Single-cell recording

Salamanders were dark adapted for a minimum of 15 h, to ensure complete regeneration of rhodopsin in green-sensitive (GS) rods (Kefalov et al., 2005). Retinal tissue was isolated under infrared illumination, whereupon a small sample was mechanically dissociated and perfused continuously in the recording chamber with Ringer’s solution consisting of: 108 mM NaCl, 2.5 mM KCl, 1.0 mM MgCl₂, 1.5 mM CaCl₂, 0.02 mM EDTA, 10 mM glucose, 0.8 µM bovine serum albumin (BSA, essentially fatty acid free; Sigma, Saint Louis, MO), 10 mM HEPES; pH 7.6, that was bubbled with 95% O₂/5% CO₂ at room temperature (20–22°C). Photocurrents were recorded from the inner segment of a single rod or cone with a suction electrode using a current-to-voltage converter (Axopatch 200A; Axon Instruments, Foster City, CA), low-pass filtered at 30 Hz with an eight-pole Bessel filter (−3 dB; Frequency Devices, Haverhill, MA), and digitized at 400 Hz. No corrections were made for the delay introduced by filtering. Cells were stimulated with light from a 75 W xenon arc lamp that passed through a six-cavity interference filter (10-nm bandwidth at half-maximal transmission; Omega Optical, Brattleboro, VT), quartz neutral density filters (Omega Optical), and an electronic shutter. A nominal duration of 22 ms was used for test flashes. The rod or cone spectral type was determined by inspection of cell shape and by responses to flashes at 434, 500, and 626 nm. The light was calibrated with a photometer (UDT 350; Graseby, Orlando, FL) through a 200-µm–diameter pinhole (Melles Griot, Carlsbad, CA) placed at the level of the recording chamber. To compare spectral sensitivity by single-cell recording and absorption spectrum by MSP, intensity was measured through a linear polarizer that separated two components whose electric vectors were orthogonal and parallel to the long axis of the recorded rod outer segment.

Vitamin A (all-trans retinol) and vitamin E ((+)-α-tocopherol) were purchased from Sigma. Stocks were made in absolute ethanol, protected from light, and stored at −80°C until use. Prior to an experiment, the purities and concentrations were checked from absorption spectra measured with a Beckman DU-640 ultraviolet (UV)-visible spectrophotometer using extinction coefficients: ε_vitA = 52,800/M/cm at 325 nm for vitamin A and ε_vitE = 3260/M/cm at 292 nm for vitamin E (Robeson et al., 1955; Podda et al., 1996). The vitamin A stock was diluted first with ethanol and then with Ringer’s solution containing BSA to obtain the desired concentration. In preliminary experiments, instability of vitamin A (Fig. S1A) was problematic because the time course of degradation was variable and because the degradation product proved to be more potent than vitamin A (Fig. S2). Addition of vitamin E effectively stabilized the vitamin A for many hours (Fig. S1B) as previously reported (Crouch et al., 1992). The final concentrations of vitamin E and BSA were 10 times lower than that of vitamin A and the final concentration of ethanol was ≤0.1%. In control experiments, perfusion with 0.1% ethanol, 6 µM BSA, or vitamin E at concentrations up to 10 µM had no effect on the flash responses of rods. Vitamin A was introduced to the cell through one channel of a theta tube and Ringer’s containing 6 µM BSA and 0.1% ethanol ran through the other channel to wash out vitamin A. A waste port positioned across from the theta tube established laminar flow from the two channels with little interfacial mixing. The theta tube was gravity fed through delivery tubes from two glass syringe reservoirs chilled with freezer packs. The delivery tubes were not chilled so the solutions warmed to room temperature before entering the experimental chamber. Because vitamin A was hydrophobic and adsorbed to the walls of the syringe and the delivery tube, the actual concentration in the solution exiting the theta tube was determined spectrophotometrically at the end of each experiment.

The stimulus-response relation was fit with the Hill equation:

where r is the response amplitude, r_max is the saturating response amplitude, i is flash strength, H is the Hill coefficient, and i_0.5 is the flash strength required for a response of half-maximal amplitude.

Spectral sensitivities were determined (Naka & Rushton, 1966; Baylor & Hodgkin, 1973) before and during treatment and after washout. Relative sensitivity was found by comparing the flash strength required for a criterion, dim flash response at each test wavelength to that at the reference wavelength, 500 nm.

Single-cell MSP

Under infrared illumination, cells were dissociated mechanically on a quartz coverslip that had been coated with poly-l-lysine to make the cells adhere. Double-sided tape was placed lengthwise on opposite edges of the coverslip and a shorter coverslip was placed on top. About 50 µl of Ringer’s solution containing small pieces of retina and single cells could be stabilized between the coverslips. This chamber was placed on the stage of the Williams–Webbers microspectrophotometer (Makino et al., 1990).

A suitable rod was located with a closed-circuit infrared television system. Baseline measurements were taken with the probe beam passing through a cell-free area. The rod was then positioned in the path of the probe beam. The probe beam was polarized with a Nicol prism. An OD measurement will be referred to as “perpendicular” (⊥) when the electric vector was oriented orthogonal to the long axis of the outer segment and “parallel” (∥) when the rod was rotated 90 degrees. Measurements were made at room temperature (21–23°C) at 2-nm intervals over the range 250–700 nm and absorbance was calculated online.

Vitamin A in Ringer’s was delivered to the cells by placing a drop on one open side of the chamber. The fluid was drawn through the chamber by touching absorbent paper to the opposite side. The chamber was perfused one to three times in rapid succession after periods ranging from 10 to 102 min. The frequency of perfusions did not appear to affect the rate of uptake. To account for adsorption losses associated with vitamin A delivery, the concentration passing through the chamber was measured spectrophotometrically in control experiments. Absorption spectra were measured from the same cells before and during treatment with vitamin A whenever possible. In addition, absorbance was measured after washing rods with Ringer’s containing 6 µM BSA, to simulate conditions used in electrophysiological experiments.

The molar ratio of vitamin A to rhodopsin was determined for individual rods in order to chart the time course of uptake. The average OD_∥ spectrum of untreated rods was scaled to match the OD_∥ spectrum of a given rod being treated with vitamin A between 480 and 600 nm. The difference spectrum was taken as the absorbance due to vitamin A and fitted with a Gaussian. The range of the fit was typically 300–400 nm, although the short wavelength limit was sometimes varied to avoid intrusion from imperfect subtraction of the protein absorption band. The OD_⊥ spectrum of a treated rod was used to determine rhodopsin content. If an OD_⊥ spectrum was not available, the dichroic ratio was found for untreated rods as OD_⊥/OD_∥ at the absorbance maximum near 500 nm. The OD_∥ spectrum for the treated rod was then multiplied by this dichroic ratio, and the spectrum fit over the range 460–650 nm with the sum of two template functions for 11-cis retinal (A1) and 11-cis dehydroretinal (A2) components of visual pigment (Govardovskii et al., 2000) with maxima at 501 and 519 nm, respectively. The vitamin A to rhodopsin ratio was then calculated as:

where OD refers to the optical density of vitamin A, A1 visual pigment or A2 visual pigment, and ε is the corresponding molar extinction coefficient: 52,800/M/cm for vitamin A, 42,000/M/cm for A1 visual pigment (Matthews et al., 1963), and 30,000/M/cm for A2 visual pigment (Brown et al., 1963).

Vitamin A binding assay

Rod outer segments were isolated from frozen bovine retinas (W.L. Lawson, Lincoln, NE) under infrared illumination using a stepwise sucrose density gradient (McDowell, 1993). Rhodopsin was purified according to Litman (1982) with slight modifications. The outer segments were suspended in solubilizing buffer consisting of: 0.1% (w/v) sucrose monolaurate (SM) (Fluka, Neu-Ulm, Germany), 0.5 mM MgCl₂, 140 mM NaCl, 1 mM dithiothreitol, 10 mM HEPES; pH 7.5, and were stirred at 4°C for 1 h. The supernatant containing solubilized rhodopsin was loaded onto a concanavalin A sepharose 4B (Sigma) column equilibrated with the same buffer. The column was washed with the same buffer until unbound protein was no longer present in the collected fractions. The concentration of SM in the column was decreased gradually by first rinsing with the solubilizing buffer containing 0.05% SM and then with enough solubilizing buffer containing 0.02% SM to exchange the bed volume of the column 15 times. Bound rhodopsin was eluted with solubilizing buffer containing 0.5 m methyl α-d-mannopyranoside (Sigma) and 0.02% SM. Rhodopsin concentration was determined from the difference in absorbance at 500 nm before and after complete bleaching in the presence of 10 mM hydroxylamine. Rhodopsin purity was confirmed by an absorbance at 280 nm that was less than two times that at 500 nm (Irreverre et al., 1969).

For binding assays, samples contained 1 µM rhodopsin in solubilizing buffer containing 0.5 m methyl α-d-mannopyranoside and 0.02% SM. The requisite amounts of vitamin A in 1.5 µl of ethanol were added to 150 µl of sample to yield final concentrations varying from 0.23 to 120 µM. After incubating the sample at room temperature (21–23°C) for 10 min, the concentration of vitamin A dissolved in the solution was determined from its absorbance at 325 nm. For high concentrations of vitamin A, samples were diluted in order to measure absorbance. Control experiments were carried out with 1-µM recombinant human vascular cell adhesion molecule (VCAM-1) (R&D Systems, Minneapolis, MN) in the same buffer or with the buffer alone. An extinction coefficient of 52,800/M/cm for vitamin A was assumed to be applicable for all conditions.

Sensitization of rods to UV light by vitamin A

Some effects of vitamin A were wavelength dependent; during perfusion with 8–10 µM vitamin A, sensitivity to flashes declined at all visible wavelengths but there was little or no change at UV wavelengths (Fig. 3A, gray circles). After normalizing to match relative sensitivity between pretreatment and treatment conditions at the two longest wavelengths, vitamin A was found to increase relative sensitivity to UV light by 3.9-fold (Fig. 3B and 3C). Hence, some “GS” rods became maximally sensitive to ~330 nm. The change in relative sensitivity during treatment matched well to the relative absorption spectrum of vitamin A (Fig. 3C). Flash response kinetics were similar between UV and long wavelengths (Fig. 3D), consistent with the principle of univariance; once absorbed, UV and visible photons induced the same active state of rhodopsin. Vitamin A also enhanced the relative sensitivity to UV light in two BS rods and two RS cones (data not shown). It was suggested that in some insect photoreceptors, vitamin A absorbs UV light and then donates the energy to the retinal chromophore of rhodopsin by Förster resonance energy transfer (FRET) to make the photoreceptors more sensitive to UV light (reviewed in Vogt, 1989). Thus, our results suggested that vitamin A was also an accessory chromophore for FRET with vertebrate rod and cone visual pigments.

The concentration dependence of sensitization to UV light, determined after exposure to vitamin A for 30–60 min, is shown in Fig. 3E. But in three GS rods, there was a further increase in relative sensitivity to UV wavelengths after washing, indicating that the additional exposure to vitamin A between the initial measurement of UV sensitivity and washing allowed for a greater enhancement. Therefore, this dose–response relation represented a snapshot in time rather than an equilibrium condition because a maximal effect was not necessarily obtained for each applied concentration of vitamin A. As will be described below, uptake continued throughout the exposure to vitamin A. A notable point was that the effect of vitamin A on sensitivity saturated with 500 nm light but did not with UV light (compare Figs. 2D and ​and3E).3E). Subsequent washing of GS rods with Ringer’s containing BSA for 20–30 min increased uniformly the absolute sensitivity to all wavelengths (Fig. 3A, open triangles). Thus, the relative sensitivity to UV wavelengths remained elevated. In no case did UV sensitivity return to pretreatment values after washing for at least 30 min even though the sensitivity to other wavelengths returned completely (Figs. 2D and ​and3E).3E). The marked difference in the dose–response relations for sensitivity to UV and visible wavelengths and their reversibility of the effect on UV sensitivity implied that vitamin A affected sensitivity to the two wavelength domains by acting at different sites.

Uptake of vitamin A into the outer segments of rods

In order to quantify the amount of vitamin A in the outer segment and the time course of uptake, the absorbance of single GS rods was measured by MSP. In the outer segments of untreated rods, there was a prominent absorbance peak near 280 nm due to the tryptophans and tyrosines in protein, as well as a peak near 515 ± 2 nm (n = 9 salamanders, where the value for each salamander derived from measurements of 13–31 rods) arising from rhodopsin (e.g., Fig. 4A, thin black line). The alignment of rhodopsin molecules within the disk membranes of the outer segment caused absorbance at long wavelengths to be strongest when the long axis of the outer segment was perpendicular to the electric vector of the probe beam with a dichroic ratio, D = OD_⊥/OD_∥ = 3.00 ± 0.08 (n = 9 salamanders). The protein band was not dichroic, D = 0.94 ± 0.06 (n = 9 salamanders). These observations are in general agreement with absorbance measurements by others (Liebman, 1972; Hárosi, 1975; Cornwall et al., 1984).

Vitamin A readily partitioned into the membranes of the rod. Perfusion with vitamin A increased UV absorbance in the inner segments of rods (data not shown) as well as in their outer segments with a maximal change near 330 nm (Fig. 4A and 4B, gray and thick black lines). Since baseline measurements were taken in a cell-free area through the bathing solution containing vitamin A, the change in UV absorbance represented an accumulation of vitamin A on or within the cell. In contrast to rhodopsin, vitamin A absorbed light best when the electric vector of the measuring beam was parallel to the long axis of the outer segment, consistent with an alignment of the molecule perpendicular to the interior disk membrane surfaces. The dichroic ratio for vitamin A was approximately 0.50 ± 0.03 (n = 20 rods) after long treatments extending from 30 to 193 min. Endogenously formed vitamin A assumes a similar orientation within the outer segment after the bleaching of rhodopsin (Denton, 1959; Liebman, 1972). Although the behavior of individual rods was variable, the overall trend was for uptake to exhibit an initial rapid component, followed by a slower component that did not equilibrate even after perfusion for several hours (Fig. 4C and 4D). Both the initial level of uptake and the rate of uptake of the slow component increased with higher bathing concentrations of vitamin A. The initial uptake ranged from 0.05 to 0.2 vitamin A per rhodopsin (Fig. 4D, regression line). For applied vitamin A concentrations less than 10 µM, the slower rate of uptake did not increase greatly the vitamin A content in the outer segment over the treatment period during which measurements of flash response sensitivity and kinetics were made in the single-cell recordings (Fig. 4E). For higher applied concentrations of vitamin A, the rate of uptake was more rapid so measurements in single-cell recordings were limited to a 30- to 45-min treatment period (Fig. 4E, red line). Uptake seemed anomalously high in some rods treated with 6–10 µM or with 16–20 µM vitamin A, perhaps because the cell integrity was compromised during dissociation. Attempts to remove vitamin A from the outer segments of rods by perfusion with 6 µM BSA were unsuccessful when such high vitamin A concentrations were used (data not shown), which could explain the persistence of enhanced UV sensitivity after washing in single-cell recordings (Fig. 3).

Absorbance measurements by MSP were compared to spectral sensitivity measurements with suction electrode recording (Fig. 4F). For that purpose, the polarization of light stimulating the rod in single-cell recording was determined in order to generate a weighted mean between OD_⊥ and OD_∥ spectra. The spectral sensitivity matched this resultant absorption spectrum before and during perfusion with vitamin A, signifying that photic energy transfer from vitamin A to the 11-cis retinal chromophore was as effective in photoisomerizing rhodopsin as direct absorption of light by rhodopsin.

Characterization of vitamin A binding

There was some uncertainty in calculating the stoichiometry of binding based on measurements of absorbance because the extinction coefficient for vitamin A is sensitive to environmental factors. Values range from 17,800/M/cm in water to 46,000/M/cm for vitamin A bound to retinol-binding protein and to 52,800/M/cm in ethanol (Robeson et al., 1955; Horwitz & Heller, 1973; Szuts & Hárosi, 1991). We used the last value to estimate [vitamin A] in our solutions as well as in cells; a lower value would yield higher ratios of vitamin A to rhodopsin.

Rods bathed for 40 min in [vitamin A] = 16–20 µM amassed approximately one vitamin A molecule per two rhodopsins (Fig. 4E) and consequently became 2.6-fold less sensitive to visible light (Fig. 2D). An equivalent loss in sensitivity is produced by steady light that isomerizes approximately 60 rhodopsins/s (cf. Pugh & Lamb, 2000). Assuming that there are a few billion rhodopsins in the rod (e.g., Pugh & Lamb, 2000), half of which have a vitamin A bound and are catalytically active, then the integrated activity of one of these rhodopsins would be approximately 10 million times lower than that of a light-activated rhodopsin. This level of activity is comparable to that of apo-opsin (Jones et al., 1996). The saturation of sensitivity loss at [vitamin A] in which the vitamin A to rhodopsin ratio was less than one could be explained by a scenario, in which vitamin A only activated a certain subpopulation of rhodopsins: a certain conformation, a multimer, rhodopsins interacting with an accessory protein as is the case for some GPCRs (Christopoulos & Kenakin, 2002), or rhodopsins at a particular location. For example, allosteric binding of vitamin A on the extracellular surface of rhodopsins in the plasma membrane could explain why activation disappeared soon after washing despite a large content of vitamin A in the disk membranes and UV sensitization. Depending upon the relative size of the subpopulation, the specific activity of rhodopsin activated by vitamin A could be considerably higher.

The effects of extracellularly applied vitamin A on GS rods may be compared to those of other retinoids. β-Ionone and 9-cis and 11-cis retinals, at concentrations ~50 µM, inhibited the CNG channel but did not activate rhodopsin (Kefalov et al., 2005; He et al., 2006; Isayama et al., 2009). Four micromolar 11-cis vitamin A in liposomes desensitized GS rods by 10-fold (Jones et al., 1989), suggesting that the cis isomer might be a more powerful agonist than the trans isomer. Unexpectedly, a degradation product(s) of vitamin A at a concentration <10 µM produced the greatest activation of rhodopsin (Fig. 6; Fig. S2). The most active ingredient was not identified, but the spectral shift to shorter wavelengths (Fig. S1) was consistent with a loss in conjugation. These results suggested that a retinoid whose separation of β-ionone ring to terminal aldehyde or alcohol group was longer than that of β-ionone but shorter than that of all-trans vitamin A, could activate GS rod pigment most effectively. Candidates for the binding site include several of the proposed sites on GS rod opsin for retinal (reviewed in Heck et al., 2003) and a recently identified site for β-ionone on GS rod rhodopsin (Makino et al., 2010). The mechanism for rhodopsin activation has yet to be identified. Taking a more global view, many other GPCRs are subject to allosteric regulation at various sites by compounds that contain ring structures (reviewed in May et al., 2007; Yanamala & Klein-Seetharaman, 2010), so vitamin A might even be pharmacologically active in tissues that do not express opsins.

Mechanism underlying sensitization of rhodopsin to UV light by vitamin A

The enhanced relative UV sensitivity of photoreceptors upon treatment with vitamin A (Fig. 3) indicated that vitamin A absorbed UV light and then transferred the energy to the chromophore of rhodopsin. The high efficiency of energy transfer (Fig. 4) ruled out the trivial mechanism, whereby rhodopsin simply absorbed vitamin A fluorescence because fluorescence would have spread in all directions and most of the energy would have been lost. Instead, there must have been radiationless transfer. Such a mechanism could occur in two ways. Dexter type requires contact between donor and acceptor and involves collisional exchange of energy. On the other hand, FRET can occur over greater distances if the donor and acceptor are aligned and if there is spectral overlap between the emission of the donor and absorption by the acceptor. The critical Förster distance at which half the energy is transferred is typically 20–90 Å (Lakowicz, 2006). Rods contain huge amounts of rhodopsin with an average distance between centers of rhodopsin of 45 Å (Fotiadis et al., 2004). Thus, a vitamin A inserted in the middle of a unit cell of a triangular matrix of rhodopsins would be 26 Å from each of three rhodopsins. The broad emission of vitamin A at 480 nm (Fig. 5F) falls well within the main absorption band of rhodopsin. Although the orientation of the vitamin A with respect to the chromophore of rhodopsin is not optimal for FRET (Fig. 4A and 4B), the lengthened radiative lifetime of vitamin A in phospholipid or bound to protein (Radda & Smith, 1970) coupled with the large oscillator strength of rhodopsin and the extreme rapidity of its isomerization (Wang et al., 1994) may allow energy transfer to occur with high efficiency. If so, then vitamin A may sensitize vertebrate rhodopsins by FRET as was suggested for insect rhodopsins (Vogt, 1989).

We thank Dr. E.R. Makino for helpful advice regarding the in vitro assays, Drs. X.-H. Wen and T.R. Middendorf for insightful discussions, and Dr. A.L. Zimmerman for comments on the manuscript. This research was supported by Award Numbers {"type":"entrez-nucleotide","attrs":{"text":"EY011358","term_id":"159077629","term_text":"EY011358"}}EY011358 and {"type":"entrez-nucleotide","attrs":{"text":"EY014104","term_id":"159076760","term_text":"EY014104"}}EY014104 from the National Eye Institute and the Lions of Massachusetts. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Eye Institute or the National Institutes of Health.