From the Departments of Biochemistry and Molecular Biology, Oregon Health Sciences University, Portland, Oregon 97201-3098
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ABSTRACT |
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A recent proposal for the formation of
functionally active rhodopsin has placed critical importance on a
movement of one of its transmembrane helices (Farrens, D. L.,
Altenbach, C., Yang, K., Hubbell, W. L., and Khorana, H. G. (1996) Science 274, 768-770). We investigated this
hypothesis using a series of eight rhodopsin mutants containing single
reactive cysteine residues in the region (helix F) where movement was
previously detected. The cysteine mutants were studied in two ways, by
measuring their reactivity to a cysteine-specific reagent
(PyMPO-maleimide), and by labeling the cysteines with a fluorescent
label (monobromobimane) followed by fluorescence spectroscopic
analysis. The chemical reactivity data showed sequence-specific
variations in reactivity for the mutants in the dark state, resulting
in a pattern suggestive of an G-protein-coupled receptors
(GPCRs)1 are membrane
proteins that act as the initial input stages in many sensory systems.
Arguably, the best characterized GPCR is the bovine visual
photoreceptor, rhodopsin (1). Approximately half the amino acids in
rhodopsin are found in a cluster of seven membrane-spanning helices
(see Fig. 1). The rhodopsin "antagonist"
(11-cis-retinal) is covalently bound in the middle of these
helices, inactivating the protein in the dark state. Rhodopsin becomes
activated when it absorbs light (absorption A central question remains as to why only the MII form of rhodopsin can
bind and activate transducin. It has been noted that transducin can be
activated by peptide analogues of the cytoplasmic loops of rhodopsin
(3), suggesting that dark state rhodopsin is inactive because the loops
are physically inaccessible. Thus, a reasonable hypothesis is that MII
formation involves some kind of conformational change that exposes the
cytoplasmic loops and allows transducin to bind and become activated.
Recently, the nature of these conformational changes has been studied
by Khorana, Hubbell, and co-workers using an approach called
site-directed spin labeling (SDSL). Briefly, their SDSL studies
involved constructing mutant rhodopsin proteins with unique reactive
cysteine residues in the cytoplasmic domain in loops C-D and E-F (4,
5), labeling these cysteines with a methanethiosulfonate spin label,
and then measuring the electron paramagnetic resonance (EPR) spectrum
of the labeled mutant in the dark state and MII state. By assessing the
mobility and environment of each spin label, a probable structure of
the cytoplasmic domain could be proposed. Interestingly, their
experiments suggested that many conformational changes in MII occur
near helix F (6, 7).2 A model
was proposed, suggesting MII formation involves an outward movement of
this helix away from helix C (7, 8). The potential importance of this
movement was implied from observations that linking helix F to helix C
(either by disulfide bonds or metal chelating agents) abolished
transducin activation (8, 9). Furthermore, a wealth of data is
accumulating that suggests a movement of this helix plays a crucial
role in forming the active, signaling state in other GPCRs.
In this paper, we directly tested the hypothesis that helix F movement
occurs in rhodopsin upon MII formation. We measured differences in
chemical reactivity of cysteines engineered into this helix, and
studied fluorescence labels attached to these cysteines. Our results
clearly detect a conformational change in helix F, and the results are
consistent with a previously proposed movement of this helix upon
rhodopsin activation. Finally, the methods we describe here can be
broadly applied to studies of other GPCRs that may be available in
limited quantities.
Materials
All buffers were purchased from either Fisher or Sigma, except
the spectroscopy grade HEPES that was purchased from United States
Biochemical Corp. PyMPO-maleimide and mBBr were purchased from
Molecular Probes (Eugene, OR). DM was purchased from Anatrace (Maumee,
OH), and GBX red light filters from Eastman Kodak Corp. Polystyrene
columns (2-ml bed volume) were purchased from Pierce. The 9-mer peptide
corresponding to the rhodopsin C terminus was acquired from the Emory
University Microchemical Facility (Atlanta, GA). The 1D4 antibody was
purchased from the National Cell Culture Center (Minneapolis, MN).
11-cis-Retinal was a generous gift from Dr. R. Crouch
(Medical University of South Carolina and the National Eye Institute).
L-Cysteine was purchased from Sigma, and 4 × 4-mm and
3 × 3-mm fluorescence cuvettes were purchased from Uvonics (Plainview, NY). The 298-435-nm ( The definitions of the buffers used are as follows: PBSSC (0.137 M NaCl, 2.7 mM KCl, 1.5 mM
KH2PO4, 8 mM
Na2HPO4), Buffer A (1% DM, PBSSC (pH 7.2)),
Buffer B (0.1 M NaHCO3 (pH 7.6), 0.5 M NaCl, 0.02% NaN3), Buffer C (2 mM ATP, 0.1% DM, 1 M NaCl, 2 mM
MgCl2), Buffer D (0.05% DM, PBSSC (pH 7.0)), Buffer E
(0.05% DM, MES (pH 6.0)), and Buffer F (5 mM MES, 50 mM HEPES, 0.025% DM, 1 mM EDTA).
Methods
Construction and Expression of Rhodopsin Cysteine
Mutants--
The single cysteine rhodopsin mutants used in the present
study have been described previously (5). Briefly, the mutants were
constructed in the plasmid PMT4, containing a synthetic gene of
rhodopsin in which the potentially reactive "background" cysteine residues 140, 316, 322, and 323 were replaced with serines (10, 11)
(see Fig. 1). The mutant rhodopsin proteins were transiently expressed
in COS cells using the DEAE-dextran method (12, 13, 14). Two days after
transfection, the cells were harvested and stored at Purification of Rhodopsin Mutants--
All steps in this
procedure were in dark room conditions under filtered red light.
Purification of mutant rhodopsins followed previously described
procedures (8, 11) except that samples were purified using small
disposable polystyrene columns. Before purification, the columns were
prepared by adding 400 µl of the 1D4-coupled antibody bead slurry (a
50% bead v:v slurry). The columns were stored in Buffer B at 4 °C
and primed with 5 ml of Buffer A before use.
For a typical purification, the harvested cells from five plates of
transfected COS-1 cells were thawed in an ice bath for 30 min, then
solubilized in 5 ml of Buffer A + 0.5 mM
phenylmethylsulfonyl fluoride and placed on a nutator at 4 °C for
1 h. The samples were then centrifuged at 4,000 × g for 10 min, and the supernatant removed. This was followed
by a second centrifugation for 15 min at 20,000 × g at
4 °C. The supernatant from this second centrifugation was applied to
the 1D4 columns and allowed to flow through by gravity. The
pass-through was collected and reapplied to the column a total of five
times. The columns were next washed with 5 ml of Buffer C, then with 40 ml of Buffer D, followed by another 40 ml of Buffer E. During the last
5 ml of washing with Buffer E, a 27-gauge 0.5-inch needle was attached
to the column to slow the flow rate further. The samples were eluted in
200-µl fractions of Buffer E containing 250 µM 9-mer
peptide. A spectrum of each elution sample was measured using a
Shimadzu UV-1601 spectrophotometer, and the purified samples were then
stored at either 4 °C in the dark or at Reactivity Measurements of Rhodopsin Cysteine Mutants with
PyMPO--
The reactivity of the rhodopsin cysteine mutants in the
dark state was probed by reacting the samples (at pH 6.8 in 0.05% DM,
10 mM NaHCO3, 1 mM EDTA) with a
50-fold excess of PyMPO (Fig. 2A). The reaction was carried
out for 16 h at room temperature while the mutants were bound to
the 1D4 antibody columns. This approach allowed extensive washing of
the beads to remove any unreacted PyMPO label. After elution from the
column, PyMPO labeling was determined for each mutant by comparing its
absorption spectrum with an "ideal" rod outer segment (ROS)
rhodopsin spectrum. The ideal rhodopsin spectrum was then subtracted
from that of the labeled sample, allowing the amount of attached PyMPO
to be determined from the remaining absorbance at 380 nm (using an
extinction coefficient of 23,000 for PyMPO).
Reactivity Measurement of Purified Rhodopsin Cysteine Mutant
V250C with PyMPO in the MII State--
Three µg of mutant rhodopsin
in 70 µl was reacted with 20-fold excess PyMPO for 1 min at pH 7.2 (100 mM HEPES and 0.05% DM) and 4 °C in either the dark
( Labeling of Rhodopsin Mutants with Monobromobimane
(mBBr)--
Labeling of the cysteine mutants with mBBr (Fig.
2B) was carried out with the samples bound to the 1D4 beads,
similar to previous methods (10, 14). In brief, the 1D4 beads in Buffer
C were incubated with each solubilized mutant in 15-ml Falcon tubes at 4 °C on a nutator for 5 h. The suspension was then centrifuged at 4,000 × g for 2.5 min and the supernatant removed.
A series of washes were performed by adding the respective buffer to
the rhodopsin-bound 1D4 beads, followed by removal of the supernatant as described above. The washes were as follows: 10 ml of Buffer D, 10 ml of Buffer D (10-min incubation), 10 ml of Buffer F (10-min incubation). The 1D4-bound mutants were then incubated with a 20-fold
excess of mBBr in Buffer F at pH 6.7 for 18 h at 4 °C. Before
elution from the column, the reaction of the mBBr with samples was
quenched by the addition of a 20-fold excess of L-cysteine. The beads were then washed with 12 ml of Buffer F containing varying amounts of detergent (0.025%, 0.2%, 0.2%, and 0.025% DM) before continuing with the normal purification protocol (see above).
Characterization of mBBr-labeled Rhodopsin Mutants--
The mBBr
incorporation in each mutant was determined by comparison to a standard
curve of mBBr reacted with L-cysteine. Covalent attachment
of the labels was verified by subjecting the samples to
SDS-polyacrylamide gel electrophoresis analysis visualized by UV
irradiation. The effect of labeling on each mutant was assessed by
measuring its absorption spectra and rates of retinal release from MII.
Spectra were measured using a Shimadzu 1601 UV-visible spectrophotometer both in the dark state and in the MII state after
photoconverting the samples for 30 s with >500 nm light using a
150-watt fiber optic illuminator (Techni-Quip Corp). The MII stability
was assessed by measuring the time course of retinal release after MII
formation (14) using a Photon Technologies QM-1 steady-state
fluorescence spectrophotometer. Each measurement was carried out at
20 °C using 125 µl of a 0.25 µM mutant sample in 5 mM MES, pH 6.0 and 0.05% DM. After photoconverting the
samples to the MII state (see above), the retinal release measurements were carried out by exciting the sample for 3 s (excitation
wavelength = 295 nm, 1/4-nm bandpass slit setting) then blocking
the excitation beam for 42 s, to avoid photobleaching the samples.
This cycle was repeated for 90 min during each measurement. The
sensitivity of the measurements was enhanced by removing the emission
monochromator and monitoring the tryptophan fluorescence emission
through a combination of one >310-nm long pass filter and a
298-435-nm band pass filter. Results were analyzed, using Sigma Plot
(Jandel Scientific) to obtain the t1/2 values
for retinal release. All fits had r2 values
ranging from 0.99 to 1.1.
Fluorescence Excitation Scans of mBBr-labeled Mutants
K248C-R252C--
Excitation scans were carried out using the PTI QM-1
fluorescence spectrometer described above. The excitation bandpass was 1/3 nm, and the fluorescence emission was detected through two >470-nm
long pass filters (Oriel). Each measurement used 200 µl of a 2-µg
sample (250 nM). The samples were first scanned in the dark
state and then again after photoconverting to the MII state. Two such
scans from 300 to 450 nm (step size = 1 nm, integration time of
0.25 s) were averaged for each mutant, and two separate sample
preparations were measured. Under these conditions, less than 2%
bleaching of the samples was observed as judged by UV-visible spectroscopy. Excitation maxima were determined by importing the spectra into the program LabCalc (Galactic Industries), and then taking
the first derivative of the spectra, using a 21-point window. The peak
maximum was taken to be the crossover point, i.e. the wavelength where the first derivative crossed zero, going from positive
to negative.
Fluorescence Lifetime Measurement of mBBr-labeled Rhodopsin
Mutant V250C--
Fluorescence lifetime measurements of the
bimane-labeled rhodopsin mutants were carried out using a PTI
Laserstrobe fluorescence lifetime instrument. Measurements were taken
at 10 °C, using 381-nm excitation pulses (full width at half maximum
(FWHM) ~1.5 ns) while monitoring the emission through two long pass
filters (>450 nm and >470 nm). Measurements used 225 µl of a 1 µM sample placed in a 4-mm black jacketed cuvette, and
represented three averages of 5 shots per point, collected in 100 channels. Under these conditions, each measurement took less than 5 min
and resulted in less than 2% of the rhodopsin spectra being converted
to MII, as judged by UV-visible spectroscopy. Fluorescence lifetimes of
the samples in the MII state were measured as described above, except
the samples were irradiated with >490 nm light for 1 min before
measurement. The fluorescence decays were fit to a single exponential
using the commercial PTI program. Note that less than 6% of the sample had decayed to opsin and free retinal during the 5-min duration of the
measurement, as calculated from the retinal release rates at
10 °C.
Fluorescence Quenching Measurements--
Steady-state
fluorescence quenching measurements were carried out using the PTI
steady-state fluorescence spectrophotometer and KI as the fluorescence
quenching agent. Measurements used 125 µl of a 0.25 µM
sample in a 2-mm cuvette. Excitation was set at 380 nm using 1/3-nm
bandpass setting. The emission was monitored through two >470-nm long
pass filters. Five separate samples with different KI concentrations
(ranging from 0 to 25 mM) were measured. The salt
concentration in the sample was kept constant at 25 mM by
the addition of a corresponding amount of KCl, and
Na2S2O3 was added to 0.10 mM to inhibit formation of I3
The data were plotted as fluorescence intensity versus
concentration of quenching agent, to calculate the Stern-Volmer
quenching constant, KSV.
Expression, Purification, and Spectral Characterization of Single
Cysteine Rhodopsin Mutants K248C-I256C--
In the present paper, we
have used rhodopsin mutants containing unique, single cysteine residues
in the cytoplasmic end of helix F to investigate whether movement
occurs in this region of the protein upon MII formation. The mutants
(K248C-I256C, shown in Fig. 1) were
previously constructed (5) in the "background" mutant ( Accessibility of the Rhodopsin Cysteine Mutants to the
Cysteine-specific Reagent PyMPO-maleimide--
The accessibility of
the cysteine mutants K248C-I256C in dark state rhodopsin mutants was
investigated using the cysteine-specific reagent PyMPO-maleimide (Fig.
2A). As anticipated, this
large, rigid molecule demonstrated differences in its ability to
penetrate and react with cysteine residues in sterically constrained
regions of the protein (Fig. 3). The
spectra of each PyMPO-labeled mutant (purified after reaction with for
16 h with a 50-fold excess of PyMPO while bound to the 1D4 column)
is shown in Fig. 3A, and a plot of the labeling efficiency
is shown in Fig. 3B. While several mutants (K248C, E249C,
T251C, R252C, and I255C) showed a near one-to-one labeling efficiency
with the PyMPO, several others (V250C,M253C and V254C,I256C) showed
little to no reactivity. The reactivity of V250C with PyMPO in the MII
state was explored further (described below).
Measurement of Light-dependent Changes in Reactivity of
Mutant V250C with PyMPO--
Mutant V250C, which showed almost no
reactivity with PyMPO in the dark state, was next studied to see
whether it might show increased reactivity in the MII state (Fig.
4). Indeed, a dramatic increase in
labeling of this residue was observed after the sample was converted to
the MII state. In contrast, the control "background" mutant Labeling the Rhodopsin Mutants with Monobromobimane and
Characterizing the Samples--
The single cysteine mutants were next
labeled with the cysteine-specific label monobromobimane for subsequent
fluorescent spectroscopic studies. Monobromobimane was chosen because
it is a relatively small label (approximately the size of a spin label or tryptophan residue) with a long fluorescence lifetime and high quantum yield. Additionally, monobromobimane is sensitive to changes in
the polarity of the surrounding solvent (16).
The labeling efficiency for each mutant was determined by comparison to
a "standard" graph of bimane (Table
I). Mutants K248C through M252C could
each be reacted with approximately one bimane label per protein.
However, the labeling efficiencies of mutants I255C and I256C were much
less than one label per protein, and consistent with the PyMPO labeling
results, mutants M253C and V254C showed essentially no reactivity with
the bimane label. Thus, the latter four mutants were not used in
further analysis. Note that only ~0.2 mBBr/protein were observed to
react with the "background" cysteine-less mutant
The effect of the bimane label on each mutant was characterized by
measuring the labeled mutant's absorption spectra and rate of retinal
release from the MII state. None of the absorption spectra for any of
the mutants showed obvious differences with respect to wild-type
rhodopsin in either the dark state or MII state (Table I). Furthermore,
all of the bimane-labeled mutants displayed retinal release rates
similar to wild-type rhodopsin (Table I). These results suggest that no
dramatic perturbation of the protein structure was induced by the
bimane label.
Fluorescence Excitation Scans of Monobromobimane-labeled
Mutants--
Fluorescence excitation spectra were measured for the
mutants to assess the environment surrounding each bimane label (Fig. 5). The excitation and emission spectra
of bimane are sensitive to the polarity of the surrounding solvent
(16). In our own experiments,3 we have found
that the excitation maxima of bimane can range from 388 nm in water
(dielectric constant = 78.5) to 371 nm in dioxane (dielectric
constant ~ 2.2). We used this property of bimane to monitor
conformational changes by detecting shifts in the excitation maxima of
the attached bimane labels. This approach has a key advantage for
rhodopsin studies. During fluorescence excitation measurements, the
samples are subjected to a minimal amount of measuring beam light, due
to the small bandpass excitation slit settings used to obtain high
wavelength resolution data. UV-visible spectra taken before and after
the measurements showed less than 2% bleaching of the rhodopsin
samples had occurred (data not shown).
As can be seen in Table I, four of the mutants (K248C, E249C, T251C,
and R252C) showed small changes in their bimane excitation maxima upon
conversion to the MII form. (Note that the total fluorescence intensity
was approximately 2-fold higher for all of the mutants in the MII state
(not shown) because of a decrease in the fluorescence overlap and thus
energy transfer of bimane with retinal.) In contrast, the excitation
spectra of mutant V250C-mBBr showed a large blue shift in its
Fluorescence Quenching Studies of V250C-mBBr in the Dark State and
MII Form--
The conformational change detected at V250C-mBBr was
further investigated by fluorescence quenching studies. As can be seen from the Stern-Volmer plot of these studies (Fig.
6A), steady-state fluorescence
quenching by KI is much greater for the MII species. However, these
results must be corrected for the differences in fluorescence lifetimes
of the dark state and MII state V250C-mBBr, as shown in Fig.
6B. The most appropriate way to do this is to compare
bimolecular quenching constants (kq) rather than
the Stern-Volmer quenching constants, KSV,
obtained from the steady-state fluorescence measurements. This can be
accomplished by using the relationship kq = KSV/
The Stern-Volmer quenching constants (KSV)
obtained from the steady-state fluorescence quenching measurements, the
fluorescence lifetimes ( In this paper, we set out to test the proposal that a movement of
helix F in rhodopsin occurs upon MII formation (7-9). Our experiments
employed a series of eight mutants, each of which contained a single
reactive cysteine in a different part of helix F (Fig. 1). We used two
different methods to analyze these mutants. The reactivity of the
engineered cysteine residues was investigated (4, 17, 18) using a
large, rigid cysteine reactive reagent, PyMPO (Fig. 2A), and
the fluorescence properties of a solvent-sensitive fluorescent label
(mBBr, Fig. 2B) attached to the cysteine residues was
studied (15).
The first striking observation from these studies was that the cysteine
residues varied in their reactivity toward PyMPO. Furthermore, the
reactivity varied in a sequence-specific fashion (Fig. 3) that is
consistent with models showing this region of rhodopsin to be an helix. Interestingly, only upon
photoactivation to the MII form did residues found on the inner
"face" of this helix react with the PyMPO-maleimide. The ability of
the dark state mutants to react with the fluorescent label
monobromobimane followed a similar pattern. Furthermore, fluorescence
measurements indicate that a bimane label on the inner face of the
helix (at V250C) detects changes in the polarity of its environment and
accessibility to a fluorescence quenching agent upon MII formation.
Viewed together, the data provide further direct evidence that
rhodopsin activation involves a conformational change at helix F.
INTRODUCTION
Top
Abstract
Introduction
Procedures
Results
Discussion
References
max = 500 nm) and the retinal is converted to an all-trans form (2).
This activated rhodopsin species, called MII (
max = 380 nm), is the only form of rhodopsin that can readily bind and activate
the G-protein transducin and initiate the subsequent biochemical events
involved in vision.
EXPERIMENTAL PROCEDURES
Top
Abstract
Introduction
Procedures
Results
Discussion
References
max = 355 nm) band
pass filter and the >310-nm and >470-nm long pass filters were
purchased from Oriel (Stratford, CT). All materials for gel
electrophoresis were purchased from Hoefer (San Francisco, CA) and
Fischer Biotech (Pittsburgh, PA), and the 30% acrylamide/bisacrylamide
solution (37.5:1) was purchased from Bio-Rad.
80 °C.
80 °C for longer term storage.
) or MII (+) state (bleached for 30 s). The reaction was then
quenched by the addition of L-cysteine to a 1 mM final concentration. Samples were resolved by
SDS-polyacrylamide gel electrophoresis. The amount of label
incorporation was determined using a Bio-Rad Gel Documentation
Instrument by comparing the signal intensity with that of a series of
PyMPO-labeled rhodopsin standards of known concentration. The
background mutant,
(containing no reactive cysteine residues) (10,
11), was used to show specificity in PyMPO labeling. After
visualization of the gels, the samples were subjected to Coomassie
staining to check for equal distribution of protein in each of the
sample wells.
.
Measurements of the MII state were carried out after bleaching the
samples with >490 nm light for 30 s, as described previously.
Fo and F are the intensities
of the fluorescence before and after addition of the KI quenching agent
(Q), respectively. The plots were analyzed using Sigma Plot
(Jandel Scientific). The KSV values thus
obtained were then used with the measured fluorescence lifetimes
(
(Eq. 1)
0) to determine the bimolecular quenching constant, kq, from the following form of the Stern-Volmer
relationship (15).
(Eq. 2)
RESULTS
Top
Abstract
Introduction
Procedures
Results
Discussion
References
)
containing no other reactive cysteine residues (10). The mutants were
transiently expressed in COS-1 cells and purified using an
immunoaffinity procedure (12, 13). Similar to the previous study, we
found these mutants are all wild-type-like, as judged by their ability
to bind 11-cis-retinal and form a 500-nm absorbing species.
Further, each mutant could be obtained in the correctly folded form, as
judged by the
A280 nm/A500 nm ratio
(which ranged from 1.6 to 1.8).
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Fig. 1.
Suggested secondary structure model of
rhodopsin. The cysteine residues introduced into this protein (5)
are shown in black. The model also shows in gray
the native cysteine residues at positions 140, 316, 322, and 323 that
were changed to serines to produce a protein with no "background"
reactive cysteines. The cytoplasmic side is on the top in
this drawing. Labels A-G refer to the helices below.
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Fig. 2.
Structures of the cysteine reactive agents
used in this study. A, PyMPO-maleimide; B,
mBBr.
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Fig. 3.
Reactivity of PyMPO-maleimide with dark state
rhodopsin cysteine mutants. A, spectrum of each
PyMPO-labeled mutant. Each spectrum shows the labeled mutant, a
wild-type ideal rhodopsin spectrum, and the difference between the two,
from which the amount of PyMPO was calculated. B, plot of
the number of PyMPO labels/mutant determined from A. Details
are given under "Experimental Procedures."
showed no reactivity toward the PyMPO either in the dark state or after
photobleaching to the MII form. The extent of PyMPO labeling was
determined to be one PyMPO per mutant V250C by comparison to a series
of rhodopsin standards previously labeled with PyMPO (Fig. 4,
B and C). The specificity of the PyMPO for
residue V250C was established further by monitoring the time course of
the labeling reaction (Fig. 4D). The results from these
experiments show that one PyMPO specifically reacts with the cysteine
residue at V250C, and does so only upon MII formation. Note that,
during the 5-min time span used in this labeling experiment, less than
3% of the sample has decayed from MII to opsin (calculated from a
t1/2 for MII = 115 min at 4 °C; Ref.
14).
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Fig. 4.
Reactivity of the purified single cysteine
rhodopsin mutants free in solution with PyMPO-maleimide.
A, 3 µg (75 pmol) of mutants V250C and (no cytoplasmic
cysteines) were reacted at pH 7.2, at 4 °C with a 20-fold excess of
PyMPO-maleimide for 1 min either in the dark (
) or after bleaching to
the MII state (+). The reaction was quenched by the addition of
cysteine to a final concentration of 1 mM and the samples
were then analyzed by SDS-polyacrylamide gel electrophoresis. Gels were
visualized using UV irradiation and documented using a Bio-Rad gel
documentation instrument. The picture on the right shows the
Coomassie staining of the same gel. B, quantization of
PyMPO-maleimide incorporation into mutant V250C. Three µg (75 pmol)
of V250C in the dark and after photoconversion to the MII form were
reacted with PyMPO under the same conditions described above. A series
of PyMPO-labeled rhodopsin standards (a = 4.5 pmol,
b = 11 pmol, c = 23 pmol,
d = 34 pmol, e = 56 pmol,
f = 68 pmol) was included on this gel. C,
analysis of data in B showing that the PyMPO label
incorporation into mutant V250C is ~0.05 in the dark state, and
~0.8 mol of label/mol of rhodopsin in the MII state. D,
measurement of the extent and rate of incorporation of PyMPO into V250C
under the same conditions as in A, except that the reaction
was quenched at the various indicated time points. Data represent an
average of two separate experiments.
, consistent
with previous reports showing little reactivity of
with different
cysteine-specific reagents (10, 11).
Characterization of monobromobimane-labeled rhodopsin cysteine mutants
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Fig. 5.
Fluorescence excitation studies of
bimane-labeled rhodopsin cysteine mutants E248C-R252C in both the dark
state and MII state. A, normalized representative
fluorescence excitation spectra of two of the bimane-labeled mutants
(E249C, V250C) in the dark state (heavy line) and
MII state (light line). B, plot of the
excitation max obtained from the spectra of the
bimane-labeled mutants. The plot demonstrates the periodicity in the
data and indicates the large change in polarity sensed by the bimane
label at V250C. Samples were in 0.025% DM, pH 6.0, at 4 °C.
Excitation slits were equal to 0.25-nm bandpass, and the emission was
collected at >475 nm using long pass filters. Each measurement took
approximately 5 min, and resulted in less than 2% bleaching, as judged
by UV-visible spectra. Results are from an average of two measurements,
and the error bars represent the variance between
the two measurements.
max (approximately 5-6 nm), which corresponds to a
change in the apparent dielectric constant from ~44 in the dark state
to ~17 in the MII form. Thus, the label at this position becomes more
hydrophobic in the MII state, and clearly indicates that a
conformational change occurred in this region of the protein.
0. Note that the different
lifetimes are most likely due to the decreased fluorescence overlap
(and thus energy transfer) from bimane to retinal in the MII state.
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Fig. 6.
Fluorescence quenching studies of V250C-mBBr
in the dark state and MII form. A, Stern-Volmer plot of
fluorescence quenching of V250C-mBBr with KI. The quenching is much
greater for the MII species. B, fluorescence lifetime
measurements of V250C-mBBr in the dark and MII states, used to
calculate the bimolecular quenching constant kq.
The KSV results from A and the
fluorescence lifetimes from B are given in Table II. These
values were used to calculate kq, the
bimolecular quenching constant. The higher kq in
the MII state indicates more collisions of the bimane label occur with
the aqueous quenching agent I . Detailed discussion of
these measurements is given under "Experimental Procedures."
0), and the bimolecular
quenching constant (kq) calculated for
V250C-mBBr in the dark and MII states are given in Table
II. Note that the higher
kq obtained for V250-mBBr in the MII state clearly shows that the bimane label experiences more collisions with
the aqueous quenching agent I
than it does in the dark
state. These results again clearly show that some conformational change
has occurred in the protein at this location. Details of these
measurements are given under "Experimental Procedures."
Fluorescence quenching studies of V250C-mBBr
DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References
helix (7, 8, 19, 20). Fig. 7A
indicates on a rhodopsin model (7) the location of the mutants'
reactivity with PyMPO.
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Fig. 7.
Models of the region studied by cysteine
reactivity and fluorescence spectroscopy. A, model
showing the location of cysteine residues on helix F, based on models
previously presented (4, 7). The residues that labeled well with
PyMPO-maleimide in the dark state are shown as dark
circles. Mutant I256C, which showed some labeling, is
indicated as a gray circle. The three cysteine
residues that showed no reactivity toward PyMPO in the dark state are
shown as white circles. Note that mutant V250C,
which showed changes upon MII formation as detected by chemical
reactivity and fluorescence spectroscopy, is located in the middle face
of this helix. B, model showing helical movement during
conversion of dark state rhodopsin to the MII form, based on a model
previously proposed from EPR studies (8). Such a movement would be
consistent with the present data, as it would make mutant V250C (shown)
more accessible to labeling with PyMPO-maleimide, and increase the
accessibility of an attached fluorescent label to quenching agents.
However, at this stage the data cannot yet distinguish between a helix
rotation or tilt.
The results also provide direct evidence for a conformational change around helix F during MII formation. For example, residues on the inner face of this helix could only react with the PyMPO in the MII form (V250C (Fig. 4) and also M253C (data not shown)). Similarly, a fluorescent bimane label at V250C also detected changes both in the polarity of its environment (Fig. 5) and in its accessibility to fluorescence quenching agents (Fig. 6) upon formation of the MII state. While our results are consistent with the previously proposed outward tilt and/or rotation of helix F (8) (shown in schematic form in Fig. 7B), they do not resolve the exact nature of the movement or rule out a possible concomitant movement of helix C (6, 21). One thing clear from these studies is that the packing of the inner face of helix F is quite tight. Neither the PyMPO nor the mBBr could react with some residues on the inner face of this helix, even after extended incubation times (16 h). In contrast, the reaction of PyMPO with V250C in the MII state was complete in less than 30 s (Fig. 4D).
The hydrophobic shift detected by the bimane label at V250C is noteworthy and has not been reported before. This shift, going from an apparent dielectric constant of ~44 to ~17, is most likely is due to a restructuring of the protein that introduces the bimane label into new contacts with neighboring hydrophobic residues. Examination of recent rhodopsin models (19-22) shows several "hydrophobic patches" in the helices neighboring V250C. The role that a new, more hydrophobic patch may play in transducin activation is unknown. However, it is tempting to speculate that it may be involved in increasing Arg-135 exposure during MII formation. Arg-135 is part of a highly conserved pair of residues required for normal transducin binding and activation (23-26). The new region of hydrophobicity might change the hydrogen bonding propensity and/or location of Arg-135 in the MII state.
Several other indirect clues from biochemical and mutagenesis studies have suggested a movement of helix F is required to form an active MII rhodopsin species. For example, linking helix F with helix C, through either disulfide linkages (8) or metal chelating agents (9) abolishes the ability of the mutants to activate transducin. Substitutions of M257L on the inner face of this helix have also been found to result in constitutively active mutants, presumably through a steric mechanism (27).
A growing body of evidence from mutagenesis and modeling studies
suggests that activation of other GPCRs involves the same helix
(28-32). Interestingly, studies of the m2 muscarinic receptors suggest
the region studied in the present paper (which includes the so-called
VTIL region; Ref. 32), might become more exposed upon receptor
activation (33), and thus allow direct coupling with the C-terminal
sequence of i/o (34). The movement we describe here may
be involved in a similar way, effectively exposing this region in
rhodopsin and enabling direct coupling with the G-protein transducin.
Alternatively, a helix F/helix VI movement may be required to reorient
the C-terminal tail (35). Recently, a movement of the same region in
helix VI in the
-adrenergic receptor was suggested from fluorescence
spectroscopy studies (36). This latter result raises the intriguing
possibility that a movement of helix F/helix VI is a conserved and
primary step in the activation of GPCRs.
In future work we intend to define further the nature of the
conformational change we detect here in helix F, and to attempt to
resolve whether the helix movement involves exclusively a rotation or a
tilt, or some combination of the two movements. It would also be of
interest to test the functional role of the movement by studying
defective rhodopsin mutants. Finally, we point out that the approaches
described in this paper allow experiments requiring only minimal
amounts of sample (~2-3 µg), and thus should work with other GPCR
systems of limited availability. The combination of PyMPO as a probe
with SDS-polyacrylamide gel electrophoresis analysis is especially
advantageous, as it provides a simple, sensitive, and direct method to
measure conformational changes in GPCRs.
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ACKNOWLEDGEMENTS |
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We thank Dr. C. Gatto for discussions and S. E. Mansoor for the fluorescence lifetime measurements.
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FOOTNOTES |
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* This work was supported in part by Grant R01 EY12095-01 from the National Institute of Health, Bethesda, MD, and a grant from the Medical Research Foundation of Oregon.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: Dept. of Biochemistry
and Molecular Biology, Oregon Health Sciences University, 3181 S.W. Sam
Jackson Park Dr., Portland, OR 97201-3098. Tel.: 503-494-0583; Fax:
503-494-8393.
The abbreviations used are: GPCR, G-protein-coupled receptor; MES, 2-(4-morpholino)ethanesulfonic acid; DM, dodecyl maltoside; PyMPO, N-(maleimidylethyl)-5-((4-methoxyphenyl)oxazol-2-yl)pyridi-nium methanesulfonate; MII, metarhodopsin II; mBBr, monobromobimane; SDSL, site-directed spin labeling.
2 Note that results from SDSL studies of helices A and B have not yet been reported, although such studies are currently under way (W. L. Hubbell and H. G. Khorana, personal communication).
3 S. E. Mansoor, H. S. Mchaourab, and D. L. Farrens, manuscript in preparation.
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REFERENCES |
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