Correspondence to: Jack M. Sullivan, Assistant Professor of Ophthalmology and Biochemistry, SUNY Health Science Center, Dept. of Biochemistry Weiskotten Hall, 4255, 750 East Adams St., Syracuse, NY 13210. Fax: 315-464-8750; E-mail:sullivaj{at}vax.cs.hscsyr.edu.
Released online: 11 October 1999
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The early receptor current (ERC) represents molecular charge movement during rhodopsin conformational dynamics. To determine whether this time-resolved assay can probe various aspects of structurefunction relationships in rhodopsin, we first measured properties of expressed normal human rhodopsin with ERC recordings. These studies were conducted in single fused giant cells containing on the order of a picogram of regenerated pigment. The action spectrum of the ERC of normal human opsin regenerated with 11-cis-retinal was fit by the human rhodopsin absorbance spectrum. Successive flashes extinguished ERC signals consistent with bleaching of a rhodopsin photopigment with a normal range of photosensitivity. ERC signals followed the univariance principle since millisecond-order relaxation kinetics were independent of the wavelength of the flash stimulus. After signal extinction, dark adaptation without added 11-cis-retinal resulted in spontaneous pigment regeneration from an intracellular store of chromophore remaining from earlier loading. After the ERC was extinguished, 350-nm flashes overlapping metarhodopsin-II absorption promoted immediate recovery of ERC charge motions identified by subsequent 500-nm flashes. Small inverted R2 signals were seen in response to some 350-nm flashes. These results indicate that the ERC can be photoregenerated from the metarhodopsin-II state. Regeneration with 9-cis-retinal permits recording of ERC signals consistent with flash activation of isorhodopsin. We initiated structurefunction studies by measuring ERC signals in cells expressing the D83N and E134Q mutant human rhodopsin pigments. D83N ERCs were simplified in comparison with normal rhodopsin, while E134Q ERCs had only the early phase of charge motion. This study demonstrates that properties of normal rhodopsin can be accurately measured with the ERC assay and that a structurefunction investigation of rapid activation processes in analogue and mutant visual pigments is feasible in a live unicellular environment.
Key Words: photoreceptor, gating currents, conformational activation, phototransduction
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Rhodopsin is the visual pigment of the rod photoreceptor and catalyzes the activation of the G-protein, transducin. Seven transmembrane segments of opsin form a pocket to bind 11-cis-retinal (11cRet),1 forming a chromophore with the lysine (K296) as a protonated Schiff base (PSB+-H). The chromophore isomerizes to all-trans-retinal within 200 fs (max 380 nm). The transition from the Meta-I to the Meta-II state is the only endothermic state change that occurs during the thermal dark reactions. This indicates that the spontaneous transition into these states is associated with a large positive entropy. A significant molecular volume increase occurs during the lifetime of the Meta-II states (
helices, and the configuration of the cytoplasmic loops that are temporally correlated with formation of the R* state, which allows transducin docking (
Other tools such as Fourier transform infrared spectroscopy or electron spin resonance can sample conformation changes outside the chromophore environment. But, like time-resolved absorption studies, these are currently limited by the need for hundreds of micrograms or milligrams of detergent-extracted and purified rhodopsin, or cysteine mutagenic engineering to allow site-specific attachment of spin probes. Compared with Fourier transform infrared spectroscopy, only the more sensitive electron spin resonance technology can resolve environmental transitions on a millisecond time scale. These tools have nonetheless contributed greatly to our current understanding of the rhodopsin activation process (
The early receptor potential (ERP) is a charge redistribution in rhodopsin associated with protein conformational changes (
The early receptor current (ERC) of rhodopsin activation is the direct measure of charge flow that underlies the ERP. This nonlinear capacitative current shows saturation, dependent upon the amount of rhodopsin molecules available for activation (
The ERC of rhodopsin activation in intact photoreceptors has been elegantly studied using gigaohm-seal, whole-cell patch clamping techniques (
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cell Culture and Fusion
Human opsin-expressing HEK293S cell lines were used for ERC recordings (
Cellular Pigment Regeneration
Cells on coverslips were washed in regeneration buffer and placed in a light-tight container in the darkroom at room temperature (2225°C). Regeneration buffer was (mM): 140 NaCl, 5.4 KCl, 1.8 CaCl2, 1.0 MgCl2, 10 glucose, 10 HEPES-NaOH, pH 7.2, and 2% (wt/vol) (~290 µM) fatty acidfree bovine serum albumin (FAF-BSA; Sigma Chemical Co.). Concentrated (mM) 11cRet or 9cRet stocks (in ethanol) were added in small volumes to this solution to a final concentration of 50 µM in preliminary experiments and 25 µM in the final protocol with 0.025% (vol/vol) -D-tocopherol (vitamin E) added as an antioxidant. Vitamin E is found in high concentrations in photoreceptor cells and may serve as an antioxidant (
Flash Photolysis
Rhodopsin regenerated in fused giant cells was activated by an intense flash microbeam apparatus described in detail elsewhere ( 374 nm) such that isomerization (cis
trans or trans
cis) of any free chromophore is not expected during flashes used to elicit ERCs. Unless otherwise mentioned, flashes were delivered at the maximum capacity of the instrument. Intensities were 108109 photons/µm2 across the near UV/visible band. Flash microbeam intensities were measured using a calibrated photodiode placed over the specimen plane of the microscope. To regulate flash intensity output (see Figure 5), the voltage on the flash tube energy storage capacitor was adjusted. Flash duration was only ~14 µs, insuring that the Meta-I
Meta-II equilibrium (milliseconds) generated at room temperature in these experiments was not perturbed by photoregeneration to other states (
Meta-II transition. Shielding and fiber optic transmission prevent contamination of the patch-clamp electronics with flash-associated noise.
|
|
|
|
|
Photosensitivity (Pt) is the product of quantal efficiency () and the wavelength-dependent absorbance cross section (
). The absorbance cross section of wild-type human rhodopsin is 1.53 x 10-8 µm2 (calculated from an extinction coefficient of 40,000 M-1 cm-1 at 493 nm (
is 0.67, leading to a Pt of 10-8 µm2 for normal human rhodopsin at peak extinction (493 nm). Pt can be used to estimate the fraction of rhodopsin molecules absorbing at least one photon per flash using the zero-order term of the Poisson equation [1 - Po = 1 - exp(-Pt · i)], where i is the flash intensity (photons/µm2) and Po is the fraction that absorbs no photons [Poisson Eq.: Pn = (Pt*i)n*exp(-Pt*i)/n!, where n is the number of absorptions per chromophore]. In this calculation, one adjusts
by the ratio of absorbance at the wavelength of interest to that at peak extinction.
is assumed to be constant and independent of wavelength. For the 70-nm bandpass filters used in these experiments (centered at 350, 430, 500, and 570 nm), the fraction of molecules absorbing at least one photon were estimated to be 0.159, 0.716, 0.963, and 0.273, respectively. For the 30- and 10-nm bandpass filters used in these experiments (centered at 400, 440, 480, 500, 520, 540, 580, and 620 nm), the fraction of rhodopsin molecules absorbing at least one photon were estimated at 0.226, 0.626, 0.831, 0.80, 0.733, 0.44, 0.122, and 0.013, respectively. These calculations assume no orientational factors, no self-screening effects, and transparent cellular media. Thus, microbeam flash intensities were not expected to be experimentally limiting for flash photolytic stimulation of expressed rhodopsin pigments, except perhaps for the 620-nm stimulus. The maximum extent of rhodopsin bleaching (i.e., formation of Meta-II) after a single flash is 50% (
Whole-Cell ERC Recording
Cells on coverslips were imaged using infrared light (high pass cutoff 830 nm) at 80160x by an inverted microscope (Diaphot; Nikon Inc.) equipped with Nomarski differential interference contrast, a CCD camera, and a TV monitor. The microscope was housed in a Faraday cage in a dark room. Microelectrodes were fashioned from borosilicate glass using two stage pulls and coated with Black Sylgard (Dow Corning Corp.). Electrodes were routinely filled with one of two intracellular solutions (with or without 10 mM HEPES-CsOH) containing (mM) 70 tetramethylammonium (TMA)-OH, 70 Mes-H, 70 TMA-F, 10 EGTA-CsOH, 10 HEPES-CsOH, pH 6.5; these solutions yielded ERCs with no qualitative changes and are called I-1. The internal pH was chosen to be 6.5 to forward bias the Meta-I Meta-II equilibrium strongly in favor of Meta-II (
The patch-clamp instrument was an Axopatch 1C with a CV-4 resistive feedback headstage and the later was used with a gain of 1 (0.5 Gigaohm feedback resistor; Axon Instruments). Since the ERC is a capacitative current, whole-cell capacitance (Cmem) and series resistance were not compensated, because this has the potential to alter the waveform. Membrane holding potential was clamped at 0 mV unless otherwise noted in the legends. Whole-cell capacity current was measured by a +20 mV/4 ms test pulse from a holding potential of -80 mV and Cmem was computed by integrating the capacitative current waveform to obtain charge (Q) (Q/V = Cmem) after substraction of ohmic current. Cell surface area was computed from the measured Cmem (1 µF/cm2). Ramp voltage clamps were delivered to test for a high resistance membrane and low level leakage. Cells with large leakage were discarded. Whole-cell currents were recorded at 5 kHz bandwidth using an eight-pole Bessel filter. Flashes were controlled and ERC and flash stimulation data acquired using pCLAMP 5.51 (CLAMPEX) and digitized (200 µs/point) by a Labmaster (100 kHz) interface board (Scientific Solutions Inc.). This acquisition rate was selected to provide the best possible representation of the R1 signal, which is still undersampled, while critically allowing the full time course of the R2 component to be acquired out to 100 ms. All ERC data was processed and analyzed using the Origin4.1 package (MicroCal Software, Inc.). Nonlinear least squares fitting was conducted using a Levinberg-Marquardt algorithm.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Giant Cells Generate Large ERCs that Recover Spontaneously with Dark Adaptation
Polyethylene-glycolfused giant HEK293S cells are used to amplify the amount of regenerable plasma membrane opsin (WT or mutant) in single large cells to improve the signal-to-noise ratio (SNR) during ERC data acquisition. These are prepared from single cells that are stably transformed to constitutively express WT or mutant human opsins in the range of 110 x 106 molecules (
Figure 1 A shows a giant cell ERC obtained on the first flash series (500 nm) after a 30-min regeneration. Flash stimuli were given at 500 nm to extinguish the ERC signal into background whole-cell noise. Both the submillisecond negative R1 current and the millisecond-order larger, and positive R2 current were routinely observed and essentially identical to those recorded in photoreceptors (see
Figure 1 D shows the kinetics of the spontaneous recovery of total ERC charge upon dark adaptation for two similarly sized fused cells (67.5 and 65.6 µm diameter). Cells were bleached and dark adaptation was allowed to occur for variable time periods before additional 500-nm flashes were given in rapid succession to extinguish the ERC signal. The total charge (Q) present at each time point of dark adaptation was obtained by summing all Qi values to yield Q
at that time point. Q
values were normalized to the maximum charge determined for each cell so that regeneration data could be compared. The data were fit by a single exponential accumulation curve. The initial regeneration rate is fast and the process begins to approach steady state by 10 min of dark adaptation. The half time of ERC regeneration was 3.2 ± 1.1 min. If primary regeneration in 2% BSA fully equilibrates WT-HEK293S cells with 11cRet at a concentration (25 µM) in considerable excess over the expected molarity of opsin in fused giant cells (
2.4 µM), then spontaneous regeneration can be described as a process with pseudofirst order kinetics from a single compartment containing chromophore. We routinely dark adapted cells for either 10 or 15 min between successive extinctions to allow visual pigment and ERC signal recovery to the stationary plateau or, at a minimum, to permit a criterion level of regeneration for any comparison of charge motions under different conditions tested on a single cell.
The Source of Chromophore during Dark Adaptation Is Intracellular
The source of chromophore that promoted spontaneous ERC recovery was investigated. Single fused giant cells regenerated with 11cRet were subjected to primary extinction by successive flashes at 500 nm in normal bath solution (E-1 without chromophore). Dark adaptation promoted spontaneous pigment regeneration and allowed for a secondary extinction of the ERC. Immediately after the second extinction and throughout the next dark adaptation, the chamber volume was replaced with E-1 containing 10 mM hydroxylamine (NH2OH, pH 7.0) and an additional 5 min of dark adaptation preceded the next series of successive 500-nm flashes. ERCs were elicited and had Qi values comparable with those seen during the primary and secondary extinctions without NH2OH. Qi extinctions before and after NH2OH are shown (Figure 2 A). When NH2OH was present in the bath many more flashes (in this cell 23 flashes) were required to completely extinguish the ERC R2 charge in comparison to three to five flashes during the primary and secondary extinctions. This suggested that 10 mM NH2OH decreased photosensitivity and this effect will be examined fully in a subsequent study. Two dark adaptations (10 min each) in 10 mM NH2OH were followed by some but not full ERC signal recovery, which was then subsequently extinguished by additional flashes. ERC signals continued to be recordable on the time scale of tens of minutes in the presence of constant 10 mM extracellular NH2OH, a concentration far greater than the initial loading of chromophore (25 µM). Figure 2 B shows the exhaustion course of Qi vs. flash number for another giant cell before and after introduction of 10 mM NH2OH into the bath solution. In NH2OH total R2 charge decreased with successive extinctions and dark regenerations until no significant signals remained. Bath solution was then replaced with fresh E-1 containing 25 µM 11cRet in 2% FAF-BSA (without NH2OH). Strong ERC R2 signals were regenerated comparable in total charge with that seen during the primary extinction. These experiments demonstrate strong evidence that the source of 11cRet during spontaneous dark regeneration is internal to the cell that is recorded.
Additional observations support an intracellular origin for 11cRet during dark adaptation. First, the cell repeats dark-adaptive ERC regeneration until the signals expire, and then they do not again regenerate unless 11cRet is reapplied to the cell. This rundown is evident in the extinctions in NH2OH in Figure 2 and suggests that cells exhaust their store of 11cRet. Second, in some cells the plasma membrane appeared wrinkled or the cytoplasm appeared optically smooth after expiration of ERC signals. This suggested a change in the structure of internal membranes associated with regeneration ability. Finally, the greatest number of "visual cycles" in this study was eight and was encountered in the largest fused giant cells (80 µm diameter) and, although not yet systematically investigated, the number of regenerations appeared to scale in proportion to cell size. Given the hydrophobic nature of 11cRet, it is likely that it is stored by partitioning into internal cellular membranes, and then repartitions back to the plasma membrane during dark adaptation to regenerate visual pigment (see DISCUSSION).
The ERC Action Spectrum Is Consistent with the WT Human Rhodopsin Photopigment
A previous study of ERC spectral sensitivity used 70-nm bandpass filters and demonstrated a broad action spectrum consistent with the ground state of human rhodopsin pigment (
Instead of fitting spectral sensitivity data with a Lorenztian/Gaussian peak function (e.g., Voigt) ( band) and overlaid with the ERC action spectrum. WT rhodopsin was immunoaffinity purified from the same WT-expressing cell line used in these ERC experiments (
band peak at 493 nm) provides an excellent fit to the ERC action spectrum obtained from cells expressing WT human opsin that was regenerated with 11cRet. The ERC action spectrum peaked around 493 nm, consistent with the major absorption band of ground state human rhodopsin regenerated with 11cRet. Moreover, the bandwidth of the
peak of the human rhodopsin absorbance spectrum also fits the ERC data well. It is important to mention that all of the action spectra data were obtained from cells after they had undergone a primary bleach; that is, under conditions where long-lived bleaching intermediates or photoregenerated pigments might have been present. Normal human rhodopsin has an absorbance spectrum that is slightly blue shifted (
493 nm) with respect to the bovine pigment (498 nm) (
ERC Extinction Measures Bleaching of Rod Rhodopsin
Successive flashes at a single wavelength and intensity promoted progressive loss of ERC R2 charge until no further signal was obtained above background current noise. Since the interstimulus intervals between successive flashes were only ~10 s, pigment regeneration was minimal between stimuli, and regeneration should not contribute to the extinction progression. The spectral sensitivity of the ERC governs not only the efficacy of successive bleaches, but also single bleaches. Figure 4 A shows the Qi extinction of R2 in response to successive flashes at 570 and then 500 nm in a single giant cell. Cumulative flash intensity delivered is used as the dependent variable. After ERC charge was effectively extinguished into noise by 570-nm flashes, 500-nm flashes of greater effective intensity were immediately delivered. The additional extinguishable ERC charge found with 500-nm flashes indicated residual ground state rhodopsin in the cell after the 570-nm flashes. This resulted because 570-nm flashes are not as effective at eliciting ERC currents as are flashes at 500 nm given the relative absorbance of WT human rhodopsin at 570 vs. 493 nm (peak absorbance) (OD570/OD493 = 0.436) (570/
493 = 0.115), assuming equal photon density at the two wavelengths. At the maximal flash strengths used, the fraction of rhodopsin molecules absorbing at least one photon at 500 vs. 570 nm was estimated to be ~0.96 and 0.27, respectively. Thus, the flash system does not deliver sufficient photons at 570 nm to compensate for the lower probability of activation. This illustrates that detection of rhodopsin charge motions depends on the unitary charge motion, which should be the same at any wavelength (see univariance below), and the number of activated rhodopsin molecules that mobilize charge and sum into an ensemble ERC current. Even at peak wavelength (
500 nm), a given flash intensity may not be sufficient to generate ERC currents above noise because the ensemble ERC current lies within the noise band.
Single exponential exhaustion curves are fit for both data sets at the two respective wavelengths. ERC extinction data for 570-, 500-, or 430-nm flashes from many experiments were always fit by single exponential curves. This is consistent with the ERC charge motion being proportional to the amount of rhodopsin that remains unactivated before each flash is given. Extinction data were fit by the following model using a nonlinear least squares method:
![]() |
(1) |
where Qi is the charge motion resulting from a single flash, Q is the total charge before any flashes are given, I is the cumulative flash intensity, and Pt is the photosensitivity. The bleaching process follows an exponential extinction that was further tested by a natural logarithmic transform of Qi values and linear fitting (Figure 4 B). Equation 1 was used to determine Pt for several giant cells subjected to flash photolytic exhaustion under conditions of different flash stimulation wavelength. The charge extinction data for the initial (primary) and three subsequent (secondary) bleaches at 500 nm and single bleaches at 430 and 570 nm in a single large giant cell (
80 µm) are shown in Figure 4 C. The charge extinction data sets for each bleach were normalized to the maximum charge on the first flash, and then Equation 1 was fit to each data set and overlaid. The fit by the single exponential model implies that each flash promoted activation and extinction (bleaching) of a fraction of remaining ground state pigment. In some flash stimulation series, there was a residual content of charge that was slow to extinguish (1020%). Pt for the primary bleach at 500 nm was estimated to be ~3.3 x 10-9 µm2, but this is less reliable as there were only three points to fit. The Pt values obtained from fitted curves at 500 nm for the three secondary bleaches were 3.28 x 10-9, 3.04 x 10-9, and 2.24 x 10-9 µm2. Pt was stable over successive secondary bleaches and similar to that of the primary bleach. This is evidence in addition to the action spectrum that the ground state of rhodopsin is regenerated with 11cRet during dark adaptation between successive flash cycles and there was no accumulation of other intermediates with significantly different photosensitivities. The mean (±SEM) for Pt values from several cells are shown in Figure 4 D at 500(70), 430(70), 570(70), and 500(30) nm, where the filter bandwidth is indicated parenthetically. Both parametric and nonparametric (Kruskal-Wallis) analysis of variance tests were used to evaluate whether the means of the five conditions were different. No statistically significant difference was found between Pt estimates of human rhodopsin during the initial and secondary extinctions after spontaneous regeneration. However, the trend toward a lower Pt at 430 nm is consistent with the ratio of absorbance of rhodopsin at 430 vs. 500 nm (
430/
493 = 0.423) (
The Pt of WT human rhodopsin is estimated to be ~1.0 x 10-8 µm2 (see MATERIALS AND METHODS). The mean value of Pt [500(70) nm] determined from 11 extinctions in 6 cells, all after spontaneous regeneration was 2.6 ± 0.4 x 10-9 µm2 and the maximum value measured was 5.0 x 10-9µm2. Pt values measured using the extinction of R2 charge are consistent with but lower than that expected of a rhodopsin chromophore. This is likely due to the suppressive effect on Pt of photoregeneration resulting from multiple photon absorptions per rhodopsin molecule at the flash intensities used. For example, with each 500(70)-nm flash, the estimates on even numbered absorptions are ~50%, which would underestimate Pt by a similar amount (see DISCUSSION). In recent experiments, stimulus intensity (at 500 nm) was reduced (by 85%) to decrease the probability of multiple hits per molecule, allowing Pt estimation to a mean value of 8.5 x 10-9 µm2, which approximates that expected from the extinction coefficient (Brueggemann and Sullivan, manuscript in preparation).
Tests of Linearity and the Univariance Principle with ERC Measurements
As first demonstrated by = 0.67) of successful activation and will contribute to the kinetics of ERC charge flow. At fixed wavelength, variation in stimulus intensity is expected to affect the probability of absorption if each activated rhodopsin molecule makes an independent and additive (linear) contribution to the ERC, and photoreversal to other states by second photon absorptions is minimal. Figure 5 shows ERCs resulting from 500-nm stimulation at two different flash strengths. When these responses are normalized and overlaid for comparison, the kinetics of the R2 relaxation at different intensities are not distinguishable. The ERC response versus flash intensity was measured. The Qi response of the first flash in an extinction series at a constant intensity is plotted versus intensity. A linear fit of the charge motion versus absolute intensity was found. This indicated that the flash intensities used were below saturation for the cellular expression system. This result is consistent with known properties of the ERC/ERP. Each activated rhodopsin molecule undergoes conformational changes to contribute a quantum of charge motion to the overall R2 signal (
According to the univariance principle, the energy of the photon (wavelength) should not affect the activation kinetics of independent rhodopsin molecules. Photon energy only affects the probability of absorption because the molecular cross section is a function of wavelength. Figure 6 shows ERCs acquired from a giant cell at 430, 500, and 570 nm. ERC waveforms were normalized to the smoothed peak of the R2 current for comparison. A double exponential curve was generated to fit the relaxation kinetics of the R2 signal from the 500-nm stimulus. This template was then overlaid with the R2 signals from the 430- and 570-nm responses. The 500-nm template fits the R2 relaxations of the 430- and 570-nm responses rather well, even though the SNR was lower with 570-nm stimuli because the absolute response was smaller. Although the amplitudes of the ERCs and total charge motion of the R2 signal vary with wavelength, the kinetics of R2 relaxation are similar. Similarly, the large 500-nm R2 response in Figure 3 was fit with a double exponential function, and this template was overlaid with the large 440-, 480-, and 520-nm ERC responses and provided a good fit. The 400- and 540-nm ERC responses were of lower SNR and were not fit well by the template. ERC data in Figure 3 was collected with 30 nm FWHM stimuli. In the rhodopsin expression system, photon energy does not affect the kinetics of the state transitions in rhodopsin, which is consistent with the univariance principle.
|
Ground State Rhodopsin ERC Can Be Photoregenerated from Metarhodopsin-II
Previous studies have shown that the ground state of rhodopsin can be photoregenerated from Meta-II380 by near UV flashes delivered concurrent with its lifetime (
|
In most cells receiving UV stimuli, no apparent ERC charge motion occurred above the noise level of the cell. However, in a few large giant cells, apparent UV flashinduced ERC signals were identified even without any signal smoothing to suppress noise. An example is shown in Figure 7 C, where a response to a UV (350-nm) flash is generated immediately after extinction of ERC signals with 500-nm stimuli. The UV-induced ERC signal is small and has a negative (inverted) R2-like response. An unconstrained third order polynomial fit the UV R2 signal and also demonstrated the inverted R2 signals. An ERC generated with a 500-nm flash after dark adaptation in the same cell is shown to demonstrate the magnitude of the normal R2 charge motion. The time to peak of the positive 500- and negative 350-nminduced R2 signals was 5.9 and 12.2 ms, respectively. The ratio of the inverted to noninverted R2 charge was 0.175 (0.099 in another smaller cell).
9-cis-Retinal Regeneration Results in ERC Signals Consistent with Isorhodopsin
Cells were regenerated with 9cRet to test the feasibility of ERC investigation of rhodopsin activation in analogue visual pigments. Analogue visual pigments are usually formed from WT opsin and a synthetic retinal known to have unique properties (e.g., to block Meta-II formation), but could also be formed from synthetic retinals and site-specific opsin mutants. The naturally occurring 9cRet analogue forms isorhodopsin, a stable ground state pigment that is generated in a photostationary state with rhodopsin and bathorhodopsin () from the extinction coefficient for isorhodopsin at 483 nm (44,000 M-1 · cm-1) (
= 3.82 x 10-21 *
) (
= 1.68 x 10-8 µm2) and multiplying
by the quantal efficiency of isorhodopsin of 0.33 (
|
Kinetic Comparison of Mutant and WT Visual Pigments with the ERC
One way to exploit the sensitivity of the ERC technique is to investigate charge motions in mutant rhodopsins. We generated stable, high-level producing (~106 opsins/single cell), HEK293S cell lines of several human rod opsin mutants altered at single amino acids that could support proton exchange processes in the membrane region (
Figure 9 shows ERC signals in response to the first 500-nm flash after primary regeneration and secondary recovery in fused giant cells containing WT, D83N, or E134Q human rhodopsins. The WT pigment generated strong R1 signals during the primary bleach. R1 signals are rarely seen during the secondary or subsequent extinctions indicating that, if present, the size is below the limits of detection at the flash intensities used. Large (>40 pA) WT ERC signals typically require two exponentials to fit the time course of the R2 relaxation over the first 100 ms. Residuals are shown beneath the ERC waveforms. R1 signals were not observed in D83N rhodopsin during primary extinction in cells with R2 charges of the same order as seen in fused WT cells that had R1 signals. Like WT, the D83N R2 relaxation typically requires two exponentials to reliably fit its relaxation. However, D83N signals appear to lack the "stretched" exponential appearance seen in many large WT signals during the 100 ms after the flash. The E134Q ERC signal was distinctly different from the WT signal. R1 signals were not observed during primary extinctions. Moreover, the outward R2 signals in E134Q rhodopsin-expressing cells were markedly simplified in comparison with WT or D83N ERCs. The relaxation was brief and required only a single exponential to fit its decay. Like WT ERCs, D83N and E134Q ERC signals extinguished with successive flashes and had spectral sensitivity consistent with pigments absorbing ~500 nm (
|
To begin to characterize R2 relaxation, single or double exponential functions were fit to a large number of WT, D83N, and E134Q R2 signals from many cells of similar size range. Time constants associated with R2 relaxation, but not R1, are essentially independent of Cmem (a,
b). Since the identification of the respective components is dependent upon their weighting, it is possible that the
b constant could be assigned to the
a data set if the weighting of the
a component is small or unreliable (e.g., lower SNR). Therefore, all the time constants obtained for R2 relaxation were placed into a total ensemble, and histograms were generated from these populations for the WT, D83N, and E134Q datasets (Figure 10, AC). The WT pigment demonstrated a broad skewed distribution suggestive of density around three time constant ranges, whereas the E134Q distribution was simple and symmetrical and the D83N distribution was intermediate. To quantitatively characterize the ensemble of time constants, Gaussian distribution functions were fit to each histogram. The WT histogram was reliably fit by a sum of three gaussian distributions, centered at 4.1, 12.5, and 26.4 ms (Table 1). There were also residuals with time constants longer than the three fitted distributions. This analysis demonstrates the kinetic complexity of the WT R2 relaxation and supported the conclusion of a minimum of three charge states with distinct lifetimes (see DISCUSSION). The D83N histogram was reliably fit to the sum of two Gaussian functions centered at 3.7 and 10.7 ms, still leaving some residuals. These time constants are comparable with the two fastest time constants measured in the first 100 ms of WT R2 relaxations, whereas the third
found in WT appears to be missing (Table 1). The E134Q histogram was distinctly different from both WT and D83N, requiring only a single Gaussian function with a peak centered at 4.4 ms. This single peak overlaps with the fastest time constant seen in the WT and D83N pigments (Table 1). This work establishes an initial approach to parameterize the R2 relaxation. The intent is to use this approach as a means to quantify differences between WT and mutant ERC kinetics during the biochemically important time period of the R2 signal.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In earlier work, we established that, after regeneration with 11cRet, ERC signals could be recorded from both single and fused HEK293S cells expressing high levels of WT human opsin to their plasma membranes (
In the current experiments, we applied the expression ERC tool to investigate physical properties of WT rhodopsin expressed in HEK293S cells. ERC measurements are consistent with normal properties of WT human rhodopsin such as the absorbance spectrum, photosensitivity, univariance, and photoreversibility from Meta-II. The utility of the expression ERC tool was expanded on in the experiments reported here by demonstrating successful measurements of ERCs in cells regenerated with the analogue chromophore 9cRet. Moreover, ERCs of two mutant pigments D83N and E134Q demonstrate qualitative and quantitative differences with respect to WT ERCs. These results strongly suggest that the expression ERC approach could be productively expanded to investigate a broad range of rhodopsin activation properties of analogue visual pigments and mutant visual pigments, thus embracing a structurefunction approach applied to both the chromophore and remote environments of rhodopsin.
On Recording ERCs from Expressed and Regenerated Visual Pigments in Giant HEK293S Cells
When fused giant cells are regenerated (primary) with 11cRet in the dark, ERC signals are obtained on high intensity flash stimulation (108 photons/µm2) with essentially identical waveform to the ERCs of vertebrate amphibian photoreceptors (
To regenerate rhodopsin from opsin apoprotein constitutively expressed in these cells, a FAF-BSA technique previously used to regenerate rhodopsin from opsin in intact rod outer segments (3 min) are similar to rates in intact photoreceptors (
1/2
1 min, complete by 10 min;
Experiments reported here indicated that the source of 11cRet during spontaneous dark regeneration is internal to the cell that is recorded. Several bleach and recovery cycles were required to completely exhaust the source of chromophore in the presence of extracellular NH2OH. If regenerating 11cRet chromophore originated from the outer surface of the plasma membrane or external to the cell (e.g., released by other cells on the coverslip or from recording chamber surfaces), then NH2OH should rapidly prevent ERC recovery by converting retinaldehydes to oximes, which do not form visual pigments. The only other potential source of chromophore is internal to the cell under recording, shielded from reaction with extracellular 10 mM NH2OH. Once NH2OH was applied to the bath, plasma membrane visual pigment was partially bleached and spontaneously regenerated over several cycles before the source of intracellular chromophore became exhausted and ERCs did not recover. Prompt recovery followed washout of NH2OH and perfusion of 25 µM 11cRet plus FAF-BSA in recording buffer. These experiments suggest that 11cRet can enter the retinal binding pocket when presented from either side of the membrane. The data also provides indirect evidence that NH2OH is impeded from permeating the plasma membrane of the HEK293 cells in E-1/I-1 solutions because if it had, then the spontaneous regeneration should have been promptly quenched. That ERCs are recordable when NH2OH has access to the extracellular surface of membrane-oriented rhodopsin indicates that visual pigment regenerated in the dark is not reacting with this agent, consistent with the resistance of the ground state of rod rhodopsin to NH2OH. NH2OH does not affect the rate of formation of Meta-II (
One of the major advantages of the FAF-BSA/-D-tocopherol regeneration technique in fused giant cells is the spontaneous recovery of visual pigment and ERC signals without the need for reinstallation of 11cRet into the recording chamber. A similar process has been reported for retina isolated away from pigment epithelium and apparently reflects limited photoreceptor stores of 11cRet (
1/2
10 min) was similar to the decay of Meta-II at 25°C (
7 min half-life) and proposed that 11cRet enters the ligand-binding pocket only after all-trans-retinal has vacated the environment. FAF-BSA can be viewed as a nonspecific retinoid binding protein (
Spontaneous regeneration of visual pigment after 11cRet loading in FAF-BSA/vitamin E provides parsimonious support of ERC experiments. The regeneration of ERCs in WT-HEK293S cells from an internal source of 11cRet provides a clear advantage for complex experiments where charge motions must be compared against different conditions (e.g., action spectra determination). Two properties of ERCs measured in this environment are under further active investigation (Brueggemann and Sullivan, manuscript in preparation). First is the loss of the inward R1 signal after post-bleach spontaneous regeneration. The time to peak of the R2 signal is slowed approximately twofold when the R1 signal is present (
The Ground State of Rhodopsin Is the Source of the ERC Signal
Strong evidence is demonstrated that ERC signals result from activation of the ground state of human rhodopsin in the plasma membrane of fused giant cells and that no other additional spectral states are contributing to these measurements. Evidence supports the conclusion that the identity of the chromophore regenerating visual pigment is 11cRet, which forms a PSB+-H with K296 of the opsin apoprotein. The action spectrum of the ERC of WT human rhodopsin, when stimulated through 30-nm bandpass filters, was well fit by a scaled normalized template of the absorbance spectrum of WT human rhodopsin purified from the same cell line used for ERC studies. The action spectrum was obtained from cells that had already had their ERCs extinguished and had undergone spontaneous recovery. These experiments allow the conclusion that the ground state of rhodopsin is the one that regenerates upon dark adaptation after primary ERC extinction. Thus, during dark adaptation, the Meta-II state must decay with hydrolysis of the Schiff base to form opsin, which then can subsequently react with 11cRet to form a protonated Schiff base at K296. From our previous work on spectral sensitivity, which was conducted with 70-nm bandpass filters, the action spectra was much broader than rhodopsin absorbance. Data from the experiments reported here shows that this outcome was the result of stimulus bandwidth, and not the presence of additional spectral states that were contributing to charge motion. Specifically, there were three pigments that could have contributed to "recovery" of the ERC when later stimulated with 500-nm stimuli: Meta-III465 and Meta470, both resulting from thermal Meta-II380 decay (3.0 min). However, if these states contributed substantially to the ERC, the action spectrum would not have been fit by the absorbance of rhodopsin and the signals should have been promptly extinguished by 10 mM NH2OH (
R2 Extinction Photosensitivity Is Consistent with Rhodopsin Activation
The ERC signal can be extinguished with successive flashes in a fashion similar to rhodopsin bleaching in photoreceptors and with similar photosensitivity. The mean and maximum Pt values (2.6 x 10-9 and 5.0 x 10-9 µm2, respectively) determined from exponential extinction decays are consistent with a rhodopsin photopigment but are lower than expected (1.0 x 10-8 µm2), taking the product of the molecular cross section of human rhodopsin at 493 nm (493 = 1.528 x 10-8 µm2, see MATERIALS AND METHODS) and the known quantal efficiency (
= 0.67) (
30,000 vs. 40,000 M-1 · cm-1) (
Photoregeneration from early intermediate states with high quantal efficiency for photoconversion (e.g., bathorhodopsin, lumirhodopsin) can definitively lower Pt estimates for the absorbing pigment ( Meta-II equilibrium. Flashes of this duration were, however, unlikely to overlap significantly with the lifetimes of earlier intermediates that have higher quantal efficiency of photoconversion (
Meta-II equilibrium, which correlates in part with the R2 time course (
of 0.21 that is only ~32% of the known value of 0.67 determined from small bleaches.
How could be decreased to explain the suppression of Pt? In fact,
is a function of light intensity. When the flash intensity is infinitely strong,
should approach zero.
is expected to decrease in proportion to the intensity of the flash stimulation because photoconversion becomes increasingly likely (even-numbered absorptions) from intermediates with lifetimes of the same order as the flash duration.
5 µs). High intensities were used to obtain good estimates of the cross section (
) in the spectral measurements and were maintained throughout because of the high SNR obtained in ERC recordings. In fact, Pt approximates the value expected when stimulus intensities (500 nm) are nearly a log-fold lower (see RESULTS). In future experiments, the effects of a range of flash intensities on Pt and quantal efficiency will be investigated. In addition, we will also investigate the mechanism for the marked suppressive effect (
1 log) of NH2OH on Pt, which appears to be a novel finding. We speculate that NH2OH has the capacity to bind in retinal binding pocket and perhaps alter the local chromophore environment in the ground state in such a manner as to result in altered photochemical properties.
Photoregeneration of Meta-II Can Be Detected with ERC Measurements
Experiments were conducted to test whether the ERC can be used to study rhodopsin photoregeneration from the Meta-II state. These experiments were motivated by earlier studies that reported reversed ERP signals from the Meta-II state upon near UV stimulation (
9-cis-Retinal Regenerates ERCs from WT Opsin
When WT-HEK293S cells are regenerated with 9cRet, ERC signals are recordable upon 500-nm flash photolysis. 9c-Ret regenerates a visual pigment, presumed to be isorhodopsin (peak 483 nm), which would broadly overlap with the 70-nm band stimulus. That ERC signals can be recorded from a "natural" analogue pigment provides evidence to support a role for the ERC in investigation of a wide variety of analogue visual pigments that are known to have unique properties. For example, certain locked analogues are known to prevent energy uptake by preventing isomerization (
Kinetic Analysis of R2 ERC Signals of WT and Mutant (D83N and E134Q) Pigments
A major challenge in structurefunction studies of visual pigments has been the preparation of sufficient mutant protein in expression systems for biophysical or biochemical analysis. The ERC method improves detection sensitivity by 107108-fold, allowing measurements of conformational activation in an ensemble of regenerated rhodopsin molecules in the physiologically intact environment of a single fused giant cell ( helix was found to perturb the kinetics of the R2 ERC phase. A mutation on the cytoplasmic face of the third
helix (E134Q), at a residue known to be a gatekeeper to both proton uptake and the related generation of the transducin docking environment, results in loss of most of the ERC R2 signal except for the fast initial process.
The R2 signal is well resolved in cellular ERC measurements and overlaps temporally with critical events leading to biochemical activation of rhodopsin. WT rhodopsin R2 relaxation kinetics are an easily accessible aspect of the total ERC signal and are invariant to intensity or wavelength of stimulation as found in these experiments. Therefore, the R2 relaxation should be a reliable parameter to evaluate for understanding conformational dynamics. A quantitative analysis of R2 relaxation in WT rhodopsin should serve not only to begin characterization of the state complexity of charge motion on the millisecond time scale, but also serve for comparison of mutant pigments that might be affected in some aspect of R2 electrical state transitions. WT rhodopsin R2 relaxation was first characterized by fitting sums of exponentials to a set of WT ERC waveforms and generating an ensemble of time constants (a,b). We assumed that the ensemble of time constants would represent the kinetic aspects of the signal even with any slight heterogeneity (e.g., due to cell size) that might affect only the fastest aspects of the relaxation (
a or
b is somewhat arbitrary given that the short time constant may not have been weighted or fit in a particular ERC signal and a longer time constant was then assigned to
a. In fact, there is some overlap in the assignments (
a,b ensemble. We attempted fits of one, two, three, and four Gaussians to the data. With this data set, a reliable fit of the sum of three Gaussians was obtained that was independent of bin width. Three Gaussian peaks were centered around 4.1, 12.45, and 26.4 ms, and the errors in the fitting results are shown in Table 1. The natural conclusion from this analysis is that there are a minimum of three distinct electrically active states in the WT R2 relaxation at room temperature in WT rhodopsin. We speculate that the underlying molecular basis of the three fitted time constants are likely to represent processes related to Schiff base deprotonation, proton uptake, and
helical movements associated with conformational activation. However, other interpretations are possible. For example, the capacitative, or AC-coupled nature of the ERC could reflect forward and reverse movements of charges (e.g., protons, sidechains), in particular molecular environments, leading to a series of coordinated charge separation and recombination events, each satisfying a zero-time integral expected for pure capacitative components (e.g., see
ERC signals from D83N rhodopsin (regenerated with 11cRet) appear slightly simplified in comparison with WT. The R2 signal appears WT in nature, but shorter in comparison to the broad stretched relaxation of the WT pigment. D83N ERCs were subjected to R2 relaxation time constant analysis, and the sum of two Gaussian distributions was required to best fit the total time constant histogram. Peaks were identified at 3.67 and 10.67 ms, and the errors in the fitting are shown in Table 1. These times appear comparable to the fast and medium time constants in the WT pigment that was measured under identical conditions. The third and longest time constant of the R2 relaxation in WT (26.4 ms) is missing in the D83N pigment. We suspect this is likely to represent a distinct property of the D83N pigment since R2 relaxation in D83N ERCs have a different waveform compared with WT. In summary, D83N loses the R1 signal during primary bleaches and its R2 relaxation loses the "stretched exponential" characteristic of the WT R2 relaxation.
The D83N mutation is known to slightly blue-shift the absorbance of rhodopsin, which is consistent with an alteration of the Schiff base environment in the ground state pigment, resulting from the isomorphic loss of the protonated carboxyl group of aspartate (
ERC signals from E134Q rhodopsin have little complexity in comparison with WT and demonstrate no R1 and a brief R2 relaxation always well fit by a single exponential. When R2 relaxation time constant analysis was applied to the E134Q pigment, a single Gaussian was needed to fit the time-constant histogram. The value of the peak was 4.4 ms, which was comparable within error to WT fast phase. The timing of the fast phase did not appear to change when the cytoplasmic pH was held at different constant values over the range from 6.0 to 8.0. The E134 sidechain is known to be essential to proton uptake into the cytoplasmic face of rhodopsin during normal Meta-II formation given that the E134Q mutant binds zero protons in comparison with the two adsorbed by the WT pigment ( helix VI relative to
helix III (the location of E134/R135) has been detected. This results in helix VI being displaced outward from the disk membrane into the cytoplasm, although the magnitude of the vectorial movements are not yet clear (
helices in the dark that are not affected by light and do not affect the subsequent movement of
helix VI. What results is a partial activation of the receptor in the darkness or constitutive activity (
helices may be normally coupled, but can still occur independently in the presence of environmental perturbation (e.g., mutation). Since spectral Meta-II formation is not perturbed in E134Q, we suggest that the residual fast ERC signal in this mutant represents the charge motion associated with Schiff base deprotonation. In preliminary experiments, the residual R2 time constant of E134Q did not demonstrate sensitivity to intracellular pH over the range of 6.0 to 8.0 and might reflect the pH-insensitive component of the ERP related to Schiff base deprotonation (
helices). If so, then these later time constants in the WT pigment should be responsive to changes in intracellular pH or to molecular conditions that prevent the conformational activation of the molecule that allows transducin binding. Further experiments will be necessary to decipher the molecular origin of the charge motions during R2 and how local environment affects these transitions.
We were surprised that no R1 signals were found in fused E134Q giant cells given that the E134 sidechain resides at the cytoplasmic face of the third helix remote from the Schiff base environment where R1 is thought to originate. However, total charge motion in E134Q cells is smaller than in WT cells and the R1 signal may simply have been lost in noise, as we found for R1 in single unfused cells expressing WT pigment regenerated with 11cRet (
Although we do not yet have a molecular understanding of the simplification of the ERC R2 signal in D83N or E134Q, such mutant pigments with uniquely different ERC signals constitute a substrate to understand the components of the normal ERC signal in WT rhodopsin. At this point, our efforts have clearly established the feasibility of using the ERC to study time-resolved and electrically active conformational changes during rhodopsin activation in WT and two mutant pigments. Future studies will focus on the molecular mechanism of R* formation and the forces that govern the molecular volume increase of the pigment during the Meta-IIa to Meta-IIb transition in WT and mutant pigments (
Summary and Conclusions
The sensitivity of the expression ERC method is greatly improved when compared with other contemporary time-resolved techniques applied to rhodopsin activation and allows measurement of rhodopsin activation in single cells containing about a picogram of visual pigment. The sensitivity of this approach is dependent upon the use of gigaohm-seal, whole-cell patch clamping techniques that are capable of resolving small currents with fast time resolution. The ERC is a conformation-dependent charge motion of the same family as ionic channel gating currents. Gating current analysis of expressed mutant ionic channels has significantly extended knowledge of molecular processes of conformational activation of ionic channels, and for this reason we are applying the same approach to visual pigments that are known to be electrically active proteins. In this work, we apply the expression ERC approach to measure properties of WT rhodopsin and found that the ERC signal results from activation of the ground state of rhodopsin, given the action spectra and bleaching photosensitivity. ERC kinetics are independent of wavelength, as expected for the univariance principle, and rhodopsin can be photoregenerated from the Meta-II state. The major impact of this study resides in our success at recording ERCs from an analogue visual pigment (isorhodopsin) and from two mutant human opsins regenerated with 11cRet. This study establishes the feasibility of an ERC structure/function investigation applied to the ligand binding pocket, as probed by artificial chromophores or site-specific mutations, or remote regions of the pigment, as probed by engineered mutations or chemically reactive ligands. The ERP has been similarly used in expressed mutant bacteriorhodopsin pigments to understand proton exchange mechanisms (e.g.,
![]() |
Acknowledgements |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The authors thank Drs. Michael Sheets and J. Hugh McDowell for sharing their methods on recording ionic channel gating currents from fused transfected HEK293 cells and on FAF-BSA rhodopsin regeneration, respectively. The authors acknowledge the technical assistance of Michael F. Satchwell with cell cultures. The authors appreciate the critical review of the manuscript by Drs. Bob Birge, Ken Foster, and Barry Knox before its submission and the comments of the reviewers before publication.
This work was funded by an R29 award (EY11384) from the National Eye Institute to J.M. Sullivan. Also supported, in part, by start-up funds from the Department of Ophthalmology at SUNY (which is a recipient of a Research to Prevent Blindness Award).
Submitted: 11 November 1999
Revised: 2 August 1999
Accepted: 9 August 1999
1used in this paper: 11cRet, 11-cis-retinal; Cmem, membrane capacitance; ERC, early receptor current; ERP, early receptor potential; FAF, fatty acidfree; FWHM, full-width-half-maximum; PSB+-H, protonated Schiff base; Pt, photosensitivity; SNR, signal-to-noise ratio; WT, wild type
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Altenbach, C., Yang, K., Farrens, D.L., Farahbakhsh, Z.T., Khorana, H.G., Hubbell, W.L. 1996. Structural features and light-dependent changes in the cytoplasmic interhelical E-F loop region of rhodopsin: a site-directed spin-labeling study. Biochemistry. 35:13470-13478.
Arden, G.B., Ikeda, H., Siegel, I.M. 1966. New components of the mammalian receptor potential and their relation to visual photochemistry. Vision Res. 6:373-384[Medline].
Arnis, S., Hofmann, K.P. 1993. Two different forms of metarhodopsin II: Schiff base deprotonation precedes proton uptake and signaling state. Proc. Natl. Acad. Sci. USA. 90:7849-7853
Arnis, S., Hofmann, K.P. 1995. Photoregeneration of bovine rhodopsin from its signaling state. Biochemistry. 34:9333-9340[Medline].
Arnis, S., Fahmy, K., Hofmann, K.P., Sakmar, T.P. 1994. A conserved carboxylic acid group mediates light-dependent proton uptake and signaling by rhodopsin. J. Biol. Chem. 269:23879-23881
Baldwin, J.M. 1993. The probable arrangement of the helices in G proteincoupled receptors. EMBO (Eur. Mol. Biol. Organ.) J. 12:1693-1703[Abstract].
Bennett, N., Michel-Villaz, M., Dupont, Y. 1980. Cyanine dye measurement of a light-induced transient membrane potential associated with the Metarhodopsin II intermediate in rod-outer-segment membranes. Eur. J. Biochem. 111:105-109[Abstract].
Bezanilla, F., Stefani, E. 1994. Voltage-dependent gating of ionic channels. Annu. Rev. Biophys. Biomol. Struct. 23:819-846[Medline].
Bhattacharya, S., Ridge, K.D., Knox, B.E., Khorana, H.G. 1992. Light-stable rhodopsin. I. A rhodopsin analog reconstituted with a nonisomerizable 11-cis retinal derivative. J. Biol. Chem. 267:6763-6769
Birge, R.R., Einterz, C.M., Knapp, H.M., Murray, L.P. 1988. The nature of the primary photochemical events in rhodopsin and isorhodopsin. Biophys. J. 53:367-385[Abstract].
Birge, R.R. 1990a. Nature of the primary photochemical events in rhodopsin and bacteriorhodopsin. Biochim. Biophys. Acta. 1016:293-327[Medline].
Birge, R.R. 1990b. Photophysics and molecular electronic applications of the rhodopsins. Annu. Rev. Phys. Chem. 41:683-733[Medline].
van Bruegel, P.J.G.M., Bovee-Geurts, P.H.M., Bonting, S.L., Daemen, F.J.M. 1979. Biochemical aspects of the visual process XL. Spectral and chemical aspects of metarhodopsin III in photoreceptor membrane suspensions. Biochim. Biophys. Acta. 557:188-198[Medline].
Cafiso, D.S., Hubbell, W.L. 1980. Light-induced interfacial potentials in photoreceptor membranes. Biophys. J. 30:243-264[Abstract].
Chen, C., Okayama, H. 1988. Calcium phosphate-mediated gene transfer: a highly efficient transfection system for stably transforming cells with plasmid DNA. Biotechniques. 6:632-637[Medline].
Cohen, G.B., Yang, T., Robinson, P.R., Oprian, D.D. 1993. Constitutive activation of opsin: influence of charge at position 134 and size at position 296. Biochemistry. 32:6111-6115[Medline].
Cone, R.A. 1967. Early receptor potential: photoreversible charge displacement in rhodopsin. Science. 155:1128-1131[Medline].
Cone, R.A., Brown, P.K. 1967. Dependence of the early receptor potential on the orientation of rhodopsin. Science. 156:536.
Cone, R.A., Brown, P.K. 1969. Spontaneous regeneration of rhodopsin in the isolated rat retina. Nature. 221:818-820[Medline].
Cone, R.A., Cobbs, W.H. 1969. Rhodopsin cycle in the living eye of the rat. Nature. 221:820-822[Medline].
Cone, R.A., Pak, W.L. 1971. The early receptor potential. In Lowenstein W.R., ed. Handbook of Sensory Physiology: Principles of Receptor Physiology. Volume 1. Berlin, Springer Verlag, 345-365.
Crouch, R., Purvin, V., Nakanishi, K., Ebrey, T. 1975. Isorhodopsin II: artificial photosensitive pigment formed from 9,13-Dicis retinal. Proc. Natl. Acad. Sci. USA. 72:1538-1542[Abstract].
Dartnall, H.J.A. 1968. The photosensitivities of visual pigments in the presence of hydroxylamine. Vision Res. 8:339-358[Medline].
Dilley, R.A., McConnell, D.G. 1970. Alpha-tocopherol in the retinal outer segment of bovine eyes. J. Membr. Biol. 2:317-323.
Drachev, L.A., Kalamkarov, G.R., Kaulen, A.D., Ostrovsky, M.A., Skulachev, V.P. 1981. Fast stages of photoelectric processes in biological membranes. II. Visual rhodopsin. Eur. J. Biochem. 117:471-481[Abstract].
Ebrey, T.G. 1968. The thermal decay of the intermediates of rhodopsin in situ. Vision Res. 8:965-982[Medline].
Ernst, O.P., Hofmann, K.P., Sakmar, T.P. 1995. Characterization of rhodopsin mutants that bind transducin but fail to induce GTP nucleotide uptake. J. Biol. Chem. 270:10580-10586
Fahmy, K., Jager, F., Beck, M., Zvyaga, T.A., Sakmar, T.P., Siebert, F. 1993. Protonation states of membrane-embedded carboxylic acid groups in rhodopsin and metarhodopsin II: a Fourier-transform infrared spectroscopy study of site-directed mutants. Proc. Natl. Acad. Sci. USA. 90:10206-10210[Abstract].
Fahmy, K., Sakmar, T.P. 1993. Regulation of the rhodopsintransducin interaction by a highly conserved carboxylic acid group. Biochemistry. 32:7229-7236[Medline].
Fahmy, K., Siebert, F., Sakmar, T.P. 1995. Photoactivated state of rhodopsin and how it can form. Biophys. Chem. 56:171-181[Medline].
Farahbakhsh, Z.T., Hideg, K., Hubbell, W.L. 1993. Photoactivated conformational changes in rhodopsin: a time-resolved spin label study. Nature. 262:1416-1419.
Farahbakhsh, Z.T., Ridge, K.D., Khorana, H.G., Hubbell, W.L. 1995. Mapping light-dependent structural changes in the cytoplasmic loop connecting helices C and D in rhodopsin: a site-directed spin labeling study. Biochemistry. 34:8812-8819[Medline].
Farrens, D.L., Altenbach, C., Yang, K., Hubbell, W.L., Khorana, H.G. 1996. Requirement of rigid-body motion of transmembrane helices for light activation of rhodopsin. Science. 274:768-770
Franke, R.R., Sakmar, T.P., Graham, R.M., Khorana, H.G. 1992. Structure and function in rhodopsin. Studies of the interaction between the rhodopsin cytoplasmic domain and transducin. J. Biol. Chem. 267:14767-14774
Ganter, U.M., Gartner, W., Siebert, F. 1988. Rhodopsin-lumirhodopsin phototransition of bovine rhodopsin investigated by Fourier transform infrared difference spectroscopy. Biochemistry. 27:7480-7488[Medline].
Ganter, U.M., Schmid, E.D., Perez-Sala, D., Rando, R.R., Siebert, F. 1989. Removal of the 9-methyl group of retinal inhibits signal transduction in the visual process. A Fourier transform infrared and biochemical investigation. Biochemistry. 28:5954-5962[Medline].
Govardovskii, V.I. 1979. Mechanism of the generation of the early receptor potential and an electrical model of the rod of the intact rat retina. Biophysics. 23:520-526.
Haeseleer, F., Huang, J., Lebioda, L., Saari, J.C., Palczewski, K. 1998. Molecular characterization of a novel short-chain dehydrogenase/reductase that reduces all-trans-retinal. J. Biol. Chem. 273:21790-21799
Hagins, W.A. 1955. The quantum efficiency of bleaching of rhodopsin in situ. J. Physiol. 129:22P.
Han, M., Lin, S.W., Minkova, M., Smith, S.O., Sakmar, T.P. 1996. Functional interaction of transmembrane helices 3 and 6 in rhodopsin. J. Biol Chem. 271:32337-32342
Hestrin, S., Korenbrot, J.I. 1990. Activation kinetics of retinal cones and rods: response to intense flashes of light. J. Neurosci. 10:1967-1973[Abstract].
Hodgkin, A.L., O'Bryan, P.M. 1977. Internal recording of the early receptor potential in turtle cones. J. Physiol. 267:737-766[Abstract].
Hofmann, K.P., Pulvermuller, A., Buczylko, J., Van Hooser, P., Palczewski, K. 1992. The role of arrestin and retinoids in the regeneration pathway of rhodopsin. J. Biol. Chem. 267:15701-15706
Hong, F.T., Hong, F.H., Needleman, R.B., Ni, B., Chang, M. 1992. Modifying the photoelectric behavior or bacteriorhodopsin by site-directed mutagenesis: electrochemical and genetic engineering approaches to molecular events. In Aviram A., ed. Molecular ElectronicsScience and Technology. AIP Conference Proceedings, American Institute of Physics, 262. 204217.New York, NY.
Honig, B.H., Hubbell, W.L., Flewelling, R.F. 1986. Electrostatic interactions in membranes and proteins. Annu. Rev. Biophys. Chem. 15:163-193[Medline].
Jager, F., Fahmy, K., Sakmar, T.P., Siebert, F. 1994. Identification of glutamic acid 113 as the Schiff base proton acceptor in the metarhodopsin II photointermediate of rhodopsin. Biochemistry. 33:10878-10882[Medline].
Johnson, R.H. 1970. Absence of effect of hydroxylamine upon production rates of some rhodopsin photo intermediates. Vision Res. 10:897-900[Medline].
Jones, G.J., Crouch, R.K., Wiggert, B., Cornwall, M.C., Chader, G.J. 1989. Retinoid requirements for recovery of sensitivity after visual pigment bleaching in isolated photoreceptors. Proc. Natl. Acad. Sci. USA. 86:9606-9610[Abstract].
Kersting, U., Joha, H., Steigner, W., Gassner, B., Gstaunthaler, G., Pfaller, W., Oberleithner, H. 1989. Fusion of cultured dog kidney (MDCK) cells: I. technique, fate of plasma membranes and of cell nuclei. J. Membr. Biol. 111:37-48[Medline].
Khorana, G. 1992. Rhodopsin, photoreceptor of the rod cell. An emerging pattern for structure and function. J. Biol. Chem. 267:1-4
Kim, J.-M., Altenbach, C., Thurmond, R.L., Khorana, H.G., Hubbell, W.L. 1997. Structure and function in rhodopsin: rhodopsin mutants with a neutral amino acid at E134 have a partially activated conformation in the dark state. Proc. Natl. Acad. Sci. USA. 94:14273-14278
Knowles, A., Dartnall, H.J.A. 1977. The photobiology of vision. In Davson H., ed. The Eye. New York, NY, Academic Press, Inc., 247-497.
Lamola, A.A., Yamane, T., Zipp, A. 1974. Effect of detergents and high pressure upon the metarhodopsin I metarhodopsin II equilibrium. Biochemistry. 13:738-745[Medline].
Lewis, J.W., Einterz, C.M., Hug, S.J., Kliger, D.S. 1989. Transition dipole orientations in the early photolysis intermediates of rhodopsin. Biophys. J. 56:1101-1111[Abstract].
Lewis, J.W., van Kuijk, F.J.G.M., Carruthers, J.A., Kliger, D.S. 1997. Metarhodopsin III formation and decay kinetics: comparison of bovine and human rhodopsin. Vision Res. 37:1-8[Medline].
Lindau, M., Ruppel, H. 1985. On the nature of the fast light-induced charge displacement in vertebrate photoreceptors. Photobiochem. Photobiophys. 9:43-56.
Livrea, M.A., Tesoriere, L., Bongiorno, A. 1991. All-trans to 11-cis retinol isomerization in nuclear membrane fraction from bovine retinal pigment epithelium. Exp. Eye Res. 52:451-459[Medline].
Longstaff, C., Calhoon, R., Rando, R.R. 1986. Deprotonation of the Schiff base of rhodopsin is obligatory in the activation of the G protein. Proc. Natl. Acad. Sci. USA. 83:4209-4213[Abstract].
Makino, C.L., Taylor, W.R., Baylor, D.A. 1991. Rapid charge movements and photosensitivity of visual pigments in salamander rods and cones. J. Physiol. 442:761-780[Abstract].
McDowell, J.H. 1993. Preparing rod outer segment membranes, regenerating rhodopsin, and determining rhodopsin concentration. Methods Neurosci. 15:123-130.
Misra, S. 1998. Contribution of proton release to the B2 photocurrent of bacteriorhodopsin. Biophys. J. 75:382-388
Moltke, S., Heyn, M.P., Krebs, M.P., Mollaaghababa, R., Khorana, H.G. 1992. Low pH photovoltage kinetics of bacteriorhodopsin with replacements of Asp-96, -85, -212 and Arg-82. In Rigaud J.L., ed. Structures and Functions of Retinal Proteins. Paris, France, Colloque INSERM, John Libbey Eurotext Ltd, 221:201-204.
Nathans, J., Weitz, C.J., Agarwal, N., Nir, I., Papermaster, D.S. 1989. Production of bovine rhodopsin by mammalian cell lines expressing cloned cDNA: spectrophotometry and subcellular localization. Vision Res. 29:907-914[Medline].
Nathans, J. 1990. Determinants of visual pigment absorbance: role of charged amino acids in the putative transmembrane segments. Biochemistry. 29:937-942[Medline].
Noy, N., Xu, Z.-J. 1990. Thermodynamic parameters of the binding of retinol to binding proteins and to membranes. Biochemistry. 29:3888-3892[Medline].
Okajima, T.L, Hong, F.T. 1986. Kinetic analysis of displacement photocurrents elicited in two types of bacteriorhodopsin model membranes. Biophys. J. 50:901-912[Abstract].
Ostroy, S.E. 1974. Hydrogen ion changes of rhodopsin. pK changes and the thermal decay of metarhodopsin II380. Arch. Biochem. Biophys. 164:275-284[Medline].
Parkes, J.H., Liebman, P.A. 1984. Temperature and pH dependence of the metarhodopsin Imetarhodopsin II kinetics and equilibria in bovine rod disk membrane suspensions. Biochemistry. 23:5054-5061[Medline].
Rath, P., DeCaluwe, L.L.J., Bovee-Geurts, P.H.M., DeGrip, W.J., Rothschild, K.J. 1993. Fourier transform infrared difference spectroscopy of rhodopsin mutants: light activation of rhodopsin causes hydrogen-bonding change in residue aspartic acid-83 during Meta II formation. Biochemistry. 32:10277-10282[Medline].
Reeves, P.J., Thurmond, R.L., Khorana, H.G. 1996. Structure and function in rhodopsin: high level expression of a synthetic bovine opsin gene and its mutants in stable mammalian cell lines. Proc. Natl. Acad. Sci. USA. 93:11487-11492
Rotmans, J.P., Daemen, F.J.M, Bonting, S.L. 1974. Biochemical aspects of the visual process. XXVI. Binding site and migration of retinaldehyde during rhodopsin photolysis. Biochim. Biophys. Acta. 357:151-158[Medline].
Sakmar, T.P., Franke, R.R., Khorana, H.G. 1989. Glutamic acid 113 serves as the retinylidene Schiff base counterion in bovine rhodopsin. Proc. Natl. Acad. Sci. USA. 86:8309-8313[Abstract].
Schneider, E.E., Goodeve, C.F., Lythgoe, R.J. 1939. The spectral variation of the photosensitivity of visual purple. Proc. R. Soc. Lond. A. 170:102-112.
Schoenlein, R.W., Peteanu, L.A., Mathies, R.A., Shank, C.V. 1991. The first step in vision: femtosecond isomerization of rhodopsin. Science. 254:412-415[Medline].
Sengbusch, G.V., Stieve, H. 1971. Flash photolysis of rhodopsin. I. Measurements on bovine rod outer segments. Z. Naturforschung. 26B:488-489.
Sheets, M.F., Kyle, J.W., Krueger, S., Hanck, D.A. 1996. Optimization of a mammalian expression system for the measurement of sodium channel gating currents. Am. J. Physiol. 271:C1001-C1006
Shieh, T., Han, M., Sakmar, T.P., Smith, S.O. 1997. The steric trigger in rhodopsin activation. J. Mol. Biol. 269:373-384[Medline].
Shukla, P., Sullivan, J.M. 1998. Rhodopsin early receptor currents from fused giant cells expressing opsin. Invest. Ophthal. Vis. Sci. 39:974a. (Abstr.).
Spalink, J.D., Stieve, H. 1980. Direct correlation between the R2 component of the early receptor potential and the formation of metarhodopsin II in the excised bovine retina. Biophys. Struct. Mech. 6:171-174[Medline].
Stecher, H., Gelb, M.H., Saari, J.C., Palczewski, K. 1999. Preferential release of 11-cis-retinol from retinal pigment epithelial cells in the presence of cellular retinaldehyde-binding protein. J. Biol. Chem. 274:8577-8585
Sullivan, J.M. 1996. Rhodopsin early receptor currents following flash photolysis of single cells expressing human rhodopsin. Invest. Ophthal. Vis. Sci. 37:811a. (Abstr.).
Sullivan, J.M. 1998. Low-cost monochromatic microsecond flash microbeam apparatus for single-cell photolysis of rhodopsin or other photolabile pigments. Rev. Sci. Instrum. 69:527-539.
Sullivan, J.M., Brueggemann, L., Shukla, P. 2000. An electrical approach to study rhodopsin activation in single cells with the early receptor current assay. Methods Enzymol. In press.
Sullivan, J.M., Satchwell, M.F. 2000. Development of stable cell lines expressing high levels of point mutants of human opsin for biochemical and biophysical studies. Methods Enzymol. In press.
Sullivan, J.M., Shukla, P. 1999. Time-resolved rhodopsin activation currents in a unicellular expression system. Biophys. J. In press.
Szuts, E.Z., Harosi, F.I. 1991. Solubility of retinoids in water. Arch. Biochem. Biophys. 287:297-304[Medline].
Trissl, H.W. 1982. On the rise time of the R1-component of the "early receptor potential": evidence for a fast light-induced charge separation in rhodopsin. Biophys. Struct. Mech. 8:213-230[Medline].
Wald, G., Brown, P.K. 1958. Human rhodopsin. Science. 127:222-226[Medline].
Weitz, C.J., Nathans, J. 1993. Rhodopsin activation: effects on the metarhodopsin Imetarhodopsin II equilibrium of neutralization or introduction of charged amino acids within putative transmembrane segments. Biochemistry. 32:14176-14182[Medline].
Williams, T.P. 1964. Photoreversal of rhodopsin bleaching. J. Gen. Physiol. 47:679-689
Williams, T.P. 1965. Rhodopsin bleaching: relative effectiveness of high and low intensity flashes. Vision Res. 5:633-638[Medline].
Williams, T.P. 1966. Limitations on the use of the concept of quantum efficiency in rhodopsin bleaching. Nature. 209:1350-1351[Medline].
Williams, T.P. 1968. Photolysis of metarhodopsin II. Rates of production of P470 and rhodopsin. Vision Res. 8:1457-1466[Medline].
Williams, T.P. 1974. Upper limits to the bleaching of rhodopsin by high intensity flashes. Vision Res. 14:603-607[Medline].
Williams, T.P., Breil, S.J. 1968. Kinetic measurements on rhodopsin solutions during intense flashes. Vision Res. 8:777-786[Medline].
Winston, A., Rando, R.R. 1998. Regulation of isomerohydrolase activity in the visual cycle. Biochemistry. 37:2044-2050[Medline].
Zhukovsky, E.A., Oprian, D.D. 1989. Effect of carboxylic acid side chains on the absorption maximum of visual pigments. Science. 246:928-930[Medline].