The vitelliform macular dystrophy protein defines a new family of chloride channels

Abstract

Vitelliform macular dystrophy (VMD/Best disease; MIM*153700) is an early-onset autosomal dominant disorder in which accumulation of lipofuscin-like material within and beneath the retinal pigment epithelium is associated with a progressive loss of central vision. Bestrophin, the protein product of the VMD gene, has four predicted transmembrane domains. There are multiple bestrophin homologues in the human, Drosophila, and Caenorhabditis elegans genomes, but no function has previously been ascribed to these proteins, and they show no detectable homology to other proteins of known function. Using heterologous expression, we show here that human, Drosophila, and C. elegans bestrophins form oligomeric chloride channels, and that human bestrophin is sensitive to intracellular calcium. Each of 15 missense mutations asscociated with VMD greatly reduces or abolishes the membrane current. Four of these mutant bestrophins were coexpressed with the wild type and each dominantly inhibited the wild-type membrane current, consistent with the dominant nature of the disease. These experiments establish the existence of a new chloride channel family and VMD as a channelopathy.

Figure 1

Whole-cell currents from 293 cells transiently transfected with different bestrophin cDNAs. (a) Averaged and normalized current–voltage (I–V) relations for hBest1 (n = 16 cells), ceBest1 (n = 7), dmBest1 (n = 7), and hBest2 (n = 7); error bars indicate standard deviation (SD). For each bestrophin, the I–V curves for each cell were normalized to a value of −1.0 at −150 mV before averaging was done. Recordings were performed with standard extracellular and pipette solutions. Insets show sample current traces produced by 320-ms voltage steps from a holding potential of −50 mV to voltages between −150 and +80 mV in 10-mV increments. Current amplitudes at the end of the voltage steps were used for I–V plots. Open symbols in a Left show the averaged I–V relation from 11 control EGFP-transfected cells, after scaling it at −150 mV in proportion to the average hBest1 current; the SDs for the control cells are too small to be discernible. (b) Current amplitudes ± SD at −150 mV for the cells in a. The wide variation in the currents recorded from individual cells presumably reflects variation in plasmid uptake and level of bestrophin production in transiently transfected cells.

Figure 2

Ion permeability and DIDS sensitivity of bestrophin currents. (a Left and Center) Dependence of the whole-cell current reversal potential recorded from hBest1-transfected cells on extracellular Na⁺ and Cl⁻ concentrations. Na⁺ was replaced by equimolar N-methyl-d-glucamine (2 cells, each represented by a different symbol), and Cl⁻ was replaced by gluconate (mean ± SD from 5 cells). (Right) Permeability ratio of NO to Cl⁻ for 293 cells transfected with hBest1 (4 cells) and hBest2 (3 cells) determined by replacing Cl⁻ with equimolar NO. The curves are derived from the Goldman–Hodgkin–Katz equation with P_NO3/P_Cl ratios of 2.7 and 5.8, respectively. Initial absolute currents in standard extracellular solution and at +80 mV were 293, 307, 366, 1562, and 2320 pA for the cells in Center, and 155, 251, 545, 904 pA (hBest1), and 218, 238, 519 pA (hBest2) for the cells in Right. (b) Effect of the Cl channel blocker DIDS on the whole-cell current of a hBest1-transfected cell. DIDS (0.5 mM) was bath applied. In four experiments, 89% ± 1% (mean ± SD) of the current at +80 mV was reversibly suppressed.

Figure 3

Cysteine-less hBest1 is resistant to inactivation by sulfhydryl-specific reagents. (a) Locations of the 5 cysteines in hBest1 in the transmembrane topography of bestrophin proposed by Bakall et al. (11) in which the N and C termini face the cytosol. (b) Time course of whole-cell current inactivation recorded at +80 mV after addition of MTSEA to 2.5 mM at time 0 for cysteine-less hBest1 (○; n = 4 cells) or wild-type hBest1 (●; n = 4 cells). Mean current amplitudes at +80 mV recorded immediately before MTSEA addition were 1,355 ± 928 pA for cysteine-less hBest1 and 795 ± 674 pA for wild-type hBest1. At each time point, currents were recorded from −120 to +80 mV in 40-mV steps, with an interstep holding potential of 0 mV. In situations where the time points did not coincide for individual cells, linear interpolations of the measurements were made before averaging.

Figure 4

Specificity of bestrophin oligomerization. (a) Proposed transmembrane topography of bestrophin (11) with dark regions indicating high levels of homology among family members. Epitope tags, Rim3F4 (R; ref. 16) and 6× myc (M) were appended at the extreme C terminus. Arrowheads demarcate the region coded by exon 10 that is missing in the “short” splice variant of hBest1. (b–d) Immunoprecipitation of different epitope-tagged bestrophins with anti-M antibodies followed by immunoblotting with anti-R antibodies. In each immunoblot, 2.5% of the total membrane protein input was loaded in lane T, and 10% of the immunopurified sample was loaded in lane IP. In the diagram to the left of each immunoblot, the circles that are touching indicate the cotranfected bestrophins and the inverted “Y” indicates the immunoprecipitating monoclonal antibody. These schematic diagrams should not be taken to imply any particular stoichiometry. (b) Cotransfection of hBest1-M and hBest1-R leads to efficient association of the two proteins (Upper), whereas mixing of detergent extracts from independently transfected cells does not (Lower). (c) Coprecipitation from cells triply transfected with full-length hBest1-M and full-length hBest1-R (represented by “L”), and the short splice variant of hBest1-R (represented by “S”) shows that association does not depend on an intact C-terminal domain. (d) Top two rows, coprecipitation from cells triply transfected with one M-tagged and two different R-tagged bestrophins. The relative efficiencies of association can be estimated by comparing the “T” and “IP” lanes. Bottom row, control experiments in which detergent extracts from cells cotransfected with two bestrophins were mixed with detergent extracts from cells transfected with a third bestrophin (shown in the schematic as the separated ball). In each case, mixing of proteins from independently transfected cells fails to produce a complex. Bars to the left of the immunoblots indicate molecular-mass markers, from top to bottom: 173, 111, 80, 61, 49, 36, and 25 kDa.

Figure 5

Stoichiometry of hBest1 oligomeric complexes. The 293 cells were transfected with a mixture of hBest1-R and hBest1-M expression plasmids at the indicated ratios and labeled for 24 h with [³⁵S]methionine. Bestrophin complexes were purified with either anti-M (Left) or anti-R (Right) antibodies. hBest1-R and hBest1-M monomers were resolved by SDS/PAGE, visualized by autoradiography (Upper), and the radioactivity in each bestrophin band was measured with a phosphorimager (Lower; shown as the hBest1-R/hBest1-M protein ratio). hBest1-M and hBest1-R contain the same number of methionines; the slower electrophoretic mobility of hBest1-M is caused by the presence of six tandem copies of the myc-tag. The autoradiograms and accompanying plots show the results from one experiment in which solubilized hBest1 was preenriched by using Ni-NTA resin (see Materials and Methods). (Inset) Mean and SD for four similar experiments without preenrichment. Molecular mass markers are as described for Fig. 3.

Figure 6

Calcium regulation of hBest1 current. (a) Under whole-cell recording, a hBest1-transfected cell was dialyzed with a caged-Ca²⁺ solution containing 4 mM NPEGTA/0.4 mM Ca²⁺ (free [Ca²⁺] ≈10 nM). Two light flashes, each of 2-s duration, were applied at the times indicated. From a holding potential of −50 mV, voltage-ramp pulses from −100 to +80 mV and then back to −100 mV were applied before and after each flash. (b) Conductance increase induced by the photorelease of Ca²⁺. The I–V relations marked “1” and “2” were replotted from the ramp pulses 1 and 2 in a. (c--e) Control experiments showing that the photo-induced current was caused by a rise in intracellular Ca²⁺ concentration. (c) A pipette solution containing BAPTA in addition to NPEGTA was used instead, to suppress the photo-released [Ca²⁺] in a hBest1-transfected cell. (d) Instead of NPEGTA, a photo-insensitive chelator, EGTA, was loaded into a hBest1-transfected cell. (e) NPEGTA was loaded into a control EGFP-transfected cell; otherwise as in a. In these control experiments, current amplitudes were almost identical before and after the flash marked “F”. (f) Collected data from the experiments in a–e. In each case, the difference in current amplitude at +80 mV before and after the first flash is plotted as mean and SD. The number below each bar indicates the number of cells tested.

Figure 7

Currents in 293 cells transfected with hBest1 missense mutants responsible for VMD. (a) Locations of 15 disease-associated mutations in the hBest1 model of Bakall et al. (11). (b) Immunoblot of Rim3F4-tagged wild-type and 15 mutant hBest1 proteins from transiently transfected cells. “Control” indicates untransfected cells. (c) Whole-cell currents at +80 mV from 293 cells transfected with wild-type (“WT”) or with 11 mutant hBest1 plasmids, or with EGFP expression plasmid only, measured as in Fig. 1a. Dots indicate data from individual cells, and bars represent the mean ± SD. (d–f) Four hBest1 missense mutants responsible for vitelliform macular dystrophy coassemble with wild-type hBest1 and decrease whole-cell current. (d) Coimmunoprecipitation of wild-type hBest1-M together with either wild-type or mutant hBest1-R from cotransfected cells as described for Fig. 3. IP, immunoprecipitate; IB, immunoblot. (e) Whole-cell recordings from transfected 293 cells showing that the four indicated mutant proteins gave smaller currents when transfected alone, and inhibited the wild-type current when cotransfected with wild-type hBest1. For transfection in a 3.5-cm-diameter dish, a total volume of 4 μl containing 0.2 μg of EGFP plasmid and either 2 μg of wild-type or mutant hBest1 plasmid (for single transfections) or 1 μg of each plasmid (for cotransfections) was mixed with 5 μl of Fugene 6 reagent. Indicated current amplitude was at +80 mV, measured as in Fig. 1a. (f) Experiment showing that inhibition of whole-cell current in the cotransfection experiments of e did not simply arise from competition between wild-type and mutant at the level of protein or RNA synthesis. Each 3.5-cm-diameter dish received 2 μg of wild-type hBest1 (Left), 1 μg of wild-type + 1 μg R92C mutant (Center), or 1 μg of wild-type + 1 μg Fz8-IgG (Right) plasmid, together with 0.2 μg of EGFP plasmid, in a total volume of 2 μl with 5 μl of Fugene 6 reagent. All data in an individual panel were obtained from transfections performed side by side. Each transfection of mutant alone or mutant plus wild-type produced a set of whole-cell currents that were significantly smaller than the wild-type controls measured in parallel (P < 0.05, two-tailed t test).