From the Department of Biochemistry and Molecular Biology, Oregon Health and Science University, Portland, Oregon 97201
Received for publication, October 15, 2002, and in revised form, January 18, 2003
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The rhodopsin crystal structure reveals that
intradiscal loop E-2 covers the 11-cis-retinal, creating a
"retinal plug." Recently, we noticed the ends of loop E-2 are
linked by an ion pair between residues Arg-177 and Asp-190, near the
highly conserved disulfide bond. This ion pair appears biologically
significant; it is conserved in almost all vertebrate opsins and may
occur in other G-protein-coupled receptors. We report here that the
Arg-177/Asp-190 ion pair is critical for the folding and stability of
dark state rhodopsin. We find ion pair mutants that regenerate with
retinal are functionally and spectrally wild-type-like yet thermally
unstable in their dark state because of rapid hydrolysis of the retinal
Schiff base linkage. Surprisingly, Arrhenius analysis indicates that
the activation energies for the hydrolysis process are similar between
the ion pair mutants and wild-type rhodopsin. Furthermore, the ion pair mutants do not show increased reactivity toward
hydroxylamine, suggesting that their instability is not caused by an
increased exposure to bulk solvent. Our results indicate that the loop
E-2 ion pair is important for rhodopsin stability and thus suggest that
retinitis pigmentosa observed in patients with Asp-190 mutations may in
part be the result of thermally unstable rhodopsin proteins.
Rhodopsin, the dim light photoreceptor of rod cells,
is the best characterized member of the superfamily of
G-protein-coupled receptors
(GPCRs),1 (1-8). Rhodopsin
consists of a chain of 348 amino acids, approximately half of which
form a cluster of seven membrane-spanning helices located within the
membrane (Fig. 1). The rhodopsin
chromophore, 11-cis-retinal, resides in the middle of these
helices attached to lysine 296 through a protonated Schiff base linkage
(9, 10). Interactions of amino acid side chains, as well as water molecules within the chromophore-binding pocket with the retinal, result in the 500 nm absorbance maxima for dark state rhodopsin (11,
12). Dim light vision begins when the 11-cis-retinal chromophore in rhodopsin absorbs a photon and is converted to all-trans-retinal. This change in retinal configuration
initiates a series of photo-intermediates and conformational changes in the protein, culminating in a 380 nm absorbing species called metarhodopsin II (MII), the "active conformation" which is able to bind and activate the G-protein transducin (3, 6, 13).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (83K):
[in a new window]
Fig. 1.
A, two-dimensional model of
rhodopsin adapted from Ref. 2. Intradiscal ion pair residues Arg-177
and Asp-190 in loop E-2 are colored blue and red,
respectively. Residues Cys-110 and Cys-187 that form a disulfide bond
in the intradiscal region are shaded gray.
B, three-dimensional model of the rhodopsin intradiscal
domain showing the ion pair formed by residues Arg-177 and Asp-190 on
either end of loop E-2. The figure also indicates the carbonyl backbone
of residue P7 (hydrogen-bound to Arg-177), as well as the
11-cis-retinal chromophore, and the disulfide bond between
cysteine residues Cys-110 and Cys-187 in loop E-2. These same features
are present in all three rhodopsin crystal structures (2, 14, 15). The
model shown was prepared using coordinates from 1L9H (15) using the
program WebLab.
Recently, high resolution crystal structures of rhodopsin have been
obtained (2, 14, 15). These structures confirm some of the previous
hypotheses about the rhodopsin structure, such as the general
arrangement of the transmembrane helices, the locations of
the disulfide bond, and glycosylation sites (16-21). However, they
also revealed several surprises. One of the most intriguing aspects was
the high degree of order in the intradiscal loops (the equivalent to
the extracellular loops in other GPCRs and hereby denoted as such).
Especially intriguing is loop E-2, which connects helices IV and V
(residues 173-198) and forms a twisted -hairpin that lies alongside
the retinal chromophore, potentially forming a "lid" or "plug"
across the retinal-binding pocket (2, 14, 15) (Fig. 1). This unexpected
finding has led to a number of new questions. What role does the
structure of loop E-2 play in the stability and function of rhodopsin?
Does it help provide a place for retinal to bind, or does retinal
binding induce structure in loop E-2? If the loop E-2 structure is
present in the apoprotein (opsin), how does retinal get into and out of the binding pocket?
We have recently begun to address some of these questions, and in the
process noticed an ion pair, Arg-177/Asp-190, is present on the ends of
loop E-2 (Fig. 1B). This ion pair, Arg-177/Asp-190, is
conserved in almost all vertebrate
rhodopsins2 and may also be
present in other GPCRs (Fig. 2).
Furthermore, the potentially important functional role of this ion pair
is suggested by the fact that mutations at residue Asp-190 in rhodopsin are found in patients with autosomal dominant retinitis pigmentosa (ADRP) (22-26).
|
In this work we report our investigations into the structural and
functional role of the Arg-177/Asp-190 ion pair. Our primary finding is
that the ion pair helps stabilize the dark state rhodopsin structure.
We find that mutations to the ion pair either result in opsin proteins
that do not regenerate with 11-cis-retinal or, if they do
regenerate, undergo rapid retinal Schiff base hydrolysis in the dark
state. Surprisingly, the active MII signaling state and MII decay
processes are not affected by mutations at the ion pair. We
also find that the ion pair mutations do not increase the
susceptibility of Schiff base attack by the bulk solvent (as judged by
hydroxylamine reactivity assays) nor affect the activation energy
barrier for Schiff base hydrolysis. These results illustrate the
importance of loop E-2 in retinal binding, rhodopsin stability, and
retinal release processes.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials
Except where noted below, all buffers and chemicals were
purchased from either Fisher or Sigma. Protease inhibitor tablets and
GTPS were purchased from Roche Molecular Biochemicals. Dodecyl maltoside (DM) was purchased from Anatrace (Maumee, OH), and GBX red
light filters were from Eastman Kodak Co. Polystyrene columns (2-ml bed
volume) were purchased from Pierce. Frozen bovine retinas were from
J. A. Lawson Co. (Lincoln, NE). Transducin was purified from rod
outer segments as described previously (27). DNA oligonucleotides were
purchased from Qiagen/Operon (Alameda, CA). Restriction endonucleases were from New England Biolabs (Beverly, MA). 11-cis-Retinal
was a generous gift from Dr. R. Crouch (Medical University of South Carolina and NEI, National Institutes of Health). The rho1D4 antibody was purchased from the National Cell Culture Center (Minneapolis, MN).
The nonapeptide corresponding to the C terminus of rhodopsin was
acquired from the Emory University Microchemical Facility (Atlanta,
GA). Cuvettes were purchased from Uvonics (Plainview, NY). Bandpass
filters and long-pass filters were purchased from Oriel (Stratford,
CT). The 30% acrylamide/bisacrylamide solution (37.5:1) was purchased
from Bio-Rad. Goat anti-mouse (H + L) conjugated with peroxidase and
SuperSignal West Pico Luminol/Enhancer Solution were obtained from Pierce.
Buffers
The definitions of the buffers used are as follows: PBSSC (137 mM NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, and 8 mM Na2HPO4 (pH 7.2)); buffer A (1% DM and PBSSC (pH 7.2)); buffer B (2 mM ATP, 0.1% DM, 1 M NaCl, and 2 mM MgCl2 (pH 7.2)); buffer C (0.05% DM and PBSSC (pH 7.0)); buffer D (0.05% DM and 5 mM MES (pH 6.0)); buffer E (5 mM Tris-HCl, 2 mM EDTA (pH 7.2)); and buffer F (20 mM Tris-HCl (pH 7.4), 5 mM MgCl2, 1 mM EDTA (pH 7.2)).
Construction and Expression of Rhodopsin Mutants
Site-directed mutagenesis was performed using a cassette-based strategy as described previously in the pMT4 plasmid (28, 29), as well as overlap extension PCR (30) to generate EcoRI and NotI fragments containing either the R177C, R177K, R177Q or D190C, D190E, D190N mutations in the synthetic bovine rhodopsin gene (28). The sequences for the primers are as follows: R177C, 5'CGTCGGCTGGTCTTGCTACATCCCGGAG3'; R177K, 5'GCTCGTCGGCTGGTCTAAGTACATCCCGGAGGGCATGCAGTGC3': R177Q, 5'GCTCGTCGGCTGGTCTCAGTACATCCCGGAGGGCATGCAGTGC3'; D190C, 5'CTCGTGCGGGATCTGCTACTACACGCCG3'; D190E, 5'GGAGGGCATGCAGTGCTCGTGCGGGATCGAGTACTACACGCCGC3'; and D190N, 5'GGAGGGCATGCAGTGCTCGTGCGGGATCAACTACTACACGCCGC3'. Subsequent to PCR mutagenesis, the PCR fragments were subcloned into the pMT4 vector containing the synthetic gene of rhodopsin using XhoI and PstI restriction sites, and double mutants were constructed using the BsrI restriction site. Cysteine mutants were subcloned into the pMT4 plasmid theta, a synthetic gene of rhodopsin in which the potentially reactive background cysteine residues 140, 316, 322 and 323 were replaced with serines (31, 32). All mutations were confirmed by the dideoxynucleotide sequencing method. The mutant rhodopsin proteins were transiently expressed in COS-1 cells using the DEAE-dextran method, and cells were harvested 56-72 h after transfection as described previously (33, 34).
Purification of Rhodopsin Mutants
Mutant rhodopsin proteins were expressed and harvested essentially as described previously (34). Briefly, five 15-cm plates of transfected COS-1 cells were washed twice with 7 ml of cold PBSSC buffer per plate, pelleted, and subsequently resuspended in 10 ml of cold PBSSC (pH 6.5) containing 0.5 mM PMSF. The opsin mutants were then regenerated with 10 µM 11-cis-retinal at 4 °C for 1 h, and an additional 5 µM of 11-cis retinal was then added and regeneration allowed to proceed for an additional 1 h (35).
The purification of the rhodopsin mutants proceeded essentially as the
original procedure (33), except small polystyrene columns were used for
washes and elution (36). Cells were solubilized in 5 ml of buffer A
containing 0.5 mM PMSF at 4 °C for 1 h and then
centrifuged to pellet the unsolubilized fraction. The supernatant was
mixed with 200 µl of rho1D4 antibody-Sepharose beads (binding capacity ~1 µg of rhodopsin/µg of resin) in buffer B containing 0.5 mM PMSF and nutated at 4 °C for 4-5 h. The slurry
was subsequently transferred to polystyrene columns and washed once
with 50 ml of buffer C followed by a 40-ml wash with buffer D by
gravity filtration. Samples were eluted in 350-µl fractions of buffer D containing 200 µM nonapeptide corresponding to the
rho1D4 antibody epitope (the last nine amino acids of the C terminus of
rhodopsin). A spectrum of each elution fraction was recorded (described
below), and the purified samples were either used immediately or
snap-frozen in liquid N2 and stored at 80 °C.
Immunoblot Analysis of Rhodopsin Mutant Cell Membranes
COS cells expressing rhodopsin mutants were pelleted and
resuspended in 1 ml/plate of buffer E and homogenized on ice. The homogenates were then centrifuged at 40,000 × g for 45 min at 4 °C, and the pellets were washed with 5 ml of buffer F and
subsequently resuspended in buffer F. Protein concentrations of
the resuspended membrane pellets were determined by a modified Dc
protein Assay from Bio-Rad. The manufacturer's instructions were
followed except for the addition 1.45% SDS to each well. Aliquots of
the membrane preparation were snap-frozen and stored at 80 °C
until use. SDS-PAGE was performed according to Laemmli (37), using a
5% stacking gel and a 10% resolving gel. The protein bands were
electrotransferred onto Immobilon-P transfer membranes (Millipore) and
detected using the rho1D4 monoclonal antibody as described previously
(19). Protein expression levels were determined using a Bio-Rad
PhosphorImager, and pixel densities were determined using a GS-525
molecular imaging system using supplied software.
UV-visible Absorption Spectroscopy
All UV-visible absorption spectra were recorded with a Shimadzu
UV-1601 spectrophotometer at 20 °C using a bandwidth of 2 nm, a
response time of 1 s, and a scan speed of 500 nm/min unless otherwise noted. For concentration calculations, a molar extinction coefficient value (500) for WT rhodopsin was taken to be
40 600 M
1 cm
1 (38). The samples
were photobleached in buffer D by illumination for 30 s (at a 6-Hz
flash rate) with a Machine Vision Strobe light source (EG & G) equipped
with a wavelength >490-nm long-pass filter. This light treatment was
found to be adequate for full conversion of all samples. Extinction
coefficients were determined for each dark state mutant species as
described previously in buffer D at 15 °C (34, 39). The presence of
a protonated Schiff base (PSB) in the MII state for each mutant was
verified by adding H2SO4 to a pH of 1.9 immediately after photobleaching and then measuring the absorbance
spectrum to assay the presence of a spectral species at 440 nm (which
indicates a PSB) (40).
Thermal Bleaching of Rhodopsin Samples
Absorbance Measurements-- Thermal decay rates were followed by UV-visible spectroscopy in buffer D. Specific temperatures were maintained using water-jacketed cuvette holders connected to a circulating water bath. Temperature was monitored through emersion of a digital thermometer into the sample chamber, with an accuracy of approximately ±0.2 °C. Thermal stability of the mutants was determined by first measuring the samples from 650 to 250 nm at 1-min intervals at a given temperature. Thermal decay rates were then measured by monitoring the decrease of the 500 nm absorbing dark state species from these measurements over time (41-43). Base-line drift was corrected for by normalizing all spectra to an absorbance of 0 at 650 nm.
Fluorescence Measurements-- Thermal decay rates were also measured by monitoring the increase in tryptophan fluorescence at 330 nm, caused by the release of retinal from the chromophore-binding pocket (44). The experimental set up was similar to that of the retinal release assay (described below) except that the samples were not photobleached. All thermal decay data was analyzed using mono-exponential decay (absorbance experiments) or mono-exponential rise to maxima (fluorescence experiments) fitting algorithms in Sigma Plot (Jandel Scientific software).
Thermodynamic Calculations of Thermal Decay Rates
Activation energies (Ea) were determined by
applying rate data to the Arrhenius equation: k = AeEa/(RT).
Thermodynamic parameters
H
,
G
, and
G
were calculated from the rate data as described previously (41, 45).
Briefly, the following thermodynamic Equations 1-3 were used,
![]() |
(Eq. 1) |
![]() |
(Eq. 2) |
![]() |
(Eq. 3) |
Measurement of the Rate of Retinal Release and/or MII Decay by Fluorescence Spectroscopy
The MII stability was assessed by measuring the time course of
retinal release occurring after MII formation using a Photon Technologies QM-1 steady state fluorescence spectrophotometer (44).
Each measurement was carried out using 100 µl of a 0.25 µM mutant sample in buffer D, and sample temperature was
maintained as described above. After the samples were photobleached to
the MII state (see above), the retinal release measurements were
carried out at the appropriate temperature by exciting the sample for 3 s (excitation wavelength = 295 nm, 1/4-nm bandwidth
slit setting) and then blocking the excitation beam for 42 s to
avoid further photobleaching the samples. Tryptophan fluorescence
emission was monitored at 330 nm (12-nm bandwidth slit setting), and
this cycle was repeated for up to 100 min during each measurement. To
determine the t1/2 values for retinal release, experimental data were analyzed using a mono-exponential rise to maxima
fit in Sigma Plot (Jandel Scientific software). In this manner a series
of MII decay rates was obtained at 5, 10, 15, 20, 25, 30, and 35 °C,
and their rates were applied to the Arrhenius equation,
k = AeEa/(RT),
to determine the activation energy (Ea) of the
retinal release process for each mutant rhodopsin.
Determination of Transducin (GT) Activation Rates
Activation of GT by rhodopsin was monitored using
fluorescence spectroscopy at 10 °C as described previously (34,
46-48). The excitation wavelength was 295 nm (2-nm bandwidth), and
fluorescence emission was monitored at 340 nm (12-nm bandwidth).
Photobleached mutant rhodopsin (see above) was added to a concentration
of 5 nM to the reaction mixture consisting of 250 nM GT in 10 mM Tris (pH 7.2), 2 mM MgCl2, 100 mM NaCl, 1 mM dithiothreitol, and 0.01% DM, and the mixture allowed
to stir for 300 s. The reaction was then initiated with the
addition of GTPS to a final concentration of 5 µM, and
the increase in fluorescence was followed for an additional 2000 s. To calculate the initial activation rates, the slopes of the initial
fluorescence increase following GTP
S addition were determined
through the data points covering the first 60 s.
Effect of R117Q Thermal Decay on Ability to Activate Transducin
Activation of GT by rhodopsin was monitored as described above. Activation assays were first performed on freshly thawed WT and R177Q stocks. The stocks were next incubated in the dark at 37 °C to facilitate thermal decay of the 500 nm absorbing species, and aliquots were withdrawn at the indicated time points and assayed for GT activation.
Hydroxylamine Reactivity
Hydroxylamine reactivity of the dark state was determined for
purified rhodopsins by monitoring the rate of 500 nm absorbance decrease after the addition of hydroxylamine (pH 6.0) to the samples in
buffer D to a final concentration of 50 mM at the indicated temperatures (49). Base-line drift was corrected as described above
(see "Thermal Bleaching of Rhodopsin Samples").
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Rationale for Choice of Loop E-2 Ion Pair Mutations-- Amino acid mutations were constructed based on their ability to disrupt or potentially restore the ion pair charge interaction, while introducing minimal steric perturbation. Thus, residue Arg-177 was mutated to R177K (conserved charge) and R177Q (neutral substitution). Residue Asp-190 was mutated to D190E (conserved charge) and D190N (ADRP mutation, neutral charge). Mutants R177C and D190C were constructed to enable chemical modification of the single cysteine residues. We did not analyze other ADRP-associated point mutations at site Asp-190 (D190A, D190G, and D190Y), because previous reports suggest these mutations are defective in folding, trafficking, and/or chromophore binding (23, 24, 26).
Characterization of Rhodopsin Mutants--
Expressed ion pair
mutant rhodopsins were analyzed for expression levels, proper
post-translational modifications, ability to bind
11-cis-retinal, and photobleaching properties. Immunoblot analysis of mutants expressed from transfected COS cells indicates all
mutants expressed to similar levels comparable with that of wild-type
rhodopsin (Fig. 3A). Mutants
R177C, D190C, and D190E did not regenerate in our hands and were
therefore not further characterized. These mutants also had abnormal
glycosylation patterns in comparison to WT rhodopsin in that they did
not exhibit the characteristic glycosylation smear pattern when
expressed in COS cells (Fig. 3A). Furthermore, mutants
defective in chromophore binding tended to form large molecular weight
aggregates relative to both WT and other mutants (Fig.
3A).
|
Immunoblot analysis of recombinant rhodopsins purified using the rho1D4
monoclonal antibody reveal a band pattern similar to that of wild-type
rhodopsin, with an apparent molecular mass of ~40 kDa and the
characteristic heterogeneous glycosylation smear due to overexpression
in a COS cell system (Fig. 3B), (50). Mutants capable of
regenerating with 11-cis-retinal formed characteristic rhodopsin-like pigments and could be purified to obtain spectral ratios
(A280/A500) between 1.6 and 1.8. The Arg-177 and Asp-190 single point mutants exhibited normal
photobleaching behavior with respect to formation of a blue-shifted
max
380 nm species (characteristic of the MII
intermediate) (9). Acidification of these photobleached samples
generated a
max = 440 nm species, indicating the
presence of a protonated retinal Schiff base (PSB) (40). These results
are compiled in Table I, and a
representative example of the photobleaching behavior is shown for
mutants R177Q and D190N (Fig.
4A). Similar to the single
mutants R177Q and D190N, the R177Q/D190N double mutant shows wild-type
behavior in terms of expression levels, post-translational
modifications, and chromophore binding. However, it did exhibit
perturbed photobleaching properties. Although capable of forming both a
spectral MII species and a PSB, following illumination a residual
species with a
max of ~480 nm persisted up to 10 h after illumination (data not shown). The cause of this is not known,
although similar effects have been reported for other rhodopsin
point mutations such as G90S and L226C (29, 34, 51).
|
|
Retinal Release Rates and Activation Energies for Metarhodopsin II Decay Measured by Fluorescence Spectroscopy-- To determine potential effects the ion pair mutations may have on the stability of the MII active signaling species of rhodopsin, the activation energies for retinal release were determined. The rate of retinal release occurring during the decay of the MII species was measured using a fluorescence-based assay at 20 °C (44). Under the conditions used for this assay, the t1/2 of retinal release for WT rhodopsin at 20 °C in buffer D was 13 ± 0.5 min (n = 3), comparable with the 13-15-min values reported previously (34, 43, 48, 52) for both ROS-purified and COS-expressed rhodopsin. Somewhat unexpectedly, the corresponding t1/2 values for the ion pair mutant rhodopsins were similar to that of WT rhodopsin. The values for each of the mutants are compiled in Table I. The activation energy for the metarhodopsin II decay process was obtained by monitoring the rate of fluorescence increase in buffer D at seven different temperatures (5, 10, 15, 20, 25, 30, and 35 °C). The rate of fluorescence increase in all cases was temperature-dependent, and Arrhenius plots of these measurements indicated a temperature-dependent linear relationship for all mutants (Fig. 4B). From these plots an activation energy (Ea) of 20.2 kcal/mol was obtained for purified WT rhodopsin in DM, in good agreement with values reported previously (34, 44, 53). Arrhenius plots of the retinal release rates for the ion pair mutants show nearly equal Ea values (Fig. 4B and Table I).
Transducin Activation by Ion Pair Mutants--
To assess the
potential functional effects, the ion pair mutants were tested for
their ability to activate transducin using a fluorescence-based assay
that measures the increase in tryptophan fluorescence of the
GT-GTP
S species (46, 48, 54). All ion pair mutations
that regenerated with retinal are functionally active, and
representative examples are presented in Fig. 4C. The
results for transducin activation are compiled in Table I as initial
rates of fluorescence increase relative to WT rhodopsin.
Thermal Stability in the Dark State--
The most dramatic
perturbation induced by the ion pair mutations was on the stability of
the dark state structure. Thermal stabilities of dark state WT and ion
pair mutant rhodopsins were determined by measuring the loss of the 500 nm absorbing species over time as described under "Experimental
Procedures." An example of this assay is depicted for mutant R177Q at
37 °C in Fig. 5A. The loss
of the 500 nm species directly correlates with a loss of ability to
activate transducin (Fig. 5, B and C).
Additionally, the decrease in absorbance at 500 nm reflects a loss of
the chromophore Schiff base linkage as judged by decay of the
acid-denatured 440 nm species over the duration of the thermal decay
assay (Fig. 5, D and E). Furthermore, we conclude
the retinal is leaving the chromophore binding pocket after the
hydrolysis because the rate of the loss of the 500 nm absorbing species
correlates with the rate of tryptophan fluorescence increase, and
irradiation of the sample with light following a plateau in signal does
not cause a further fluorescence increase (Fig. 5F)
(44).
|
All of the ion pair mutants showed significantly expedited
rates of thermal decay in comparison to WT rhodopsin as judged by their
loss in absorbance at 500 nm and increase in fluorescence at 330 nm
(Table II). A comparison of the thermal
decay rates at 55 °C monitored by absorbance is shown in Fig.
6. Note that the thermal decay rate of
ROS-purified rhodopsin was similar to that of WT recombinant rhodopsin
purified from COS cells (38.5 ± 3.0 and 37 min at 55 °C,
respectively). The activation energies for the thermal absorbance decay
processes were determined by monitoring the loss of the 500 nm
absorbing species over time at 7 different temperatures (37, 41, 45, 47.5, 50, 52.5, and 55 °C). In all cases, the rate of loss in 500 nm
absorbance was temperature-dependent, and Arrhenius plots
indicate a similar temperature-dependent relationship for all
mutants (Fig. 7). The Arrhenius plots are
clearly concave, suggesting at least two different rate-limiting
processes may occur during the temperature-dependent absorbance
decay. With this in mind, two linear regressions were used to
approximate the activation energies for the two apparent processes
(55-47.5 and 47.5-37 °C, respectively). From this analysis, the
Ea for WT rhodopsin was determined to be ~16
kcal/mol at 37 °C and 103 kcal/mol at 55 °C. The thermodynamic
parameters Ea, G
,
H
, and
G
were estimated from the rate data for WT and mutant rhodopsins using
equations described previously (see Table II) (41, 45).
|
|
|
Hydroxylamine Reactivity--
Hydroxylamine reactivity experiments
showed that the ion pair mutants were not more susceptible to
hydroxylamine in the dark state. These assays were carried out for
purified ROS rhodopsin and each mutant sample, and the decay of the
dark state 500 nm absorbing species was monitored in buffer D at either
20, 37, or 55 °C over time following the addition of hydroxylamine
(pH 6.0) to a final concentration of 50 mM. WT rhodopsin
purified from retinal sources and from expressed COS cells was found to be inert to hydroxylamine in the dark state, as described previously (49). Intriguingly, none of the ion pair mutants exhibited any increased reactivity toward hydroxylamine treatment in the dark state
at 20, 37, and 50 °C (Fig. 8).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Early studies by Khorana and others (50, 55-59) led to the
hypothesis that the rhodopsin intradiscal domain plays a crucial role
in maintaining proper protein folding, correct post-translational modifications, trafficking, and 11-cis-retinal binding.
Consistent with this theory, a number of point mutations that naturally
occur in this region result in ADRP, an inherited human disease causing retina degradation (23, 24, 26, 60-62). As noted in the Introduction, the rhodopsin crystal structures reveal a high degree of order and
structure in the intradiscal region. This region of the protein is
proposed to be structurally critical for maintaining the electrostatic and hydrogen-bonded network surrounding the retinal chromophore (15).
Most notably, loop E-2 forms a twisted -sheet which lies across the
retinal chromophore (2, 14, 15). Through analysis of the loop E-2
region we noticed an ion pair Arg-177/Asp-190 is present on either end
of this loop structure. Additionally, we noticed that residue Arg-177
is hydrogen-bonded to the backbone carbonyl of residue P7, a residue
found at the turn of loop E-1 in the N terminus of the proteins. The
fact that residues Arg-177 and Asp-190 interact with many
residues within the intradiscal region of rhodopsin suggests that the
ion pair may play a significant role in maintaining the structural
integrity of the "retinal plug" domain. The published rhodopsin
crystal structures show very little difference in the region around the
Arg-177/Asp-190 ion pair (analysis of all three structures using the
program Swiss-PDB Viewer shows that the 87 amino acid side chains
within 16 Å of Arg-177 show a root mean square deviation of 1.0 Å or
less (2, 14, 15)). However, we do notice a difference in the placement
of residue Asn-200, which exhibits a different rotameric flip between
structures 1HZX, 1L9H, and 1F88, and thus exhibits alternate
hydrogen bonding to residue Asp-190 in the different structures (2, 14,
15). The present report details our studies on the structural and
functional effects caused by disrupting the Arg-177/ Asp-190
intradiscal ion pair located on either end of loop E-2 (Fig.
1B).
General Characteristics of Mutants-- The majority of the single ion pair mutants we created, expressed to similar levels comparable with WT rhodopsin, underwent proper glycosylation (as judged by whole cell lysate immunoblotting, Fig. 3) and regenerated with 11-cis-retinal. Additionally, with the exceptions of mutants R177C, D190C, and D190E, the mutants were properly folded, as judged by their ability to bind the 11-cis-retinal chromophore, and produced a wild type like A280/A500 ratio (Table I). The fact that most of the Asp-190 mutations were unable to bind retinal is in agreement with previous reports of other Asp-190 mutations, which also were found to be defective in retinal binding (23, 26, 58). One possible reason for the sensitivity of this site to mutations may be that residue Asp-190 is partially buried and makes contacts with several residues (Ile-189 and Tyr-Y191), which form part of the retinal binding pocket (Fig. 1B) (2, 15, 52). Abrogation of these contacts by mutations to residue Asp-190 may thus distort the retinal-binding pocket thereby making it inaccessible or sterically unfavorable for proper binding of 11-cis-retinal. Additionally, it is also possible that the R177C and D190C mutants are not able to regenerate with 11-cis-retinal because improper disulfide bonds are formed in the final folded structure, as these residues are in close proximity to the conserved Cys-110/Cys-187 disulfide pair, as well as residue Cys-185 (Fig. 1B). This explanation is supported by recent findings, which show that improper disulfide bonds form as a result of mutations to this region (63-65). Furthermore, the fact that the glycosylation patterns of whole cell lysate immunoblots of mutants R177C, D190C, and D190E are different suggests the misfolding may have already occurred by the time the protein reached the endoplasmic reticulum, and thus the lack of regeneration is not simply due to an ultra-fast rate of retinal hydrolysis after regeneration (Fig. 3A). These three mutants also tend to form high molecular weight aggregates when analyzed on SDS gels, and such aggregation has been suggested to result in proteins that eventually become degraded in the endoplasmic reticulum rather than proceeding through the Golgi to the plasma membrane (24, 66).
The spectral behaviors for the mutants that did regenerate with
11-cis-retinal were WT-like in their ability to form an MII absorbing species and a PSB upon acid denaturation (Fig.
4A). One exception was the double mutant R177Q/D190N,
although WT-like in its MII and PSB characteristics, which demonstrated
a residual absorbing species with a max of ~480 nm
following illumination (data not shown). This ~480 nm absorbing
species may be an MII-like intermediate containing a PSB, as was
suggested for mutations at site Gly-90 (34, 67, 68), or it may be an
MIII intermediate, due to an altered equilibrium between the MII and
MIII states (69, 70). However, why the double mutation shows this
characteristic and the single mutants do not is unknown.
The Arg-177/Asp-190 Ion Pair Is Not Required for MII Formation, Signaling, or Stability-- We were surprised to find that the Arg-177/Asp-190 ion pair does not appear critical for the formation or maintaining the stability of the MII state. All ion pair mutants show retinal release rates similar to that of WT rhodopsin in buffer D at 20 °C (13 ± 0.5 min, Table I), and activation energies (Ea) for retinal release from MII are almost identical to the 20.2 kcal/mol obtained for WT rhodopsin (Fig. 4B and Table I) (34, 44, 53). These results are in contrast to our previous findings on mutations within the chromophore-binding pocket, which increased the retinal release rate from MII (although the activation energy for retinal release were unaffected (34)). Notably, disruption of the Cys-110/Cys-187 disulfide bond in the intradiscal region of rhodopsin dramatically disrupts MII decay and stability (41). In contrast, the regenerated ion pair mutants all show normal MII function and decay, suggesting they must still contain a normal Cys-110/Cys-187 disulfide bond. Furthermore, the ion pair mutants are functionally active, as they could activate the G-protein transducin, although not quite to the full extent of WT rhodopsin (Fig. 4C and Table I). Taken together, these results suggest that the intradiscal ion pair is not critical for the formation or stability of the active signaling MII state.
Arg-177/Asp-190 Ion Pair Helps Stabilize the Dark State Rhodopsin Structure-- The most dramatic effect exhibited by the ion pair mutants was a sharp decrease in the stability of their dark state structures in comparison to WT rhodopsin. Recent studies (43, 71, 72) have begun to address carefully the thermodynamics of rhodopsin protein stability. In the present work, our conclusions about rhodopsin thermal stability are primarily based on monitoring the stability of the retinal Schiff base linkage, which we measure as the loss of absorbance at 500 nm. We feel confident that these measurements report on Schiff base hydrolysis and release of retinal from the binding pocket, for the following reasons. The loss in absorbance at 500 nm correlates with both the loss of the PSB (as observed by a decrease in the acid-denatured 440 nm species over time) as well as the increase in tryptophan fluorescence at 330 nm (Fig. 5 and Table II). Furthermore, the loss of absorbance at 500 nm also correlates with the loss in ability of the protein to activate transducin (Fig. 5C). Taken together these results indicate that during the thermal decay of the dark state structure the Schiff base is hydrolyzed and the retinal leaves the chromophore-binding pocket.
Arrhenius Analysis Indicates the Thermal Decay in Rhodopsin May
Occur through More Than One Pathway--
Interestingly, the Arrhenius
analysis of the thermal decay rates shows a concave plot, with all of
the mutants decaying much faster than WT rhodopsin. To our knowledge,
this behavior has not been reported previously for rhodopsin. Concave
Arrhenius plots can be attributed to several factors, although the most common interpretation is that at least two different rate-limiting steps are involved (73). With this interpretation in mind, we fit our
WT rhodopsin data assuming two different activation energies may be
present, and we find Ea values of ~16 kcal/mol for
the lower temperature range (37-47.5 °C) and ~103 kcal/mol for
the higher temperature range (47.5-55 °C). As shown in Table II,
the ion pair mutants all showed similar Ea values ranging from 17 to 31 kcal/mol for the lower temperature range and
94-107 kcal/mol for the higher temperature range. Finally, thermodynamic analysis of the Arrhenius data show that the ion pair and
WT rhodopsins have similar H
values (see
Table II), suggesting the perturbation caused by the mutations are
generally entropic in nature (34, 41, 74).
It is informative to compare the thermal decay activation energies from this study with previous findings. For example, our Ea of 103 kcal/mol for WT rhodopsin (at the higher temperatures) is in excellent agreement with the 102.1 ± 5.8 kcal/mol value reported previously by Khorana and co-workers (41) for WT using a similar experimental set up. Furthermore, the Ea value of 16 kcal/mol we observe at the lower temperatures is similar to the 20.2 kcal/mol value obtained for the retinal release process that occurs during MII decay (34, 44), and to the Ea of hydrolysis for model Schiff base retinal compounds (75), as well as the Ea for retinal binding in rhodopsin (76).
What might be the cause of the different Ea values obtained for the Arrhenius analysis at different temperatures? The higher activation energy barriers at higher temperatures may reflect the less favorable conditions for hydrolysis of the Schiff base linkage present in the interior of unfolded rhodopsin. In other words, thermal denaturation of the protein occurs which may reposition key amino acids involved in Schiff base formation and hydrolysis, thereby increasing the Ea observed at the higher temperatures. Alternatively, the lower Ea observed at the lower temperatures suggests that at these temperatures another process also contributes to Schiff base hydrolysis, one that occurs more efficiently at lower temperatures (such as proton tunneling) than the process that dominates at higher temperatures. Another possibility is that the pre-exponential factor in the Arrhenius analysis has changed; the pre-exponential factor is related to steric factors and/or the efficiency with which the collisions lead to a productive reaction (73).
Thermodynamic Significance of the Arg-177/Asp-190 Ion
Pair--
The G
values for the ion
pair mutants are approximately
1.5 to
2.4 kcal/mol, similar to the
2.9 ± 1.2 kcal/mol reported for the C110A/C187A mutation of the
critical rhodopsin disulfide bond (41). Thus, in terms of dark state
stability, abrogation of the ion pair results in free energy changes
comparable with those observed for the loss of the Cys-110/Cys-187
disulfide bond. This is perhaps not surprising, because the
Arg-177/Asp-190 ion pair is in close proximity to the intradiscal
Cys-110/Cys-187 disulfide bond (Fig. 1B). However, it is
important to note that the ion pair mutations we report here apparently
have little effect on the stability of the MII structure, in contrast
to the disulfide bond that appears crucial for MII stability (41). This
latter point is of interest; to the best of our knowledge, the
Arg-177/Asp-190 ion pair mutants represent a previously unidentified
class of rhodopsin mutants which alter the stability and structure of
the dark state yet have little to no effect on the stability of the MII
state. One explanation for this phenomena may be that the loop E-2
region changes structure during MII formation and the ion pair is no
longer present in the MII state, thus the stability of MII would not be
affected by mutations to the Arg-177/Asp-190 ion pair. We note that
although the Arg-177 and Asp-190 residues point away from the helical
bundle, it is also possible that mutations to this ion pair may affect
the positioning or orientation of helices 5 and 6. However, there is
some precedence for conformational changes in loop E-2, as Ridge
et al. (77) have previously shown residue Cys-185 becomes
accessible to chemical labeling only in the MII state.
Disrupting the Arg-177/Asp-190 Ion Pair Does Not Appear to Increase the Exposure of the Retinal Schiff Base to Bulk Solvent-- We initially interpreted our results to mean that disrupting the Arg-177/Asp-190 ion pair causes a loosening of the loop E-2 structure, and thus we hypothesized the increased rate of Schiff base hydrolysis was due its increased exposure to external solvent. However, to our surprise, we found that the presence of 50 mM hydroxylamine had no effect on any of the mutants tested, even when tested at three different temperatures (Fig. 8).3 The lack of increased hydroxylamine sensitivity argues against the hypothesis that the thermal instability induced in the ion pair mutants is due to a structural perturbation that renders their Schiff base more susceptible to attack by the bulk solvent. Rather, the hydroxylamine data suggest that the Arg-177/Asp-190 ion pair in the intradiscal domain of rhodopsin mediates thermal stability of the dark state structure through some other mechanism.
Speculation on the Role of Arg-177/Asp-190 in Stabilizing Rhodopsin-- Previous chemical models of the retinal-binding and release pathways have speculated that the protonated retinal Schiff base linkage can spontaneously hydrolyze and thus is in dynamic equilibrium with retinal covalently bound to rhodopsin (75, 78). One interpretation of our data may be that the ion pair stabilizes the retinal plug structure. If the retinal plug functions to block the release of free 11-cis-retinal produced through spontaneous hydrolysis (through a steric mechanism which confines the transiently hydrolyzed retinal to the chromophore binding pocket), disruption of this structure might lead to an apparent increase in Schiff base hydrolysis rates. In this scenario the function of the retinal plug is to effectively force the transiently formed free 11-cis-retinal to remain in the binding pocket and reform a Schiff base linkage with rhodopsin. Alternatively, the ion pair may enhance rhodopsin stability by constraining the conformation of a network of water molecules and residues attached to loop E-2 (Glu-181 to Ser-186 to Cys-187) that directly link loop E-2 with the retinal Schiff base through residue Glu-113 (15, 52). Restraining the flexibility of this region may thus make rhodopsin more stable by inhibiting transient formation of the tetrahedral carbinolamine intermediate thought to be involved in the transition state of Schiff base hydrolysis (75, 79). We are presently carrying out further experiments to test these hypotheses.
Conclusions--
The Arg-177/Asp-190 ion pair located on either
end of intradiscal loop E-2 appears to be important for maintaining
dark state rhodopsin stability, although it does not appear to play a
critical role in formation or stability of the active MII species.
These results illustrate the importance of the rhodopsin structure
revealed by x-ray crystallography (2, 14, 15). With the structure of
rhodopsin in hand, it is now possible to assess the previously unappreciated functional role of interactions that occur within the
protein to provide receptor stability and allow receptor activation and attenuation.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. John Denu, Dr. Dan Oprian, Dr. Kevin Ridge, Dr. Mark Krebs, and numerous members of the 10th International Conference on Retinal Proteins for helpful discussions.
![]() |
FOOTNOTES |
---|
* This work was supported in part by Grants EY12095-01 (to D. L. F.) and T32-EY07123-09 (to J. M. J.) from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Oregon Health and
Science University, Mail Code L224, 3181 S.W. Sam Jackson Park Rd.,
Portland, OR 97201-3098. Tel.: 503-494-0583; Fax: 503-494-8393; E-mail: farrensd@ohsu.edu.
Published, JBC Papers in Press, January 23, 2003, DOI 10.1074/jbc.M210567200
2 Vertebrate rhodopsin alignments were carried out using ExPASy (Expert Protein Analysis System) proteomics server of the Swiss Institute of Bioinformatics (us.expasy.org). The sole exception was sheep rhodopsin, which contains a leucine residue at site 190 (Leu-190) and, interestingly, a glutamine residue at site 181 (Gln-181).
3 Normally, if solvent-accessible, a retinal Schiff base is rapidly cleaved by hydroxylamine, and this property has frequently been used as a measure of accessibility of the retinal Schiff base linkage in rhodopsin (41, 42, 52, 68, 72).
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
GPCR, G-protein-coupled receptor;
ADRP, autosomal dominant retinitis
pigmentosa;
DM, n-dodecyl--maltoside;
E-2, second
extracellular loop of rhodopsin;
Ea, energy of
activation;
G-protein, guanine nucleotide-binding regulatory protein;
GT, heterotrimeric G-protein transducin;
GT
,
subunit of transducin G-protein;
GTP
S, guanosine
5'-3-O-(thio)triphosphate;
MES, 2-(N-morpholino)ethanesulfonic acid monohydrate;
MII, metarhodopsin II;
PMSF, phenylmethylsulfonyl fluoride;
PSB, protonated
Schiff-base;
ROS, rod outer segment;
WT, wild-type.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Khorana, H. G.
(1992)
J. Biol. Chem.
267,
1-4 |
2. |
Palczewski, K.,
Kumasaka, T.,
Hori, T.,
Behnke, C. A.,
Motoshima, H.,
Fox, B. A.,
Le Trong, I.,
Teller, D. C.,
Okada, T.,
Stenkamp, R. E.,
Yamamoto, M.,
and Miyano, M.
(2000)
Science
289,
739-745 |
3. | Meng, E. C., and Bourne, H. R. (2001) Trends Pharmacol. Sci. 22, 587-593[CrossRef][Medline] [Order article via Infotrieve] |
4. | Okada, T., Ernst, O. P., Palczewski, K., and Hofmann, K. P. (2001) Trends Biochem. Sci. 26, 318-324[CrossRef][Medline] [Order article via Infotrieve] |
5. | Ebrey, T., and Koutalos, Y. (2001) Prog. Retin. Eye Res. 20, 49-94[CrossRef][Medline] [Order article via Infotrieve] |
6. | Sakmar, T. P., Menon, S. T., Marin, E. P., and Awad, E. S. (2002) Annu. Rev. Biophys. Biomol. Struct. 31, 443-484[CrossRef][Medline] [Order article via Infotrieve] |
7. | Albert, A. D., and Yeagle, P. L. (2002) Biochim. Biophys. Acta 1565, 183-195[Medline] [Order article via Infotrieve] |
8. | Filipek, S., Stenkamp, R. E., Teller, D. C., and Palczewski, K. (2002) Annu. Rev. Physiol. 65, 851-879[CrossRef][Medline] [Order article via Infotrieve] |
9. | Wald, G. (1968) Nature 219, 800-807[Medline] [Order article via Infotrieve] |
10. | Hargrave, P. A., Hamm, H. E., and Hofmann, K. P. (1993) Bioessays 15, 43-50[Medline] [Order article via Infotrieve] |
11. | Neitz, M., Neitz, J., and Jacobs, G. H. (1991) Science 252, 971-974[Medline] [Order article via Infotrieve] |
12. | Kochendoerfer, G. G., Lin, S. W., Sakmar, T. P., and Mathies, R. A. (1999) Trends Biochem. Sci. 24, 300-305[CrossRef][Medline] [Order article via Infotrieve] |
13. | Lewis, J. W., and Kliger, D. S. (2000) Methods Enzymol. 315, 164-178[CrossRef][Medline] [Order article via Infotrieve] |
14. | Teller, D. C., Okada, T., Behnke, C. A., Palczewski, K., and Stenkamp, R. E. (2001) Biochemistry 40, 7761-7772[CrossRef][Medline] [Order article via Infotrieve] |
15. |
Okada, T.,
Fujiyoshi, Y.,
Silow, M.,
Navarro, J.,
Landau, E. M.,
and Shichida, Y.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
5982-5987 |
16. | Unger, V. M., Hargrave, P. A., Baldwin, J. M., and Schertler, G. F. (1997) Nature 389, 203-206[CrossRef][Medline] [Order article via Infotrieve] |
17. | Schertler, G. F., and Hargrave, P. A. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 11578-11582[Abstract] |
18. | Baldwin, J. M., Schertler, G. F., and Unger, V. M. (1997) J. Mol. Biol. 272, 144-164[CrossRef][Medline] [Order article via Infotrieve] |
19. | Karnik, S. S., Sakmar, T. P., Chen, H. B., and Khorana, H. G. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 8459-8463[Abstract] |
20. |
Karnik, S. S.,
and Khorana, H. G.
(1990)
J. Biol. Chem.
265,
17520-17524 |
21. | Fukuda, M. N., Papermaster, D. S., and Hargrave, P. A. (1982) Methods Enzymol. 81, 214-223[Medline] [Order article via Infotrieve] |
22. | Nathans, J. (1990) Biochemistry 29, 9746-9752[Medline] [Order article via Infotrieve] |
23. | Sung, C. H., Schneider, B. G., Agarwal, N., Papermaster, D. S., and Nathans, J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 8840-8844[Abstract] |
24. | Sung, C. H., Davenport, C. M., Hennessey, J. C., Maumenee, I. H., Jacobson, S. G., Heckenlively, J. R., Nowakowski, R., Fishman, G., Gouras, P., and Nathans, J. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 6481-6485[Abstract] |
25. | Sung, C. H., Makino, C., Baylor, D., and Nathans, J. (1994) J. Neurosci. 14, 5818-5833[Abstract] |
26. | Kaushal, S., and Khorana, H. G. (1994) Biochemistry 33, 6121-6128[Medline] [Order article via Infotrieve] |
27. |
Baehr, W.,
Morita, E. A.,
Swanson, R. J.,
and Applebury, M. L.
(1982)
J. Biol. Chem.
257,
6452-6460 |
28. | Ferretti, L., Karnik, S. S., Khorana, H. G., Nassal, M., and Oprian, D. D. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 599-603[Abstract] |
29. | Yang, K., Farrens, D. L., Hubbell, W. L., and Khorana, H. G. (1996) Biochemistry 35, 12464-12469[CrossRef][Medline] [Order article via Infotrieve] |
30. | Fong, T. M. (1999) in Structure-Function Analysis Of G Protein-coupled Receptors (Wess, J., ed) , pp. 1-20, Wiley-Liss, Inc., New York |
31. | Resek, J. F., Farahbakhsh, Z. T., Hubbell, W. L., and Khorana, H. G. (1993) Biochemistry 32, 12025-12032[Medline] [Order article via Infotrieve] |
32. | Resek, J. F., Farrens, D., and Khorana, H. G. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 7643-7647[Abstract] |
33. | Oprian, D. D., Molday, R. S., Kaufman, R. J., and Khorana, H. G. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 8874-8878[Abstract] |
34. | Janz, J. M., and Farrens, D. L. (2001) Biochemistry 40, 7219-7227[Medline] [Order article via Infotrieve] |
35. |
Reeves, P. J.,
Hwa, J.,
and Khorana, H. G.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
1927-1931 |
36. |
Dunham, T. D.,
and Farrens, D. L.
(1999)
J. Biol. Chem.
274,
1683-1690 |
37. | Laemmli, U. K. (1970) Nature 227, 680-685[Medline] [Order article via Infotrieve] |
38. |
Wald, G. B.,
and Brown, P. K.
(1953)
J. Gen. Physiol.
37,
189-200 |
39. | Sakmar, T. P., Franke, R. R., and Khorana, H. G. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 8309-8313[Abstract] |
40. | Sakamoto, T., and Khorana, H. G. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 249-253[Abstract] |
41. | Davidson, F. F., Loewen, P. C., and Khorana, H. G. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 4029-4033[Abstract] |
42. |
Andres, A.,
Kosoy, A.,
Garriga, P.,
and Manyosa, J.
(2001)
Eur. J. Biochem.
268,
5696-5704 |
43. | Vogel, R., and Siebert, F. (2002) Biochemistry 41, 3529-3535[CrossRef][Medline] [Order article via Infotrieve] |
44. |
Farrens, D. L.,
and Khorana, H. G.
(1995)
J. Biol. Chem.
270,
5073-5076 |
45. | Segel, I. H. (1975) Enzyme Kinetics , pp. 931-941, Wiley Interscience, New York |
46. |
Phillips, W. J.,
and Cerione, R. A.
(1988)
J. Biol. Chem.
263,
15498-15505 |
47. | Fahmy, K., Zvyaga, T. A., Sakmar, T. P., and Siebert, F. (1996) Biochemistry 35, 15065-15073[CrossRef][Medline] [Order article via Infotrieve] |
48. |
Farrens, D. L.,
Altenbach, C.,
Yang, K.,
Hubbell, W. L.,
and Khorana, H. G.
(1996)
Science
274,
768-770 |
49. | Sakmar, T. P., Franke, R. R., and Khorana, H. G. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 3079-3083[Abstract] |
50. | Kaushal, S., Ridge, K. D., and Khorana, H. G. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 4024-4028[Abstract] |
51. | Zhukovsky, E. A., and Oprian, D. D. (1989) Science 246, 928-930[Medline] [Order article via Infotrieve] |
52. | Yan, E. C., Kazmi, M. A., De, S., Chang, B. S., Seibert, C., Marin, E. P., Mathies, R. A., and Sakmar, T. P. (2002) Biochemistry 41, 3620-3627[CrossRef][Medline] [Order article via Infotrieve] |
53. | Hubbard, R., Bownds, D., and Yoshizawa, T. (1965) Cold Spring Harbor Symp. Quant. Biol. 30, 301-315[Medline] [Order article via Infotrieve] |
54. | Fahmy, K., and Sakmar, T. P. (1993) Biochemistry 32, 7229-7236[Medline] [Order article via Infotrieve] |
55. | Doi, T., Molday, R. S., and Khorana, H. G. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 4991-4995[Abstract] |
56. | Karnik, S. S., Ridge, K. D., Bhattacharya, S., and Khorana, H. G. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 40-44[Abstract] |
57. |
Borjigin, J.,
and Nathans, J.
(1994)
J. Biol. Chem.
269,
14715-14722 |
58. |
Liu, X.,
Garriga, P.,
and Khorana, H. G.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
4554-4559 |
59. |
Cha, K.,
Reeves, P. J.,
and Khorana, H. G.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
3016-3021 |
60. |
Sung, C. H.,
Davenport, C. M.,
and Nathans, J.
(1993)
J. Biol. Chem.
268,
26645-26649 |
61. | Jacobson, S. G., Kemp, C. M., Sung, C. H., and Nathans, J. (1991) Am. J. Ophthalmol. 112, 256-271[Medline] [Order article via Infotrieve] |
62. | Kemp, C. M., Jacobson, S. G., Roman, A. J., Sung, C. H., and Nathans, J. (1992) Am. J. Ophthalmol. 113, 165-174[Medline] [Order article via Infotrieve] |
63. |
Hwa, J.,
Garriga, P.,
Liu, X.,
and Khorana, H. G.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
10571-10576 |
64. |
Hwa, J.,
Reeves, P. J.,
Klein-Seetharaman, J.,
Davidson, F.,
and Khorana, H. G.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
1932-1935 |
65. |
Hwa, J.,
Klein-Seetharaman, J.,
and Khorana, H. G.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
4872-4876 |
66. |
Illing, M. E.,
Rajan, R. S.,
Bence, N. F.,
and Kopito, R. R.
(2002)
J. Biol. Chem.
277,
34150-34160 |
67. | Rao, V. R., Cohen, G. B., and Oprian, D. D. (1994) Nature 367, 639-642[CrossRef][Medline] [Order article via Infotrieve] |
68. | Zvyaga, T. A., Fahmy, K., Siebert, F., and Sakmar, T. P. (1996) Biochemistry 35, 7536-7545[CrossRef][Medline] [Order article via Infotrieve] |
69. | Reuter, T. (1976) Vision Res. 16, 909-917[CrossRef][Medline] [Order article via Infotrieve] |
70. | Lewis, J. W., van Kuijk, F. J., Carruthers, J. A., and Kliger, D. S. (1997) Vision Res. 37, 1-8[CrossRef][Medline] [Order article via Infotrieve] |
71. | Landin, J. S., Katragadda, M., and Albert, A. D. (2001) Biochemistry 40, 11176-11183[CrossRef][Medline] [Order article via Infotrieve] |
72. | Vogel, R., and Siebert, F. (2002) Biochemistry 41, 3536-3545[CrossRef][Medline] [Order article via Infotrieve] |
73. | Swinbourne, E. S. (1971) in Analysis of Kinetic Data (Agosta, W. A., ed) , pp. 60-65, Thomas Nelson and Sons Ltd., Nairobi, Kenya |
74. | Jung, K. H., Spudich, E. N., Dag, P., and Spudich, J. L. (1999) Biochemistry 38, 13270-13274[CrossRef][Medline] [Order article via Infotrieve] |
75. | Cooper, A., Dixon, S., Nutley, M., and Robb, J. (1987) J. Am. Chem. Soc. 109, 7254-7263 |
76. | Hiroyuki Matsumoto, K. H., and Yoshizawa, T. (1978) Biochim. Biophys. Acta 501, 257-268[Medline] [Order article via Infotrieve] |
77. | Ridge, K. D., Lu, Z., Liu, X., and Khorana, H. G. (1995) Biochemistry 34, 3261-3267[Medline] [Order article via Infotrieve] |
78. | Cooper, A., and Converse, C. A. (1976) Biochemistry 15, 2970-2978[Medline] [Order article via Infotrieve] |
79. | Harosi, F. I., and Sandorfy, C. (1995) Photochem. Photobiol. 61, 510-517 |