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• W2076770170 abstract "In this study, we address the mechanism of visual arrestin release from light-activated rhodopsin using fluorescently labeled arrestin mutants. We find that two mutants, I72C and S251C, when labeled with the small, solvent-sensitive fluorophore monobromobimane, exhibit spectral changes only upon binding light-activated, phosphorylated rhodopsin. Our analysis indicates that these changes are probably due to a burying of the probes at these sites in the rhodopsin-arrestin or phospholipid-arrestin interface. Using a fluorescence approach based on this observation, we demonstrate that arrestin and retinal release are linked and are described by similar activation energies. However, at physiological temperatures, we find that arrestin slows the rate of retinal release ∼2-fold and abolishes the pH dependence of retinal release. Using fluorescence, EPR, and biochemical approaches, we also find intriguing evidence that arrestin binds to a post-Meta II photodecay product, possibly Meta III. We speculate that arrestin regulates levels of free retinal in the rod cell to help limit the formation of damaging oxidative retinal adducts. Such adducts may contribute to diseases like atrophic age-related macular degeneration (AMD). Thus, arrestin may serve to both attenuate rhodopsin signaling and protect the cell from excessive retinal levels under bright light conditions. In this study, we address the mechanism of visual arrestin release from light-activated rhodopsin using fluorescently labeled arrestin mutants. We find that two mutants, I72C and S251C, when labeled with the small, solvent-sensitive fluorophore monobromobimane, exhibit spectral changes only upon binding light-activated, phosphorylated rhodopsin. Our analysis indicates that these changes are probably due to a burying of the probes at these sites in the rhodopsin-arrestin or phospholipid-arrestin interface. Using a fluorescence approach based on this observation, we demonstrate that arrestin and retinal release are linked and are described by similar activation energies. However, at physiological temperatures, we find that arrestin slows the rate of retinal release ∼2-fold and abolishes the pH dependence of retinal release. Using fluorescence, EPR, and biochemical approaches, we also find intriguing evidence that arrestin binds to a post-Meta II photodecay product, possibly Meta III. We speculate that arrestin regulates levels of free retinal in the rod cell to help limit the formation of damaging oxidative retinal adducts. Such adducts may contribute to diseases like atrophic age-related macular degeneration (AMD). Thus, arrestin may serve to both attenuate rhodopsin signaling and protect the cell from excessive retinal levels under bright light conditions. The visual photoreceptor rhodopsin is perhaps the best model system for understanding the mechanisms used in G-protein-coupled receptor (GPCR) 1The abbreviations used are: GPCR, G-protein-coupled receptor; Rho*, light-activated rhodopsin; Rho-P, phosphorylated rhodopsin; Meta II, metarhodopsin II; Meta III, metarhodopsin III; ROS, rod outer segment, or wild-type rhodopsin in native membranes; ROS*, light-activated rhodopsin in native membranes; ROS-P, phosphorylated rhodopsin in native membranes; PIPES, 1,4-piperazinediethanesulfonic acid; MES, 2-(N-morpholino)ethanesulfonic acid; WT, wild type; RDH, retinal dehydrogenase. signaling, as detailed information exists on the structures and dynamic interactions of the protein constituents (1Ridge K.D. Abdulaev N.G. Sousa M. Palczewski K. Trends Biochem. Sci. 2003; 28: 479-487Abstract Full Text Full Text PDF PubMed Scopus (148) Google Scholar). Visual activation begins with absorption of light by the 11-cis-retinal chromophore in rhodopsin. The photoactivated form of rhodopsin, Rho* or “Meta II,” interacts with and activates the G-protein transducin, which exchanges nucleotide and then diffuses to interact with downstream effectors. Signaling by Rho* is terminated by slow thermal decay and the release of retinal. Alternatively, signaling can be quickly terminated by a process that begins with phosphorylation of rhodopsin's C-terminal tail through the action of rhodopsin kinase (2Kuhn H. Biochemistry. 1978; 17: 4389-4395Crossref PubMed Scopus (220) Google Scholar, 3Maeda T. Imanishi Y. Palczewski K. Prog. Retin. Eye Res. 2003; 22: 417-434Crossref PubMed Scopus (125) Google Scholar). The phosphorylated Rho* is then bound by arrestin, which stops signaling by physically occluding the G-protein binding site (4McBee J.K. Palczewski K. Baehr W. Pepperberg D.R. Prog. Retin. Eye Res. 2001; 20: 469-529Crossref PubMed Scopus (316) Google Scholar, 5Gurevich V.V. Gurevich E.V. Trends Pharmacol. Sci. 2004; 25: 105-111Abstract Full Text Full Text PDF PubMed Scopus (287) Google Scholar). In the present study, we address how these two inactivation mechanisms are related and, specifically, what governs arrestin release from rhodopsin. Arrestin is known to bind to phosphorylated Meta II, a form in which the photolyzed chromophore all-trans-retinal is still attached to the receptor by a deprotonated Schiff base. Arrestin does not bind phosphorylated opsin, but all-trans retinal added exogenously can stimulate arrestin binding to phosphorylated opsin (6Sachs K. Maretzki D. Hofmann K.P. Methods Enzymol. 2000; 315: 238-251Crossref PubMed Google Scholar, 7Hofmann K.P. Pulvermuller A. Buczylko J. Van Hooser P. Palczewski K. J. Biol. Chem. 1992; 267: 15701-15706Abstract Full Text PDF PubMed Google Scholar). Early studies showed indirectly that retinal release and arrestin release are probably interrelated events (7Hofmann K.P. Pulvermuller A. Buczylko J. Van Hooser P. Palczewski K. J. Biol. Chem. 1992; 267: 15701-15706Abstract Full Text PDF PubMed Google Scholar, 8Palczewski K. Jager S. Buczylko J. Crouch R.K. Bredberg D.L. Hofmann K.P. Asson-Batres M.A. Saari J.C. Biochemistry. 1994; 33: 13741-13750Crossref PubMed Scopus (133) Google Scholar). However, how these processes are linked or whether arrestin binds other photointermediates of rhodopsin (such as the storage form Meta III) is still unknown. In this paper, we demonstrate that arrestin binding and release can be directly observed in real time by monitoring spectral changes in fluorescently labeled arrestin mutants. Our studies employed two different cysteine mutants of arrestin, I72C and S251C, two sites that lie within the experimentally proposed rhodopsin-binding surface on arrestin (9Smith W.C. Dinculescu A. Peterson J.J. McDowell J.H. Mol. Vis. 2004; 10: 392-398PubMed Google Scholar, 62Pulvermuller A. Schroder K. Fischer T. Hofmann K.P. J. Biol. Chem. 2000; 275: 37679-37685Abstract Full Text Full Text PDF PubMed Scopus (69) Google Scholar, 63Dinculescu A. McDowell J.H. Amici S.A. Dugger D.R. Richards N. Hargrave P.A. Smith W.C. J. Biol. Chem. 2002; 277: 11703-11708Abstract Full Text Full Text PDF PubMed Scopus (40) Google Scholar, 64Vishnivetskiy S.A. Hosey M.M. Benovic J.L. Gurevich V.V. J. Biol. Chem. 2004; 279: 1262-1268Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). When these mutants are labeled with the fluorescent probe monobromobimane and incubated with phosphorylated rhodopsin, they exhibit changes in fluorescence upon rhodopsin photoactivation. Importantly, these changes do not occur in the presence of nonphosphorylated rhodopsin, and subsequent analysis suggests that the spectral changes are a direct result of interaction with activated, phosphorylated rhodopsin. In the present study, we demonstrate how these spectral changes can be exploited to obtain information on the rate of arrestin release. Our results provide direct evidence that arrestin and retinal release are indeed linked events and that arrestin slows the rate of retinal release from Rho*-P by ∼2-fold, possibly through a kinetic trap mechanism. In addition, we find that arrestin binding eliminates the pH dependence of retinal release. Surprisingly, we find that a subfraction of arrestin consistently remains bound to phosphorylated rhodopsin long after the decay of the active Meta II species. This latter result provides compelling evidence that arrestin binds Meta III or some other photodecay product. Materials—Frozen bovine retinas were obtained from Lawson and Lawson, Inc. (Lincoln, NE). GBX red light filters were purchased from Eastman Kodak Co., and [γ-32P]ATP was from PerkinElmer Life Sciences. Nitrocellulose filters (0.45 μm) and Biomax centrifugal concentrators (10-kDa cut-off) were from Millipore Corp. (Bedford, MA). Monobromobimane was obtained from Molecular Probes, Inc. (Eugene, OR), and [1-oxyl-2,2,5,5-tetramethyl-d-pyrroline-3-methyl]methanethiosulfonate spin label was from Toronto Research Chemicals (North York, Canada). 11-cis-Retinal was a generous gift from Rosalie Crouch (Medical University of South Carolina and NEI, National Institutes of Health). Cuvettes were purchased from Uvonics (Plainview, NY), and round capillaries for EPR measurements were from VitroCom, Inc. (Mountain Lakes, NJ). Band pass filters and long pass filters were acquired from Oriel (Stratford, CT). Acrylamide/bisacrylamine solution (37:5:1) and microcolumns were purchased from Bio-Rad. Spectroscopic grade buffers were from USB Corp. All other chemicals and reagents were obtained from Sigma. Preparation of Rod Outer Segments (ROS)—ROS were isolated from bovine retinas as described previously (10Papermaster D.S. Methods Enzymol. 1982; 81: 48-52Crossref PubMed Scopus (256) Google Scholar). All sucrose solutions were made in ROS buffer (70 mm potassium phosphate, 1 mm magnesium acetate, pH 6.8), and all procedures were done at 4 °C under red lights. Rhodopsin concentration was assessed by difference spectra in the presence of hydroxylamine (ϵ500 = 40,800 liters cm-1 mol-1). Stocks were snap-frozen and stored at -80 °C. Highly phosphorylated ROS (ROS-P) were prepared by suspending ROS (10 μm final rhodopsin concentration) in ROS buffer, with 20 μm GTP and 3 mm ATP (10-ml final volume). Phosphorylation of rhodopsin by the native rhodopsin kinase was initiated by illumination with a 15-watt bulb from a Kodak safelight (without filter) placed ∼20 cm away, and sedimentation of the membranes was prevented by gently rocking the sample. After 2 h, the reaction was stopped by a 4-fold dilution with ROS buffer (plus 50 mm hydroxylamine and 2% bovine serum albumin). Phosphorylated opsin membranes were then collected by centrifugation (40,000 × g, 50 min). Levels of phosphorylation were quantified with the use of [γ-32P]ATP (10–100 cpm/pmol) as a tracer. Aliquots were removed from the tracer reaction, spotted onto nitrocellulose filters, washed, and subjected to scintillation counting. Our assay indicated that ROS-P with ∼6.4 ± 1.5 phosphates/rhodopsin was created (11Kuhn H. Wilden U. Methods Enzymol. 1982; 81: 489-496Crossref PubMed Scopus (26) Google Scholar). Phosphorylated opsin membranes were washed by resuspending the pellet using a tissue homogenizer, followed by centrifugation. Membranes were washed twice with low ionic strength buffer (5 mm PIPES, 1 mm EDTA, pH 7.0) to remove peripheral proteins (12Kuhn H. Methods Enzymol. 1982; 81: 556-564Crossref PubMed Scopus (62) Google Scholar), followed by three additional washes with ROS buffer. The final washed opsin membranes were regenerated overnight by the addition of a 2-fold excess of 11-cis-retinal (4 °C). The regenerated samples were washed once with ROS buffer containing 2% bovine serum albumin and 50 mm hydroxylamine, twice with 2% bovine serum albumin in ROS buffer, and three times with ROS buffer alone. Nonphosphorylated ROS membranes were prepared identically, except that no ATP was added during the phosphorylation procedure. The turbidity of ROS samples was reduced by continuous sonication using a Branson 1210 bath sonicator (4 °C, 3 min). More than 95% of the rhodopsin retained function after sonication, as assessed by absorption spectroscopy. Synthesis of Synthetic Phosphopeptide 7PP—The 19-amino acid-long peptide analogous to the fully phosphorylated C-terminal tail of rhodopsin was synthesized and purified as described previously (13Puig J. Arendt A. Tomson F.L. Abdulaeva G. Miller R. Hargrave P.A. McDowell J.H. FEBS Lett. 1995; 362: 185-188Crossref PubMed Scopus (71) Google Scholar). Construction, Expression, and Purification of Arrestin—Recombinant bovine visual arrestin with an N-terminal His tag was expressed and purified from Pichia pastoris as described previously (14McDowell J.H. Smith W.C. Miller R.L. Popp M.P. Arendt A. Abdulaeva G. Hargrave P.A. Biochemistry. 1999; 38: 6119-6125Crossref PubMed Scopus (34) Google Scholar). Mutant constructs I72C and S251C were created utilizing PCR, and the constructs were confirmed by DNA sequencing on both strands. Labeling of Arrestin—Arrestin samples were buffer-exchanged and concentrated (∼50 μm) in labeling buffer (10 mm MES, 150 mm NaCl, pH 6.5) by ultrafiltration (Millipore Biomax). Monobromobimane was added from a stock in Me2SO in 10-fold molar excess to arrestin (final Me2SO concentration below 1%). After an incubation of 3 h at room temperature with gentle agitation, samples were centrifuged briefly at 100,000 × g to remove aggregates. Labeled arrestin was bound by multiple passages over His-select resin (Sigma) equilibrated with buffer (10 mm HEPES, 250 mm NaCl, pH 7.4), followed by extensive washing with buffer. Arrestin was eluted with 500 mm imidazole, and the imidazole was removed by size exclusion chromatography (Sephadex G-15; Sigma; 10 mm HEPES, 150 mm NaCl, pH 7.4). The labeling efficiency was calculated using ϵ280 = 26,360 liters cm-1 mol-1 for arrestin and ϵ393 = 5,000 liters cm-1 mol-1 for bimanep (15Mansoor S.E. McHaourab H.S. Farrens D.L. Biochemistry. 1999; 38: 16383-16393Crossref PubMed Scopus (53) Google Scholar, 16Schubert C. Hirsch J.A. Gurevich V.V. Engelman D.M. Sigler P.B. Fleming K.G. J. Biol. Chem. 1999; 274: 21186-21190Abstract Full Text Full Text PDF PubMed Scopus (71) Google Scholar). The absorbance at 392 nm was subtracted from the 280-nm value to compensate for bimane's absorbance at 280 nm. Using this method, labeling efficiencies of ∼83 and 88% were determined for arrestin mutants I72C and S251C, respectively. Since both of these mutants contain the three native cysteines of arrestin (Cys63, Cys128, and Cys143), a sample of wild-type (WT) arrestin was labeled using the same conditions as a control. The WT cysteines labeled at less than ∼2% efficiency. To assess possible free label contamination, the fluorescence of 2 μm labeled arrestin was compared with an identical sample, which had been precipitated with trichloroacetic acid (10%). For both I72B and S251B, free label contamination was well below 1%. Functional Pull-down Assay—A simple centrifugation assay was used to assess arrestin functionality. Briefly, sonicated ROS containing 12 μm rhodopsin or phosphorylated rhodopsin and 3 μm arrestin were mixed in 10 mm HEPES, 150 mm NaCl, pH 7.4, in the dark at room temperature (20 μl). Reactions were either kept in the dark or bleached for 5 min using a 150-watt fiber optic light source (>495 nm), followed by 10-fold dilution with ice-cold buffer and centrifugation at 100,000 × g for 10 min at 4 °C. The pellets were solubilized in loading buffer and subjected to SDS-PAGE (10%). Proteins were visualized by Coomassie staining, and densitometry was performed using AlphaEase FC software. To assess arrestin affinity for post-Meta II rhodopsin, ROS-P in 10 mm HEPES, 150 mm NaCl, pH 7.0, was photobleached at room temperature for 90 s and then transferred to a 35 °C water bath. The sample was kept in the dark after the initial photobleach. After 12 min, arrestin I72B was added (1 μm) and incubated at room temperature for an additional 3 min. The samples were diluted, centrifuged, and subjected to SDS-PAGE as described above. The fluorescently labeled arrestin was visualized with a gel-doc apparatus (Alpha-Innotech FluoroChem 5500). The gel was excited from above with a short wave UV source, and the fluorescent bands were detected through a cut-off filter (535 ± 50 nm) by a CCD camera (5-min exposure). Densitometry was performed using AlphaEase FC software. Steady-state Fluorescence Assays—All fluorescence measurements were made using a Photon Technologies QM-1 steady-state fluorescence spectrophotometer with a single excitation source and two emission detectors (T format). Temperature was controlled and monitored using a water-jacketed cuvette holder connected to a circulating water bath (VWR Scientific) and a digital thermometer, which was submerged into a water-filled well in the sample chamber. Typically, 100-μl samples containing 1 μm labeled arrestin in a 2-mm black-jacketed cuvette were excited at 380 nm, and the emitted fluorescence was measured from 400 to 600 nm using 2-nm increments. Each data point was integrated for 0.25 s, and the average of two scans yielded the final spectrum. Excitation bandpass was kept at 0.25 nm to avoid bleaching of rhodopsin samples (emission bandpass at 15 nm). Spectra were smoothed and analyzed using the PTI software program Felix, and the fluorescence spectra of buffer or rhodopsin alone were subtracted where appropriate. Rhodopsin was generally present at a 4-fold excess to arrestin, which was found to be sufficient for complete arrestin binding. The samples were bleached using a 150-watt fiber optic light source (>495 nm) for 40 s. Fluorescence Lifetime Measurements—A PTI Laserstrobe fluorescence lifetime instrument was used to measure I72B and S251B under different conditions. Typically, 500 nm arrestin in a 4-mm black-jacketed cuvette (200 μl) was measured at 20 °C using 381-nm excitation pulses (full-width half-maximum, ∼1.5 ns) with a 298–435-nm band pass filter on the excitation beam. The emission was monitored through two >470-nm long pass filters, and neutral density filters were used to modulate the intensity. Each data point collected represented two averages of five laser shots, and typically 150 points were collected over the lifetime decay curve. The instrument response function, which must be deconvoluted from lifetime decay data, was determined from the scatter from a solution of Ludox through a 400-nm broadband interference filter. Data points were acquired randomly to minimize the impact of laser misfires on the decay curve, and data were analyzed using the commercial PTI software TimeMaster. The “goodness of fit” was evaluated by plotting the residuals and the χ2 value (0.7 < χ2 < 1.2 was considered acceptable) (17Straume M. Johnson M.L. Methods Enzymol. 1992; 210: 87-105Crossref PubMed Scopus (110) Google Scholar, 18Lakowicz J.R. Principles of Fluorescence Spectroscopy. 2nd Ed. Kluwer Academic, New York1999Crossref Google Scholar). Fluorescent lifetimes of these labeled arrestins were also measured in the presence of the phosphopeptide 7PP (100 μm) or ROS-P (2 μm in the dark state, after photoactivation, and after the addition of 50 mm hydroxylamine). The lifetime data acquisition scheme described above was found to bleach <2% of rhodopsin, and the total time elapsed from the start of photoactivation to the conclusion of the measurement was 4 min, or ∼0.5 Meta II decay half-lives at 20 °C. ROS-P was found to introduce a significant amount of scatter and a fluorescent component with an extremely short lifetime (<1 ns) into the samples. To correct for these components, they were simply subtracted from the arrestin life-time by measuring the lifetime of ROS-P alone with matched concentration under the same conditions. Fluorescence Quenching Analysis—To measure the quenching effects of I- on free versus rhodopsin-bound arrestin, a stock of 4 μm ROS-P and 1 μm labeled arrestin was divided among five tubes (0, 10, 20, 30, or 50 mm KI). KCl was added to keep the ionic strength consistent, and 0.1 mm sodium thiosulfate was present to suppress I3− formation. The steady-state fluorescence of each sample was measured in the dark state and immediately after photoactivation, and the average of two independent experiments was used to calculate quenching constants. KSV is derived from the Stern-Volmer equation F0/F = 1 + KSV[Q], where F0 and F represent the fluorescence intensities in the absence and presence of quencher, respectively, and [Q] is the concentration of quencher. KSV values were also determined for free arrestin and arrestin in the presence of the phosphopeptide 7PP by titration with KI. The fluorescence was corrected for dilution, and an independent titration with KCl was performed to assess any potential ionic effects on the fluorescence. For time-based quenching studies of arrestin I72B, the average of a range of fluorescence values (λem = 456 nm, 35 °C) was used to derive the Stern-Volmer constants in the dark (300 s prior to activation), after photoactivation (first four points collected after activation), and at the plateau (1200–1500 s after activation). Time-based Fluorescence Assays—To measure arrestin I72B binding, the samples were excited continuously at 380 nm, and the fluorescence emission at 456 nm was monitored (two points/s). The excitation band-pass settings were kept below 0.2 nm, resulting in less than 1% rhodopsin bleaching over a 10-min period. Arrestin binding was measured at 8 °C (1 μm arrestin and 8 μm ROS-P, 55 μl) after irradiation from a Machine Vision Strobe light source for 5 s, which photoactivated 56% of the rhodopsin. Buffer-subtracted data were fit using the program Sigma Plot (SPSS Inc., Chicago, IL) to a monoexponential equation to define the rate of arrestin binding. Arrestin release was measured in a similar way, except that the excitation shutter was opened for 2 s and then closed for 8 s between measurements to avoid photobleaching. When used, 2 μl of a 300 mm hydroxylamine stock (pH 7) was added, and this was found not to have any effect on the fluorescence of labeled arrestin. Simultaneous monitoring of retinal and arrestin release from ROS-P was accomplished by exciting the sample at 295 nm for 2 s, followed by a 2-s excitation at 380 nm. Fluorescence emission at 330 and 456 nm was monitored (1 point/s), and the shutter was closed for 8 s between measurements. The sample was irradiated for 8 s with a Machine Vision Strobe light source, which photobleached >90% of the rhodopsin. In this way, the change in rhodopsin tryptophan emission due to retinal release (λex = 295 nm; λem = 330 nm) (19Farrens D.L. Khorana H.G. J. Biol. Chem. 1995; 270: 5073-5076Abstract Full Text Full Text PDF PubMed Scopus (175) Google Scholar) and the change in bimane fluorescence due to arrestin I72B binding and release (λex = 380 nm; λem = 456 nm) could be monitored simultaneously. The fluorescence contribution from arrestin's one native tryptophan was subtracted from retinal release data, since the fluorescence of this tryptophan does not change upon arrestin activation (20Wilson C.J. Copeland R.A. J. Protein Chem. 1997; 16: 755-763Crossref PubMed Scopus (23) Google Scholar). Retinal release from ROS-P without arrestin was measured in the same way, except that no arrestin was present. Data were background-subtracted and fit to either monoexponential rise or decay equations using the program Sigma Plot to derive rates. Goodness of fit was evaluated by plotting the residuals. Arrestin release rates and retinal release rates, with or without arrestin, were determined at 15, 20, 25, 30, 35, and 40 °C (pH 7.5), and the average of two independent experiments was used to derive the activation energy (Ea) of these events using the Arrhenius equation, k = Ae-Ea/(RT). The pH dependence of retinal and arrestin release was determined by mixing the phosphorylated rhodopsin and arrestin into buffer (20 mm HEPES, 150 mm NaCl) at pH 6.0, 6.5, 7.0, 7.5, 8.0, or 8.5. Rates were determined as described above (35 °C). EPR—Arrestin I72C and S251C were prepared for EPR in an identical manner as described for monobromobimane labeling, except that a 5-fold molar excess of nitroxide spin label was used during labeling. The EPR spectrum of a sample of WT arrestin, labeled using identical conditions, showed less than 10% incorporation of spin label at the native cysteines compared with I72C. For EPR measurements, a 6-μl volume of 50 μm spin-labeled arrestin and 200 μm ROS-P were measured in the dark at room temperature (19–21 °C) and at various time points following photoactivation (45 s using a 150-watt >495-nm fiber optic light source). To assess whether the observed spectral changes were due to changes in protein rotational rates, EPR spectra were also measured in the presence of 20% Ficoll 400 (21Frazier A.A. Wisner M.A. Malmberg N.J. Victor K.G. Fanucci G.E. Nalefski E.A. Falke J.J. Cafiso D.S. Biochemistry. 2002; 41: 6282-6292Crossref PubMed Scopus (100) Google Scholar). Ficoll concentrations higher than 20% could not be used, since they caused arrestin precipitation. EPR measurements were made using a Varian E-104 instrument fitted with a loop-gap resonator and the EWWIN 5.22 data acquisition package (Scientific Software Services, Plymouth, MI). Measurements were carried out at ∼9.3-GHz microwave frequency, using 2-milliwatt incident microwave power, a modulation amplitude of ∼2 gauss, and a 100-gauss sweep (29 s/scan). Multiple scans were taken and averaged where appropriate (see the legend to Fig. 7 for details). Numerous studies suggest the two concave surfaces of arrestin are involved in binding light-activated phosphorylated rhodopsin (5Gurevich V.V. Gurevich E.V. Trends Pharmacol. Sci. 2004; 25: 105-111Abstract Full Text Full Text PDF PubMed Scopus (287) Google Scholar, 9Smith W.C. Dinculescu A. Peterson J.J. McDowell J.H. Mol. Vis. 2004; 10: 392-398PubMed Google Scholar, 22Ling Y. Ascano M. Robinson P. Gregurick S.K. Biophys. J. 2004; 86: 2445-2454Abstract Full Text Full Text PDF PubMed Scopus (7) Google Scholar). In the present study, we introduced two cysteine residues between these surfaces and labeled them with the cysteine-specific fluorescent probe monobromobimane (Fig. 1). We and others have extensively characterized bimane and found that it can reliably report on protein dynamics and structure due to its small size, its sensitivity to polarity, and its ability to be quenched by nearby tryptophan and tyrosine residues (15Mansoor S.E. McHaourab H.S. Farrens D.L. Biochemistry. 1999; 38: 16383-16393Crossref PubMed Scopus (53) Google Scholar, 23Giniger R. Huppert D. Kosower E.M. Chem. Phys. Lett. 1985; 118: 240-245Crossref Scopus (18) Google Scholar, 24Kosower E.M. Giniger R. Radkowsky A. Hebel D. Shusterman A. J. Phys. Chem. 1986; 90: 5552-5557Crossref Scopus (40) Google Scholar, 25Mansoor S.E. Farrens D.L. Biochemistry. 2004; 43: 9426-9438Crossref PubMed Scopus (50) Google Scholar, 26Mansoor S.E. McHaourab H.S. Farrens D.L. Biochemistry. 2002; 41: 2475-2484Crossref PubMed Scopus (87) Google Scholar). Below, we report our studies on the bimane-labeled arrestin mutants I72C and S251C. Fluorescently Labeled Arrestin Mutants Are Functional— The relative functionality of the mutants with and without the bimane label was first assessed using a centrifugal “pull-down” assay (Fig. 2). Mutant I72C shows proper binding specificity to ROS*-P but binds ROS*-P with less affinity (43% of WT). Interestingly, bimane labeling of I72C (I72B) appears to restore the binding ability of this mutant to ∼99% of WT levels. Both unlabeled arrestin S251C and labeled S251B bind to similar levels as WT (89 and 101%, respectively). Spectral Properties of Labeled Arrestin Mutants—Both I72B and S251B fluoresce with a λmaxof ∼470 nm, and their spectra do not change in the presence of an excess of dark ROS-P. However, upon photoactivation, the emission spectrum of arrestin I72B blue-shifts to a λmax of 456 nm and increases in total integrated intensity by ∼12% (Fig. 3A). The fluorescence of arrestin S251B increases ∼100% upon photoactivation but displays no shift in its λmax (Fig. 3D). Importantly, these changes are not observed using nonphosphorylated ROS (Fig. 3, B and E) and are abolished by 50 mm hydroxylamine (data not shown), presumably because hydroxylamine catalyzes the decay of activated rhodopsin by cleaving the retinal Schiff base. Spectral Changes Require Interaction with Rhodopsin—We next assessed whether the observed spectral changes were due to conformational changes within arrestin itself rather than interactions with ROS*-P. To do this, we tested the effect of phosphopeptide 7PP, which represents the fully phosphorylated form of rhodopsin's C-terminal tail and has been shown to transactivate arrestin to bind nonphosphorylated ROS* (13Puig J. Arendt A. Tomson F.L. Abdulaeva G. Miller R. Hargrave P.A. McDowell J.H. FEBS Lett. 1995; 362: 185-188Crossref PubMed Scopus (71) Google Scholar). The peptide 7PP causes an ∼20% decrease in intensity for mutant I72B (Fig. 3C) and induces no change in arrestin S251B (Fig. 3F). We obtained similar results as for 7PP using heparin and phytic acid, two polyanionic compounds that have been reported to bind arrestin and induce activating conformational changes (data not shown) (20Wilson C.J. Copeland R.A. J. Protein Chem. 1997; 16: 755-763Crossref PubMed Scopus (23) Google Scholar, 27Palczewski K. Pulvermuller A. Buczylko J. Hofmann K.P. J. Biol. Chem. 1991; 266: 18649-18654Abstract Full Text PDF PubMed Google Scholar). Fluorescence Lifetime Analysis—To further elucidate the cause of these spectral changes, we measured the fluorescent decay lifetimes of I72B and S251B under different conditions (Table I). The fluorescence lifetime (τ) of I72B is 11.1 ns, and this value is shortened slightly by the phosphopeptide 7PP (10.6 ns). In the presence of ROS-P, the τ shortens by ∼1 ns after photoactivation but reverts to ∼10.5 ns after the addition of hydroxylamine. These values are all similar to free bimane (9.1 ± 0.1 ns; data not shown), s" @default.
• W2076770170 created "2016-06-24" @default.
• W2076770170 creator A5000618696 @default.
• W2076770170 creator A5043616114 @default.
• W2076770170 creator A5076637852 @default.
• W2076770170 date "2005-02-01" @default.
• W2076770170 modified "2023-09-27" @default.
• W2076770170 title "Dynamics of Arrestin-Rhodopsin Interactions" @default.
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