm in response to light, and
(c) the delay between feedback messenger decline and
the corresponding change in enzymatic activity (including the dissociation of bound messengers from regulatory proteins and the subsequent change in protein-
protein interactions). The rates of cGMP and Ca2+ reductions in rod outer segments (ROS) have been well
characterized. Changes in free cGMP are simply reflected
in the changes in the photocurrent that senses cGMP
concentration with only a millisecond delay (reviewed
by Yau and Baylor, 1989). Changes in free Ca2+ have
been studied by a number of laboratories (McCarthy et
al., 1994
, 1996
; Gray-Keller and Detwiler, 1994
; Younger
et al., 1996
; Sampath et al., 1997
). Ca2+ decline with
light in frog, gecko, and salamander photoreceptors is
described as the sum of at least two exponential processes, one with a subsecond time constant and another
with a time constant of several seconds. Practically nothing is known about the delay between Ca2+ and cGMP
decline and the changes in corresponding enzymatic
activities. Physiological experiments have indirectly addressed some of these questions, but few have been approached in direct biochemical experiments.
Our goal was to study how fast the reduction in free Ca2+ results in the stimulation of guanylate cyclase activity and the disinhibition of rhodopsin kinase and how fast the reduction in free cGMP results in the acceleration of transducin GTPase rate. Bullfrogs were selected as an experimental animal because all three of these feedback reactions exist in their rods, frog ROS can be obtained in a quantity sufficient for biochemical experiments and it is one of only three species where the changes in intracellular free Ca2+ are described in detail. We have found that the Ca2+-dependent increase in the activities of rhodopsin kinase and guanylate cyclase occur very quickly, on a time scale of a few hundred milliseconds. This indicates that the regulation of these enzymes takes place on the time frame of the Ca2+ change in photoreceptor cytoplasm. In contrast, changes in transducin GTPase activity do not directly follow the decline of free cGMP, but are delayed for several tens of seconds due to the slow dissociation of cGMP from PDE noncatalytic cGMP binding sites.
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METHODS |
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Materials
[8-3H]cGMP, [-32P]GTP, [
-33P]GTP, and [
-32P]ATP were purchased from Du Pont-NEN (Boston, MA), Percoll from Pharmacia LKB Biotechnology Inc. (Piscataway, NJ), potassium isethionate from Eastman Kodak Co. (Rochester, NY), nitrocellulose filters from Whatmann Inc. (Clifton, NJ), and BAPTA and Fluo-3
from Molecular Probes, Inc. (Eugene, OR). All other chemicals
were obtained from Sigma Chemical Co. (St. Louis, MO) (CaM-PDE and CaM from bovine brain were product No. P9529 and
P2277, respectively).
Solutions
The Ringer's solution used to isolate ROS contained (mM): 105 NaCl, 2 KCl, 2 MgCl2, 1 CaCl2, and 10 HEPES, pH 7.5. The
pseudointracellular medium used in all experiments contained
95 mM potassium isethionate, 15 mM sodium isethionate, 5 mM
MgCl2, 2 mM dithiothreitol, 10 µM leupeptin, 100 kallikrein U/ml
aprotinin, and 10 mM HEPES, pH 7.8. The pseudointracellular
medium was passed over a Chelex 100 column before MgCl2 addition in order to remove contaminating Ca2+. All the solutions
had a final osmolarity of 232-238 mosM. Where required, Ca2+ in
the pseudointracellular medium was buffered by BAPTA as described in Klenchin et al. (1995). Since commercially available
CaCl2 and BAPTA salts contain unpredictable levels of H2O,
which could significantly affect their concentrations on making
stock solutions, the H2O content in these salts was determined
gravimetrically and their formula weights were adjusted accordingly. 4× stock solutions with varying concentrations of CaCl2
and 20 mM BAPTA concentration were prepared in the pseudointracellular medium. Free Ca2+ concentrations of these stocks
were calculated using the program BAD (Brooks and Story,
1992). The validity of this method was checked using both a calcium selective electrode (Microelectrodes, Inc., Londonderry,
NH) and Fluo-3 indicator dye.
Preparation of Rod Outer Segments
Live bullfrogs (Rana catesbeiana or Rana grylio) were purchased
from commercial sources and maintained with feeding on a 12-h light-dark cycle for at least 2 wk before use (Woodruff and
Bownds, 1979). All manipulations were performed under infrared illumination. Animals were killed by decapitation, retinas
were removed, placed into Ringer solution containing 5% Percoll, and ROS were purified on a Percoll gradient as described in
Biernbaum and Bownds (1985a)
. Intact ROS were washed free of
Percoll in pseudointracellular medium and kept on ice. Before
each experiment, ROS were disrupted in a Potter-Elvehjem homogenizer, creating a membrane suspension with no structure
detectable under light microscopy (Dumke et al., 1994
). Rhodopsin concentration was determined spectrophotometrically according to Bownds et al. (1971)
. All experiments were carried out at 22°C.
Guanylate Cyclase Assay
Guanylate cyclase activity was determined in ROS homogenates
using thin layer chromatography as described by Dizhoor et al. (1994). The reaction was performed in 500 µl Eppendorf tubes
upon vigorous vortexing. The reaction was started by the addition of 10 µl of pseudointracellular medium supplemented with
[
-33P]GTP (either 1 mM or 200 µM), 10 µM ATP, and 10 mM
[3H]cGMP to 10 µl of the ROS suspension supplemented with
100 µM zaprinast and Ca2+-BAPTA buffer. 33P-labeled GTP, ATP,
and zaprinast were used to reduce PDE activity that would otherwise hydrolyze the cGMP formed by guanylate cyclase, confounding the measurement of its activity. 33P-labeled GTP was used
rather than 32P-GTP to avoid bleaching of rhodopsin by Cerenkov radiation, caused by the decay of 32Pi, and the subsequent activation of PDE (Biernbaum et al., 1991
). ATP was provided to
quench contaminating bleached rhodopsin in the preparation
through its phosphorylation by endogenous kinase. Zaprinast is a
potent PDE inhibitor (Gillespie and Beavo, 1989
). This strategy
reduced the hydrolysis of cGMP, as measured by the reduction in
added [3H]cGMP, to <5% over the time course of our experiments. The reaction was quenched with 100 µl of 50 mM EDTA,
pH 7.0, followed by 1 min boiling to precipitate proteins. Samples were then centrifuged on a Beckman Microfuge E for 10 min and 10-µl aliquots were loaded on PEI-cellulose thin layer
chromatography plates (EM Sciences, Gibbstown, NJ). The nucleotides were separated using 0.2 M LiCl. Spots containing
cGMP were visualized under ultraviolet illumination, cut from the
plate, eluted with 2 M LiCl, mixed with 10 ml ScintiSafe cocktail
(Fisher Scientific Co., Santa Clara, CA) and counted in a scintillation counter.
Rhodopsin Phosphorylation Assay
Rhodopsin phosphorylation was measured essentially as described by Klenchin et al. (1995). The reaction was started with
the addition 10 µl of [
-32P]ATP to 10 µl ROS containing Ca2+-BAPTA buffer and 30 µM myristoylated recombinant bovine recoverin. The reaction was quenched with 80 µl of 50 mM EDTA,
100 mM KF, and 100 mM Na-phosphate buffer, pH 7.5. This solution terminated both rhodopsin phosphorylation and dephosphorylation. 50 µl of quenched sample was applied to nitrocellulose filter and washed six times with 1 ml of 100-mM Na-phosphate buffer, pH 7.5. Filters were placed in scintillation vials,
dissolved in 2 ml glacial acetic acid, mixed with 10 ml ScintiSafe
(Fisher Scientific Co.) and counted. Recombinant recoverin was
produced using a bacterial expression system (Dizhoor et al.,
1993
). The expression system was a kind gift from Dr. J.B. Hurley
(University of Washington, Seattle, WA).
cGMP Binding Assay
cGMP binding to the PDE noncatalytic sites was determined by
the nitrocellulose filter binding technique described in detail by
Cote and Brunnock (1993). ROS were incubated for 30 min at room temperature to completely dissociate the endogenous
cGMP from the PDE noncatalytic sites. Nucleotide-depleted ROS
were incubated for 1 min with 3 µM [3H]cGMP, a time sufficient
to reach binding equilibrium (Cote and Brunnock, 1993
). Dissociation of the labeled cGMP was then initiated by a chase with either an excess of the unlabeled cGMP or a mixture of CaM-PDE
with CaM (80 U CaM per unit CaM-PDE). At different times after
the chase, 15-µl portions were added to nitrocellulose filters. The
filters were rinsed with three 1-ml portions of ice-cold pseudointracellular medium, dissolved in glacial acetic acid and counted
in ScintiSafe cocktail.
GTPase Assay
Transducin GTPase activity was determined by a modified method
of Godchaux and Zimmerman (1979) described by Arshavsky et al. (1991)
. 20 µl of bleached ROS (20 µM rhodopsin final concentration) were mixed with 10 µl of [
-32P]GTP (4 µM final
concentration) in 1.5 ml Eppendorf tubes. The reaction was
stopped in 3 s with 100 µl of 6% perchloric acid. The samples
were incubated for 10 min with 0.7 ml activated charcoal (100 mg/ml in 50 mM Na-phosphate buffer, pH 7.5), sedimented, and 32Pi in the supernatant was measured with ScintiSafe cocktail.
Control experiments carried out in darkness indicate that >90%
of GTP hydrolysis was light-dependent. Therefore, we conclude
that most of the GTPase activity in these experiments is attributable to transducin.
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RESULTS |
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Ca2+ Feedback
Guanylate cyclase activity changes quickly after a rapid change in
free Ca2+.
The regulation of guanylate cyclase activity by
Ca2+ in rod photoreceptor outer segments has been
well characterized in bovine rod photoreceptors (Koch
and Stryer, 1988; Gorczyca et al., 1994b
; Dizhoor et al.,
1994
). Ca2+ regulation of the guanylate cyclase in frog
ROS has also been established (Coccia and Cote, 1994
);
however, a detailed analysis of the Ca2+ dependence
was not performed. Such an analysis in a suspension of
frog ROS is shown in Fig. 1. The cyclase activity was expressed as the cGMP concentration produced in the
ROS cytoplasm per second. The value of 6 mM rhodopsin with respect to the ROS cytoplasm was used (see
Bownds and Arshavsky, 1995
). The Ca2+ dependence
was approximated by the Hill equation:
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(1) |
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(2) |
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Rhodopsin Kinase Activity Increases Quickly after a Rapid
Reduction in Free Ca2+
We measured how fast the rhodopsin kinase activity increased when Ca2+ was abruptly
changed from high to low concentration in experiments similar to those described for the guanylate cyclase.
Previous studies have shown that the affinity of recoverin for rhodopsin kinase is low and that significant effects of recoverin on rhodopsin phosphorylation in ROS
suspensions may be observed only with the addition of
exogenous recoverin (Kawamura, 1993; Klenchin et al., 1995
). Therefore, we supplemented ROS suspensions
with 30 µM exogenous recoverin, which approximates
the endogenous concentration in the cytoplasm of intact frog ROS (Klenchin et al., 1995
). We used myristoylated recombinant bovine recoverin, which regulates rhodopsin phosphorylation in frog ROS in the
same way as the endogenous frog recoverin (Kawamura
et al., 1993
; Klenchin et al., 1995
). Recoverin-supplemented ROS suspensions were preincubated either at
10 µM or 10 nM Ca2+ for at least 2 min before the initiation of the rhodopsin phosphorylation reaction.
Rhodopsin was fully bleached with white light immediately before initiation of the reaction to ensure that it is
not limiting in the reaction. The reaction was started by
the addition of 25 µM [
-32P]ATP according to one of
three protocols (analogous to the cyclase conditions
described above): (a) ROS in 10 µM Ca2+ were mixed
with [
-32P]ATP in 10 µM Ca2+; (b) ROS in 10 nM Ca2+
were mixed with [
-32P]ATP in 10 nM Ca2+; (c) ROS in
10 µM Ca2+ were mixed with [
-32P]ATP in the pseudointracellular medium containing 1 mM EGTA, an
amount sufficient to reduce the Ca2+ concentration to
10 nM upon the start of the reaction.
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cGMP Feedback
cGMP dissociation from the PDE noncatalytic sites after a
rapid reduction in free cGMP.
One goal of this study was
to correlate the rate of cGMP dissociation from PDE
noncatalytic cGMP binding sites with the dynamics of
transition between "slow" and "fast" transducin GTPase
measured in the same ROS suspension. To carry out
these experiments, we needed to modify the pulse-chase technique previously used for monitoring cGMP
dissociation from the noncatalytic sites (Cote and Brunnock, 1993; Cote et al., 1994
). In those studies, noncatalytic sites were first loaded with radio-labeled cGMP,
and then its dissociation from the sites was monitored
after a chase with a large excess of nonlabeled cGMP.
This approach could not be used in this study since the
nonlabeled cGMP quickly substitutes labeled cGMP in
the noncatalytic sites and, therefore, they remain occupied during the entire course of the experiment. Instead, we needed to rapidly remove free cGMP as well
as cGMP dissociating from the noncatalytic sites from
the reaction mixture. To accomplish this, we substituted the cGMP chase with the addition of an excess of
CaM-PDE from bovine brain, which hydrolyzes free but
not bound cGMP. We decided to use CaM-PDE rather
than any other phosphodiesterase type because it has a
high level of cGMP hydrolytic activity, lacks noncatalytic cGMP binding sites, and is commercially available. CaM-PDE in the chase was fully activated by CaM and
CaCl2. Control experiments have shown that Ca2+,
CaM, and CaM-PDE do not interfere with GTP hydrolysis by transducin and that addition of Ca2+ and CaM
does not alter the kinetics of cGMP dissociation from PDE noncatalytic sites as measured by a chase with excess cGMP (data not shown).
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Transition between slow and fast transducin GTPase directly follows cGMP dissociation from PDE noncatalytic binding sites.
The correlation of cGMP dissociation from
the PDE noncatalytic sites with the transition between
slow and fast transducin GTPase is shown in Fig. 6.
Bleached ROS were preincubated with [3H]cGMP to
fully occupy the PDE noncatalytic sites, and then the dissociation of bound cGMP was initiated at time zero
by a chase with CaM/CaM-PDE. Aliquots were taken
from this mixture to determine either the amount of
bound cGMP (Fig. 6 A) or the rate of transducin GTPase (Fig. 6 B). The GTPase determinations were initiated with the addition of either [-32P]GTP or a mixture of [
-32P]GTP and 1 mM cGMP. GTPase measurements were conducted for only 3 s to minimize further
cGMP dissociation during the measurement. The measurements with GTP/cGMP mixture provide a control
that shows that the rate of slow GTPase remains constant during the course of this experiment. The major
observation of this experiment is that the extent of the
noncatalytic site occupancy by cGMP precisely corresponds to the extent of the transition of transducin GTPase between slow and fast rates. Both processes were described by single exponents: the rate constant for
cGMP dissociation was 0.0030 ± 0.0002 s
1 (n = 4),
while the rate constant for the transition between slow and fast GTPase was 0.0031 ± 0.0008 s
1 (n = 4). This
coincidence indicates that there is no substantial delay
between dissociation of cGMP from the noncatalytic
binding sites and the onset of fast transducin GTPase.
Further, the correspondence of the extent of cGMP
dissociation with the extent of GTPase activation indicates that each PDE noncatalytic site plays an equal role in regulating transducin GTPase.
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cGMP dissociation from the noncatalytic sites of activated
PDE after a rapid reduction of free cGMP.
Having established
that the increase in transducin GTPase directly follows
the dissociation of cGMP from the PDE noncatalytic sites, we wanted to know how fast cGMP might dissociate from these sites during the photoreceptor light response. The dissociation rate derived from the data
presented in Fig. 5 gives only a slower limit for this process because PDE activation by transducin results in an
accelerated rate of cGMP dissociation from the noncatalytic sites (Yamazaki et al., 1982, 1996
; Cote et al., 1994
).
In Fig. 7, we compare the rates of cGMP dissociation
from the noncatalytic sites of activated and nonactivated PDE after a chase with CaM/CaM-PDE. As reported earlier (Cote and Brunnock, 1993
; Cote et al.,
1994
; Yamazaki et al., 1996
) and as shown above in Fig.
5, cGMP dissociation from nonactivated PDE is a single
exponential process, while cGMP dissociation from
transducin-activated PDE is biphasic. cGMP dissociates from 32 ± 7% of the sites with a rate constant of 0.11 ± 0.04 s
1 and from 68 ± 7% with a rate constant of 0.006 ± 0.001 s
1 (n = 5). These rates are at least threefold
faster than those obtained by Cote et al. (1994)
after a
chase with an excess of unlabeled cGMP. A reasonable
explanation for this difference is provided by Yamazaki
et al. (1996)
, who suggested that cGMP release from
the noncatalytic sites of activated PDE may be inhibited by high concentrations of cGMP. Cote et al. (1994)
and
Yamazaki et al. (1996)
proposed that the biphasic
cGMP dissociation is due to a heterogeneity in the noncatalytic cGMP binding sites. However, the mechanism
may be different considering that the maximal high affinity cGMP binding observed in our experiments corresponds to two cGMP molecules per PDE holoenzyme,
while the amplitudes of the two phases of cGMP dissociation are twofold different rather than being equal as
expected.
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DISCUSSION |
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Ca2+ Feedback on Guanylate Cyclase
In the experiments reported in this study, guanylate cyclase activity quickly increases to its maximum after a
reduction of the Ca2+ concentration with a time constant of ~200 ms (Fig. 3 A). With this parameter, and
knowing the K1/2 and the Hill coefficient for the Ca2+-dependent regulation of cyclase from Fig. 1, we can
predict the time course of the increase in cyclase activity as a result of the light-dependent Ca2+ reduction in
bullfrog ROS. For this analysis, we chose to use a 10-fold regulation of the cyclase by Ca2+, the largest extent
of regulation directly observed in our experiments and
the lower limit of this parameter in intact cells (see Fig. 2). The analysis is based on the time course of the light-dependent Ca2+ reduction in bullfrog ROS cytoplasm
as measured by McCarthy et al. (1996). Ca2+ declines as
the sum of three exponential processes as depicted in
Fig. 8 A, where the Ca2+ concentration is expressed as a
percentage of dark Ca2+ concentration. Some ambiguity remains as to the actual value of Ca2+ concentration
in the dark. McCarthy et al. (1996)
suggest that the
dark Ca2+ concentration is within the range of 200-400
nM, so we performed separate analyses using each of
these values. The predicted time courses for the onset
of the cyclase are shown in Fig. 8 B, where a 200-ms delay between the change in Ca2+ and the change in the
cyclase rate has been imposed. It is seen that the range
of the cyclase activity change is highly dependent on
the value for the dark Ca2+ concentration. A 2.6-fold increase in cyclase activity is observed when the dark Ca2+
is assumed to be 400 nM, whereas only a 1.5-fold increase is observed with 200 nM dark Ca2+. We favor the
higher value of free dark Ca2+ concentration for two
reasons. First, it uses more of the dynamic range of cyclase regulation. A value for K1/2 (255 nM, Fig. 1) that is
higher than the dark Ca2+ level will preclude a regulation of the cyclase from being more than twofold, while
physiological data (see below) argue for a larger extent
of regulation. Second, it is consistent with the estimates
of the dark ROS Ca2+ concentration in other lower vertebrates: 410 nM from Lagnado et al. (1992)
and 534 nM from Sampath et al. (1997)
for salamander, and
550 nM from Gray-Keller and Detwiler (1994)
for
gecko. Importantly, our analysis reveals that the changes
in the cyclase activity after a saturating light occur slightly
faster than the changes in free Ca2+. The cyclase rate
increases 90% within ~5 s, whereas a 90% decrease in
Ca2+ occurs in ~10 s. This is because the Ca2+ dependence of the cyclase is cooperative and the half-saturating Ca2+ concentration is close to the dark Ca2+ level. It
should be noted that our analysis is based on the spatially averaged Ca2+ concentration changes in ROS
(McCarthy et al., 1996
) and, therefore, should be considered as a spatially averaged change in the cyclase rate.
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It is interesting to compare our biochemical analysis
of guanylate cyclase regulation with indirect estimates
obtained using physiological techniques (a more detailed discussion on this topic may be found in a recent
review by Pugh et al., 1997). The first physiological
measurements of the light-dependent increase in cyclase activity were reported by Hodgkin and Nunn
(1988)
in salamander rods. By monitoring the rates of
the photo-sensitive current rise after suddenly inhibiting PDE by IBMX (3-isobutyl-1-methylxanthine), they estimated the maximum increase to be ~10-fold (Fig. 15 from Hodgkin and Nunn, 1988
). A smaller value of sixfold was later reported by Cornwall and Fain (1994)
.
To the contrary, Koutalos et al. (1995a)
made indirect
measurements of the Ca2+ dependence of cyclase activity in truncated salamander ROS and concluded that
the maximal extent of the light-dependent cyclase activation is ~30-fold (assuming 400 nM as the dark Ca2+
concentration). This difference may be explained by
the fact that Koutalos et al. (1995a)
measured the cyclase activity at high Ca2+ to be zero. In fact, their cyclase
measurements had a resolution of 1-2 µM/s (Y. Koutalos, personal communication). Thus, in their experiments, enzymatic rates below ~2 µM/s may have been
indistinguishable from zero and the actual high Ca2+
rate may have been anywhere from zero to 2 µM/s. If
we assume that the cyclase rate at high Ca2+ was the
same as the rate we measured in frogs at the highest ROS concentration, ~1 µM/s, then the dynamic range
of cyclase activation measured by Koutalos et al. (1995a)
would be approximately ninefold, in agreement with
others. This activation level is also consistent with the
estimates by Dawis et al. (1988)
who have found that
bright light causes an 8- to 10-fold increase in the cGMP
metabolic flux rate in toad retinas.
The at least 10-fold range of guanylate cyclase regulation by Ca2+ reported in this study (Fig. 2) is well consistent with the 6-10-fold dark-light difference predicted from the physiological studies. However, the actual range of cyclase stimulation by light calculated in
Fig. 8 B is only 2.6-fold. We propose three simple explanations for this discrepancy. First, the high/low Ca2+
ratio of the cyclase activity in intact photoreceptors
should be greater than the 10-fold that we were able to
measure in Fig. 2 B. Second, it is possible that the actual Ca2+ concentration in a dark adapted frog ROS is
higher than the 400 nM value used in the calculations.
This would allow cyclase to operate over a larger portion of its dynamic range. Third, it is possible that the
K1/2 value for the Ca2+ regulation of cyclase in frog is
higher than in salamander. Indeed, while we observe a
K1/2 of 255 nM in frog, Koutalos et al. (1995a) report a
K1/2 value of 87 nM in salamander. Assuming that the dark Ca2+ levels in frog and salamander are similar and
equal to ~400 nM, then the light-dependent range of
the cyclase activation covers practically the entire Ca2+-dependent range in salamander, whereas only a portion of the Ca2+-dependent range is used by frogs. Another important consequence of the position of the K1/2
for Ca2+-dependent cyclase regulation relative to the
Ca2+ concentration in the dark is that it determines the
rate of increase in the cyclase activity upon illumination. The estimates by Hodgkin and Nunn (1988; see
Fig. 12) indicate that the halftime of cyclase activation
in saturating light is at least 5 s, slower than the 2 s estimated in Fig. 8 B. This is consistent with the possibility that the K1/2 value for the Ca2+ regulation of cyclase in
salamander is lower than in frog, while values for dark
Ca2+ concentrations are similar. However, it could also
reflect an artifact of cyclase estimates by their method
at high light levels.
Ca2+ Feedback on Rhodopsin Kinase
Our study shows that rhodopsin kinase activity also increases practically without delay after a sudden drop in
Ca2+. The predicted time course of kinase activation after the onset of saturating light is less straightforward
than for guanylate cyclase due to an uncertainty in the
Ca2+ range for kinase regulation by Ca2+ recoverin in
vivo. All recent measurements performed in suspensions of disrupted ROS or in reconstituted systems indicate that the half-maximal Ca2+ concentration for the
rhodopsin kinase regulation by recoverin is above the
Ca2+ range in intact rods (Klenchin et al., 1995; Chen
et al., 1995
; Ames et al., 1995
). However, physiological
experiments performed with recoverin knock-out mice
(Dodd et al., 1995
) strongly support the idea that the
Ca2+-dependent regulation of rhodopsin kinase occurs
over the physiological range of Ca2+ concentration
changes: rods lacking recoverin have accelerated photoresponse recovery and are less able to adapt to light.
Two complementary arguments provide a possible resolution of this paradox. First, Zozulya and Stryer
(1992)
argued that Ca2+-recoverin binding to ROS
membranes increases the Ca2+ binding affinity to recoverin. Second, Klenchin et al. (1995)
have pointed
out that because the amount of recoverin in ROS is larger than the amount of rhodopsin kinase, Ca2+ binding to a relatively small fraction of recoverin might be sufficient for inhibition of a relatively large fraction of
kinase. This results in a lower K1/2 for the Ca2+-dependent regulation of kinase than the K1/2 for Ca2+ binding to recoverin. Klenchin et al. (1995)
used both of
these arguments to calculate a K1/2 of ~270 nM for the
Ca2+-dependent kinase regulation in vivo and a maximum extent of the regulation between high and low
Ca2+ of ~10-fold. Based on these numbers, the value of
400 nM for Ca2+ concentration in the dark and our observation that the kinase activity senses the decrease in
Ca2+ with a delay of ~100 ms, we predicted the time
course for the increase in the kinase activity after the
onset of saturating illumination (Fig. 8 C, solid curve).
The onset of rhodopsin kinase activation in this case is
very similar to the onset of the guanylate cyclase.
An important question raised in our study is why the
photoreceptor has two Ca2+-dependent feedback mechanisms, both designed to sense changes in Ca2+ quickly.
If the Ca2+ K1/2 values for these mechanisms are indeed
close as we discussed above, then their coincidence simply allows the photoreceptor to enhance the amplitude
of the Ca2+ feedback regulation. If, however, the actual
Ca2+ K1/2 value for rhodopsin kinase is different from
that for cyclase, then a temporal hierarchy between two
Ca2+-dependent mechanisms is established in addition
to an enhanced amplitude. To illustrate this point, we
calculated the putative time course of rhodopsin kinase
activation for the cases when the value for Ca2+ K1/2 is
twice smaller and twice larger than predicted by
Klenchin et al. (1995) (Fig. 8 C, dashed curves). A higher
K1/2 value makes the kinase feedback work faster than
the cyclase feedback; however, its amplitude is small.
Conversely, a lower K1/2 value makes the kinase feedback work slower than the cyclase feedback, but its amplitude becomes large. Further clarification of this issue requires a precise determination of the Ca2+ K1/2
value for the kinase feedback in vivo.
Unlike the case with guanylate cyclase, measurements of the Ca2+ K1/2 for rhodopsin kinase feedback
regulation based on electrophysiological techniques
are not available. The closest estimates are provided by
Koutalos et al. (1995b), who measured the Ca2+ dependence of PDE activation by light in truncated salamander rods. They reported a value of 400 nM for the
Ca2+ K1/2. A possible complication of their analysis is
that they likely measured a combination of two Ca2+ effects on the PDE activation: the effect mediated through
rhodopsin kinase and the effect on the rate of transducin activation by photoexcited rhodopsin originally described by Lagnado and Baylor (1994)
. A subsequent
study by Sagoo and Lagnado (1997)
indicates that this
second Ca2+ effect is based on the reduction in the affinity of transducin for GTP upon transducin activation
by rhodopsin. A Km of 9 µM GTP was observed at high
Ca2+ and 170 µM GTP at low Ca2+ while the Vmax remained unchanged. Since the GTP concentration in
intact amphibian rods remains in the millimolar range
at any level of illumination (Biernbaum and Bownds,
1985b
), this mechanism is unlikely to be physiological.
The individual impact of rhodopsin kinase regulation
and the modulation of transducin activation rate on
the overall Ca2+ regulation of PDE activation measured
by Koutalos et al. (1995b)
may be distinguished by performing the analysis with millimolar GTP levels rather
than with the 100 µM GTP used by these authors.
cGMP Feedback on Transducin GTPase
As opposed to the relatively fast Ca2+ dissociation from
corresponding binding sites, the dissociation of cGMP
from the PDE noncatalytic sites is a slow process with a
rate that is dependent on whether or not PDE is activated by transducin. Since we have demonstrated that
cGMP dissociation from these sites results in the immediate acceleration of transducin GTPase, the upper and lower curves from Fig. 7 represent the slowest and fastest limits for the onset of fast GTPase. Two hypotheses
for the role of cGMP dissociation from noncatalytic
binding sites on PDE are discussed in the literature.
Yamazaki et al. (1996) suggested that cGMP dissociation from the noncatalytic sites of activated PDE contributes to cGMP restoration during the recovery phase
of the photoresponse, while Cote et al. (1994)
proposed that cGMP dissociation contributes to photoreceptor light adaptation to bright continuous light through the acceleration of transducin GTPase. We favor the hypothesis by Cote et al. (1994)
for two reasons.
First, the amount of cGMP dissociating from the noncatalytic sites of activated PDE during the time frame of
the photoresponse does not appear to be sufficient for
significant restoration of hydrolyzed cGMP. Indeed,
given the frog rod PDE Km ~100 µM, Vmax ~4,000 s1
(Dumke et al., 1994
) and free cGMP concentration in
darkness ~4 µM (Pugh and Lamb, 1993), ~150 cGMP
molecules are hydrolyzed by each activated PDE during
the frog rod photoresponse. Yet the data presented in
Fig. 7 (virtually the same as in Fig. 4 B from Yamazaki et
al., 1996
) indicate that only ~0.25 cGMP molecules per
activated PDE can dissociate from the noncatalytic sites
over the time period of four seconds, enough time for
complete photoresponse recovery.
Second, the hypothesis that cGMP dissociation from
the noncatalytic sites contributes to the speeded recovery of the photoresponse during prolonged bright
background illumination is consistent with electrophysiological measurements of Coles and Yamane (1975) and Cervetto et al. (1984)
. These authors reported a
gradual acceleration of photoresponse recovery in amphibian rods after exposure to bright background illumination for tens of seconds. This time scale is substantially slower than the reduction in intracellular Ca2+
(McCarthy et al., 1994
, 1996
; Sampath et al., 1997
), indicating that this effect is unlikely to be mediated by
Ca2+. A goal of our future experiments is to directly
correlate this physiological phenomenon with the occupancy of the noncatalytic cGMP binding sites of frog
rod PDE.
![]() |
FOOTNOTES |
---|
Address correspondence to Peter D. Calvert, Howe Laboratory/ MEEI, 243 Charles St., Boston, MA 02114. Fax: 617-573-4290; E-mail: pdcalvert{at}meei.harvard.edu
Received for publication 12 August 1997 and accepted in revised form 31 October 1997.
1 Abbreviations used in this paper: CaM, calmodulin; PDE, rod photoreceptor cGMP-phosphodiesterase; ROS, rod outer segment.We thank Drs. M. Deric Bownds and Victor I. Govardovskii for many helpful discussions, Dr. Clint L. Makino for critically reading the manuscript, and Dr. Elina R. Nekrasova and Mr. Jason Handy for assistance in rapid kinetics experiments.
This work was supported by National Institutes of Health grant EY-10336 and a grant from the Massachusetts Lions Eye Research Foundation Inc. to V.Y. Arshavsky. V.Y. Arshavsky is a recipient of a Jules and Doris Stein Professorship from Research to Prevent Blindness Inc.
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