N-Terminal Fatty Acylation of Transducin Profoundly Influences Its Localization and the Kinetics of Photoresponse in Rods

Abstract

N-terminal acylation of the α-subunits of heterotrimeric G-proteins is believed to play a major role in regulating the cellular localization and signaling of G-proteins, but physiological evidence has been lacking. To examine the functional significance of N-acylation of a well understood G-protein α-subunit, transducin (Gα_t), we generated transgenic mice that expressed a mutant Gα_t lacking N-terminal acylation sequence (Gα_tG2A). Rods expressing Gα_tG2A showed a severe defect in transducin cellular localization. In contrast to native Gα_t, which resides in the outer segments of dark-adapted rods, Gα_tG2A was found predominantly in the inner compartments of the photoreceptor cells. Remarkably, transgenic rods with the outer segments containing Gα_tG2A at 5–6% of the Gα_t levels in wild-type rods showed only a sixfold reduction in sensitivity and a threefold decrease in the amplification constant. The much smaller than predicted reduction may reflect an increase in the lateral diffusion of transducin and an increased activation rate by photoexcited rhodopsin or more efficient activation of cGMP phosphodiesterase 6 by Gα_tG2A; alternatively, nonlinear relationships between concentration and the activation rate of transducin also potentially contribute to the mismatch between the amplification constant and quantitative expression analysis of Gα_tG2A rods. Furthermore, the G2A mutation reduced the GTPase activity of transducin and resulted in two to three times slower than normal recovery of flash responses of transgenic rods, indicating the role of Gα_t membrane tethering for its efficient inactivation by the regulator of G-protein signaling 9 GTPase-activating protein complex. Thus, N-acylation is critical for correct compartmentalization of transducin and controls the rate of its deactivation.

Introduction

Heterotrimeric G-proteins mediate the transduction of a variety of extracellular signals that activate G-protein-coupled receptors at the plasma membrane (Bourne, 1997; Pierce et al., 2002; Cabrera-Vera et al., 2003). The vertebrate rod phototransduction cascade represents one of the best model systems to study the mechanisms of G-protein signaling. Rod transducin (G_t) couples photoexcitation of rhodopsin (R) to the activation of cGMP phosphodiesterase 6 (PDE6). The GTP-bound α-subunit of G_t (Gα_tGTP) binds to the γ-subunit of PDE6 (Pγ), activating the enzyme. The Gα_tGTP/PDE6 complex is deactivated when bound GTP is hydrolyzed as a result of intrinsic GTPase activity of Gα_t controlled by the GTPase-activating protein (GAP) complex, regulator of G-protein signaling 9-1 (RGS9-1)/Gβ₅/RGS9 anchor protein (R9AP) (Burns and Baylor, 2001; Burns and Arshavsky, 2005).

Lipid modifications of G-proteins are thought to play a major role in the regulation of G-protein cellular localization and signaling (Wedegaertner et al., 1995; Higgins and Casey, 1996; Chen and Manning, 2001). Most G-protein γ-subunits (Gγs) are modified with a thioether-linked isoprenoid geranylgeranyl (C20) attached to Cys residues within the C-terminal “CAAX” box (Escriba et al., 2006). In contrast, the rod-specific Gγ₁ carries the farnesyl moiety (C15), which facilitates light-dependent translocation of G_t from the rod outer segment (ROS) to the inner compartments, thereby contributing to light adaptation (Kassai et al., 2005). Gα-subunits are typically modified with an amide-linked fatty acid myristate (C14:0) at the extreme N-terminal Gly residue and/or with a thioester-linked palmitate (C16:0) at Cys residues near the N termini (Wedegaertner et al., 1995; Chen and Manning, 2001). Uniquely, rod Gα_t is heterogeneously fatty N-acylated with C12:0, C14:0, C14:1, and C14:2 moieties (Neubert et al., 1992). The lower hydrophobicity of the Gα_t acyl residues also assists light-dependent redistribution of transducin between the outer segment (OS) and inner regions of rods (Lobanova et al., 2007).

Although the heterogeneity of Gα_t lipid modification may influence certain aspects of phototransduction and light adaptation (Neubert and Hurley, 1998; Lobanova et al., 2007), the significance of the lipid modification itself is expected to be much more profound. The N-terminal acylation may be required for proper transport and localization of transducin to the OS. On the disc membrane, the Gα_t lipid anchor may control the lateral diffusion and collisional coupling of Gα_t with photoexcited rhodopsin (R*). Furthermore, the Gα_t lipid is potentially critical for PDE6 activation and inactivation. The activated Gα_tGTP/PDE6 complex is membrane bound and stabilized by the Gα_t and PDE6 lipid modifications (Catty et al., 1992; Clerc and Bennett, 1992). The ability of the GAP complex to rapidly inactivate the Gα_tGTP/PDE6 complex depends on the membrane attachment of the GAP complex via the anchor protein, R9AP (Lishko et al., 2002, Hu et al., 2003).

Despite growing data supporting the roles of N-acylation of Gα subunits for localization and transduction efficiency of G-proteins, functional evidence for the importance in cell signaling is lacking. To probe the functional significance of the N-acylation of Gα_t in intact cells, we have produced and examined transgenic mice expressing an acylation-deficient mutant, Gα_tG2A, in rod photoreceptors.

Materials and Methods

Generation of Gα_tG2A transgenic mice.

A pBRH Gα_t transgenic construct contained the mouse Gα_t genomic sequence of ∼5.5 kb (Raport et al., 1989) flanked by the 4.4 kb mouse rod opsin promoter fragment (Lem et al., 1991) and polyadenylation signal (Fig. 1A). The transgene sequence was mutated previously to introduce the Glu-Glu (EE) monoclonal antibody epitope to assess expression levels of mutant transducins in mouse retinas (Kerov et al., 2005; Moussaif et al., 2006). The G2A substitution was created using the QuickChange mutagenesis kit (Stratagene, La Jolla, CA). A DNA fragment of 9 kb was released from the pBRH-Gα_tEEG2A plasmid by restriction with NotI and gel purified. The DNA fragment was microinjected into mouse embryos at the transgenic core facility at the University of Iowa. Transgenic mice were identified by PCR of mouse tail DNA with a pair of primers chosen to amplify a 300 bp fragment surrounding the junction between the Gα_t and polyadenylation signal sequences. Six potential Gα_tG2A transgenic mouse founders were identified and mated to C57BL/6 mice. The founder line with the highest expression level of Gα_tG2A was selected for breeding with Gα_t knock-out mice (Gα_t^−/−) to produce Gα_tG2A/Gα_t^+/− transgenic mice hemizygous for native Gα_t allele. Gα_tG2A/Gα_t^+/− mice were further mated with Gα_t^−/− mice to move the transgene into the knock-out (Gα_t^−/−) background. Gα_tG2A/Gα_t^−/− mice were identified by the absence of PCR product using primers designed to amplify a DNA sequence only from the wild-type Gα_t allele. All experimental procedures involving the use of mice were performed in accordance with the National Institutes of Health (NIH) guidelines and the protocol approved by the University of Iowa Animal Care and Use Committee.

Figure 1.

A, Transgenic construct used for the generation of mutant Gα_tG2A mice. B, Expression levels of G2A mutant in Gα_tG2A/Gα_t^+/− and Gα_tG2A/Gα_t^−/− mice. Retinas from transgenic and control (Gα_t^+/+) mice were homogenized and subjected to SDS-PAGE. Sample loads are in micrograms of retinal protein (ret. pr.). Immunoblot (IB) analysis was performed with anti-rod Gα_t and anti-EE antibodies. C, Quantification of protein levels from the anti-EE antibody immunoblot in B. The relative level of Gα_tG2A in Gα_tG2A/Gα_t^−/− mice is 1.5-fold higher than in Gα_tG2A/Gα_t^+/− mice (calculated as the ratio of the linear fit slopes). D, Immunoblot analyses of key phototransduction proteins. Samples contained 2.5, 5, and 10 μg of retinal homogenate from 4-month-old Gα_tG2A/Gα_t^−/− or control WT mice. The antibodies are described in Materials and Methods. E, Quantification of the relative protein levels from the immunoblot in D. Integrated densities for each protein band were plotted as a function of retinal homogenate load to determine the linear fit slopes and the ratios of the slopes for WT and mutant mice. Arr, Arrestin.

Immunoblot analysis and quantification of transgene expression.

Total mouse retinal homogenates were obtained by solubilization of two retinas in 100 μl of 10% SDS-Na using brief sonication. Protein concentrations were determined using the DC protein assay (Bio-Rad, Hercules, CA) and bovine serum albumin dissolved in 10% SDS-Na as the standard. Typically, the total protein content of a homogenate obtained from a single mouse retina was ∼500 μg. Samples of retinal homogenates were subjected to SDS-PAGE in 12% gels, transferred onto nitrocellulose membranes BA 85 (Schleicher and Schuell, Keene, NH), and probed with the following primary antibodies (1:1000 dilution): anti-rod Gα_t (K-20), anti-Gβ₁ (M-14), and anti-Gγ₁ (P-19) (Santa Cruz Biotechnology, Santa Cruz, CA); EE monoclonal (Covance, Princeton, NJ); monoclonal anti-rhodopsin 1D4 (National Cell Culture Center, Minneapolis, MN); anti-PDE6αβ MOE antibodies (Cytosignal, Irvine, CA); anti-arrestin (PA1–731; ABR, Golden, CO), and anti-RGS9 antibodies (Elmira Biologicals, Iowa City, IA). According to the manufacturer, the epitope of anti-rod Gα_t antibody K-20 is located within residues 80–120 of human Gα_t. Thus, the affinity of this antibody for Gα_t should not be affected by the G2A or EE mutations. The antibody–antigen complexes were detected using secondary anti-rabbit, anti-mouse, or anti-sheep antibodies (1:10,000) conjugated to horseradish peroxidase (Santa Cruz Biotechnology) and ECL reagent (GE Healthcare, Piscataway, NJ). Nitrocellulose membranes were exposed to film, and integrated densities of scanned individual bands were measured with Image J 1.33u software.

Immunohistochemistry.

For dark adaptation, mice were kept in the dark for at least 12 h. All dark procedures were performed under infrared illumination using night vision goggles. Eyeballs were enucleated, and the corneas were perforated with a 21 gauge needle, fixed in PBS with 4% formaldehyde for 1 h, and cut in half. The cornea and lens were removed, and the eyecups were submersed in a 30% sucrose solution in PBS for 5 h at 4°C. The eyecups were then embedded in tissue freezing medium (Triangle Biomedical Sciences, Durham, NC) and frozen on dry ice. Radial sectioning (10 μm) of the retina was performed using a cryomicrotome HM 505E. Retinal cryosections were air dried and kept at −80°C. The sections were warmed up to room temperature and incubated in PBS/0.1% Triton X-100 for 30 min. Next, 2% goat serum and 5% BSA in PBS was added, and incubation continued for 30 min. Sections were then incubated with rabbit anti-rod Gα_t antibody (K-20; 1:2000; Santa Cruz Biotechnology) and a monoclonal anti-EE antibody (1:2000; Covance) for 3 h. After a 1 h incubation with goat anti-rabbit AlexaFluor 555 or goat anti-mouse AlexaFluor 488 secondary antibodies (1:1000; Invitrogen, Carlsbad, CA), the sections were visualized using a Zeiss (Thornwood, NY) LSM 510 confocal microscope. To evaluate mouse retina morphology, eyeballs were fixed in 2.5% gluteraldehyde in 0.1 m cacodylate buffer for 2 h at 22°C, cut in half to remove the cornea and lens, dehydrated through a series of acetones, and embedded in Epon. Radial sections (1 μm) were stained with toluidine blue and photographed through a Nikon (Tokyo, Japan) Optiphot microscope using a Spot camera (Diagnostic Instruments, Sterling Heights, MI).

Biochemical analysis of transducin function in mouse ROS membranes.

Mouse ROS membranes were prepared using a procedure that allows isolation of osmotically intact and sealed ROS, thereby guarding against loss of OS proteins (Tsang et al., 1998). Rhodopsin concentrations were determined using the difference in absorbance at 500 nm before and after photobleaching of ROS preparations solubilized in 1% Ammonix LO. To measure light-dependent binding of GTPγS, ROS (10 μl) from dark-adapted Gα_tG2A/Gα_t^−/− or control Gα_t^+/+ mice containing 10 μm rhodopsin were mixed with 5 μl of 10 μm [³⁵S]GTPγS under infrared illumination. After 10 s of incubation, the amount of bound GTPγS was determined in a 6 μl aliquot by the nitrocellulose filter binding assay. The rest of the sample was bleached to determine the GTPγS binding in the light.

The single-turnover GTPase measurements were initiated by mixing photobleached ROS (12 μl) containing 10 μm rhodopsin with 3 μl of 0.1 μm [γ-³²P]GTP. The time course of ³²P_i formation was determined using the activated charcoal procedure after the reactions were stopped with perchloric acid. The GTPase rate constants were calculated by fitting the experimental data to an exponential function:

where k is a rate constant for GTP hydrolysis.

To measure PDE6 activation by transducin, photobleached ROS (3 μm R) were incubated with 3 μm cGMP for 3 min to stabilize the PDE6 basal activity (Tsang et al., 1998). The reactions were started by the addition of 4 mm [³H]cGMP with or without 10 μm GTPγS. To activate PDE by limited trypsinolysis, ROS samples were incubated with tosyl phenylalanyl chloromethyl ketone-treated trypsin (50 μg/ml) for 4 min on ice followed by incubation with soybean trypsin inhibitor (250 μg/ml) for 5 min at 25°C. cGMP hydrolysis was terminated after 30 s by heating the samples for 2 min at 100°C. After cooling, 0.1 U of bacterial alkaline phosphatase (Sigma-Aldrich, St. Louis, MO) was added to each sample followed by incubation for 20 min at 37°C. Nonhydrolyzed cGMP was removed with AG1-X2 anion exchange resin (Bio-Rad), and [8-³H]guanosine was counted in a liquid scintillation counter.

To assess pertussis toxin-catalyzed Gβ₁γ₁-dependent ADP ribosylation of Gα_t, dark-isolated or bleached wild-type (WT) ROS (0.6 μm R) and Gα_tG2A/Gα_t^−/− ROS membranes (6 μm R) were incubated with 3 μg/ml of pertussis toxin (preactivated by 15 min incubation at 30°C with 100 mm dithiothreitol and 0.2% SDS) and 5 μm [³²P]nicotinamide adenine dinucleotide ([³²P]NAD) for 1 h at 25°C in the presence or absence of 30 μm AlCl₃ and 10 mm NaF. The reactions were stopped by the addition of SDS-sample buffer and analyzed by SDS-PAGE in 12% gels and autoradiography.

Suction electrode recordings and analysis.

Mice were cared for and handled following an approved protocol from the Animal Care and Use Committee of the University of California, Davis and in compliance with NIH guidelines for the care and use of experimental animals. Before an experiment, animals were dark adapted overnight for a minimum of 12 h. Animals were anesthetized and killed under infrared light, and the retinas were dissected and stored on ice in L-15 solution with 10 mm glucose and 0.1 mg/ml bovine serum albumin (Sigma-Aldrich). For recording, a small piece of retina was chopped into small pieces in a chamber containing the L-15 solution, and the chopped tissue was then placed in a recording chamber perfused with bicarbonate solution, pH 7.4, supplemented with 10 mm glucose held at 35–37°C. Individual rods were visualized under infrared light using a CCD camera (Stanford Photonics, Palo Alto, CA). The outer segment of an individual rod was gently drawn into the tip of a capillary pipette containing HEPES solution, pH 7.4. The membrane currents were recorded by a current-to-voltage converter (Axopatch 1B; Molecular Devices, Union City, CA) and low-pass filtered (eight-pole Bessel; Frequency Devices, Haverhill, MA) using 30 Hz corner frequency. Data were digitized at 200 Hz using IGOR-National Instruments acquisition software (IgorPro for NIDAQ for Windows; Wavemetrics, Lake Oswego, OR). Rods were presented with 10 ms flashes of 500 nm light. The intensity of the light was controlled by using calibrated neutral density filters.

The form of the single-photon response was calculated by squaring the average response to 30 or more dim (linear) responses and scaling it to coincide with the time course of the ensemble light-dependent variance over the initial rising phase. Assuming the predominant source of variance over this time scale arises from Poisson fluctuations in the number of photoexcited rhodopsins, the inverse of this scaling factor corresponds to the average number of photoisomerizations per trial. The average linear response was then divided by the average number of photoisomerizations per trial to yield the average single-photon response. The effective collecting area was calculated by dividing the average number of photoisomerizations per trial by the flash strength (in photons · μm⁻²).

The time constant of recovery for dim flash responses was determined by least-squares fitting of a single exponential to the final falling phase of the response; for bright flash responses, a straight line was fitted to the relationship between the time in saturation and the natural log of the flash strength up to ln i = 9 (Krispel et al., 2006). The integration time was calculated as the time integral of the dim flash response divided by the peak amplitude.

Results

Expression and localization of Gα_tG2A mutant in transgenic rods

The G2A substitution was introduced into a transgene previously used for expression of the monoclonal antibody EE epitope-tagged transducin mutant (Fig. 1A) (Kerov et al., 2005; Moussaif et al., 2006). The EE tag facilitates determination of the expression levels and localization of the Gα_tG2A mutant in the presence of native Gα_t in transgenic rods. The EE tag does not alter biochemical properties of transducin, and the flash responses from transgenic mouse rods expressing Gα_tEE are similar to those from the wild-type rods (Kerov et al., 2005; Moussaif et al., 2006). Several transgenic mouse lines expressing the EE-tagged Gα_tG2A mutant were established. The line with the highest expression level of Gα_tG2A was selected by immunoblot analysis with anti-EE antibody and used for breeding with Gα_t knock-out mice (Calvert et al., 2000) to move the transgene into the hemizygous (Gα_t^+/−) and knock-out (Gα_t^−/−) backgrounds. Based on the immunoblot analysis of mouse retinal homogenates with anti-rod Gα_t antibody, the total expression level of transducin-α (Gα_tG2A + native Gα_t) in Gα_tG2A/Gα_t^+/− mice was similar to the Gα_t level in wild-type (Gα_t^+/+) mice (Fig. 1B). The level of Gα_tG2A in the Gα_t knock-out background was ∼30% of the wild-type level (Fig. 1B). The blot analysis with EE antibody indicated that expression of Gα_tG2A in the Gα_t^−/− background was ∼1.5-fold higher than in the Gα_t^+/− background (Fig. 1B,C), suggesting that Gα_tG2A/Gα_t^+/− mice express ∼20% Gα_tG2A and 80% native Gα_t. Control immunoblot tests demonstrated that the levels of Gβ₁ and several major phototransduction components such as rhodopsin, PDE6αβ, arrestin-1, and RGS9-1 were essentially unchanged in Gα_tG2A/Gα_t^−/− mice (Fig. 1D,E). Also, Gα_tG2A/Gα_t^+/− and Gα_tG2A/Gα_t^−/− mice showed normal gross morphology of the retina with no apparent signs of retinal degeneration (data not shown).

In dark adapted rods, native transducin localizes to the OS. We examined subcellular localization of Gα_tG2A in transgenic rods by immunofluorescence confocal microscopy of mouse retinal sections using anti-rod Gα_t and anti-EE antibodies. In contrast to the native Gα_t, Gα_tG2A displayed extensive mislocalization to the inner compartments of rod photoreceptor cells (Fig. 2A). Only a very weak EE signal was seen in the OS of Gα_tG2A/Gα_t^+/− and Gα_tG2A/Gα_t^−/− mice (Fig. 2A). ROS membranes were isolated from dark-adapted Gα_tG2A/Gα_t^−/− and control Gα_t^+/+ mice to determine the fraction of the mutant that correctly localized to the OS. The ROS immunoblot analysis revealed that the level of Gα_tG2A in the OS is ∼5–6% of the Gα_t level in control mice (Fig. 2B,C). Thus, Gα_tG2A/Gα_t^−/− mice express Gα_tG2A at ∼30% of Gα_t in wild-type mice, but only 15–20% of the mutant Gα_t is able to reach the OS. This result is in agreement with the immunofluorescence staining of Gα_tG2A in the Gα_tG2A/Gα_t^−/− retinal sections (Fig. 2A).

Figure 2.

A, Localization of total Gα_t and mutant Gα_tG2A in dark-adapted (DA) mouse retinas. Gα_tG2A/Gα_t^+/− and Gα_tG2A/Gα_t^−/− retina sections were stained with rabbit anti-rod Gα_t antibody K-20 or a monoclonal anti-EE antibody and visualized with goat anti-rabbit AlexaFluor 555 and goat anti-mouse AlexaFluor 488 secondary antibodies using a Zeiss LSM 510 confocal microscope. Scale bar, 20 μm. IS, Inner segment; ONL, outer nuclear layer; OPL, outer plexiform layer. B, Representative immunoblot (IB) analysis of relative levels of Gα_tG2A in Gα_tG2A/Gα_t^−/− ROS and native Gα_t in WT ROS. C, Quantification of the relative protein levels from the immunoblot in B. The relative level of Gα_t in WT ROS is ∼17-fold higher than the level of Gα_tG2A in transgenic ROSs (calculated as the ratio of the linear fit slopes). The average ratio of the slopes in three similar experiments is 18 ± 2 (mean ± SE).

Biochemical analysis of Gα_tG2A

The key signaling properties of Gα_tG2A such as its interaction with Gβ₁γ₁, R*-dependent binding of GTPγS, GTPase activity, and the ability to activate PDE6 were examined using preparations of ROS membranes isolated from Gα_tG2A/Gα_t^−/− and control wild-type (Gα_t^+/+) mice. To assay the ability of Gα_tG2A to bind Gβ₁γ₁, we measured the extent of pertussis toxin-induced ADP ribosylation of Gα_t at Cys347 (van Dop et al., 1984; West et al., 1985; Watkins et al., 1985). Holotransducin, Gα_tβ₁γ₁, is a much-preferred substrate in this reaction compared with Gα_t alone (Watkins et al., 1985). The extent of ADP ribosylation of Gα_t in wild-type ROS and Gα_tG2A in transgenic ROS was similar, and both were inhibited comparably by the addition of AlF₄⁻ (Fig. 3A). AlF₄⁻ ions activate GDP-bound Gα_t subunits by mimicking the γ-phosphate of GTP and causing dissociation of Gβ₁γ₁. Thus, Gα_tG2A is capable of binding Gβ₁γ₁ and is present in ROS of Gα_tG2A/Gα_t^−/− mice as a heterotrimeric protein Gα_tG2Aβ₁γ₁. The binding of wild-type Gα_tβ₁γ₁ to R* inhibited pertussis toxin-catalyzed ADP ribosylation of Gα_t, apparently by masking the Cys residue (van Dop et al., 1984; Watkins et al., 1985; West et al., 1985). Likewise, ADP ribosylation of Gα_tG2Aβ₁γ₁ was inhibited by light, suggesting specific binding of the mutant transducin to R* (Fig. 3A).

Figure 3.

A, Pertussis toxin-catalyzed ADP ribosylation of Gα_t and Gα_tG2A. Dark-isolated (D) or bleached (L) WT ROS (0.6 μm R) and Gα_tG2A/Gα_t^−/− ROS (6 μm R) were incubated with preactivated pertussis toxin (3 μg/ml) and 5 μm [³²P]NAD for 1 h at 25°C in the presence or absence of 30 μm AlCl₃ and 10 mm NaF. The reactions were stopped by the addition of SDS-PAGE sample buffer and analyzed by SDS-PAGE and autoradiography. B, Light-dependent binding of GTPγS to control WT and Gα_tG2A/Gα_t^−/− ROS. Mouse ROS (10 μl, 10 μm R) were mixed with 5 μl of 10 μm [³⁵S]GTPγS under infrared illumination. After 10 s incubation, the amount of bound GTPγS was determined in a 6 μl aliquot by the nitrocellulose filter binding assay. The rest of the sample was bleached to determine the GTPγS binding in the light. Error bars indicate SE (n = 3). C, Transducin GTPase activities in ROS from control WT and Gα_tG2A/Gα_t^−/− mice. The single-turnover GTPase measurements were initiated by mixing photobleached ROS (12 μl) containing 10 μm rhodopsin with 3 μl of 0.1 μm [γ-³²P]GTP. The time course of ³²P_i formation was determined using the activated charcoal procedure after the reactions were stopped with perchloric acid. The GTPase rate constants from single-phase exponential fits for the control (■) and Gα_tG2A/Gα_t^−/− (▴) ROS were 0.15 ± 0.0 and 0.09 ± 0.00 s⁻¹ (mean ± SE; n = 3). D, PDE6 activation in ROS membranes. cGMP hydrolysis was measured in suspensions of bleached ROS membranes (3 μm R) in the absence or presence of 10 μm GTPγS using a 30 s incubation with 4 mm [³H]cGMP. Trypsin (Tryp.)-treated ROS were prepared as described in Materials and Methods. Error bars indicate SE (n = 3).

The overall ability of Gα_tG2Aβ₁γ₁ to be activated by R* was determined by light-dependent binding of GTPγS to ROS membranes from Gα_tG2A/Gα_t^−/− mice. The level of light-dependent binding of [³⁵S]GTPγS to Gα_tG2A/Gα_t^−/− ROS was ∼16-fold lower than (6% of) that of wild-type ROS (Fig. 3B). The decrease in [³⁵S]GTPγS binding to Gα_tG2A ROS is very similar to the reduction in the expression level of the mutant Gα_t in the ROS fraction (18-fold) (Fig. 2B), indicating that the entire pool of Gα_tG2Aβ₁γ₁ can be activated by R* in transgenic rods.

The GTPase activities of Gα_tG2A and native Gα_t in Gα_tG2A/Gα_t^−/− and wild-type ROS membranes were measured under single turnover conditions ([GTP] < [G_t]). A low substrate concentration (20 nm [γ-³²P]GTP) was used to ensure that the single turnover conditions held for the transgenic ROS with low levels of Gα_tG2A. The rates of 0.15 and 0.09 s⁻¹ for control and Gα_tG2A/Gα_t^−/− ROS preparations, respectively, were determined from the single-exponential fits to the data (Fig. 3C). Hence, the G2A mutation caused an ∼1.7-fold decrease in the rate of GTP hydrolysis by Gα_t.

The ability of Gα_tG2A to activate its effector, PDE6, was determined by measuring the rate of cGMP hydrolysis induced by the addition of GTPγS. The presence of GTPγS caused marked stimulation of cGMP hydrolysis in control ROS but had an approximately sevenfold smaller effect in Gα_tG2A/Gα_t^−/− ROS (Fig. 3D). As a control, ROS preparations were also treated with trypsin, which cleaves the inhibitory Pγ subunit and produces full activation of the PDE6 present in the assay. The addition of trypsin resulted in equivalent enzyme activity in control and transgenic ROS preparations (Fig. 3D), indicating that the lower GTPγS-induced PDE6 activation in Gα_tG2A/Gα_t^−/− ROS was not attributable to a reduction in PDE6 levels. Given the ∼18-fold reduction in Gα_tG2A expression in the transgenic rods, the sevenfold reduction in GTPγS-induced PDE6 activation indicates that the mutant Gα_t not only is competent to activate its effector, but seems to do so even more efficiently than wild-type Gα_t.

Physiology of Gα_tG2A rods

To assess the ability of nonacylated Gα_t to mediate phototransduction in intact cells, suction electrodes were used to record the light-induced decrease in plasma membrane current mediated by the closure of cGMP-gated ion channels (for review, see Burns and Arshavsky, 2005). The amplitude and time course of the light response provides a quantitative measure of amplification and deactivation of the phototransduction cascade in the intact cell (for review, see Pugh and Lamb, 2000). Gα_tG2A/Gα_t^−/− rods responded to brief flashes of light in a manner similar to that of wild-type rods (Fig. 4A), indicating that Gα_tG2A indeed becomes activated by rhodopsin and in turn can bind and activate PDE6, generating sufficient cGMP hydrolysis to close cGMP-gated channels on the plasma membrane. However, Gα_tG2A/Gα_t^−/− rods were less sensitive than those of wild-type mice; the flash strength needed to generate a half-maximal response was sixfold brighter than that for wild-type rods (Fig. 4B, Table 1). This decrease in sensitivity was not associated with a change in the ability of Gα_tG2A/Gα_t^−/− rods to capture incident photons, because the effective collecting area of 0.17 ± 0.03 μm² (n = 11) calculated by variance-to-mean analysis (see Materials and Methods) was similar to that of 0.23 ± 0.03 μm² (n = 9) measured for wild-type rods (p = 0.20). Thus, the reduced sensitivity of Gα_tG2A/Gα_t^−/− rods was not caused by a decrease in photon capture, but rather reduced activation of Gα_tG2A by photoexcited rhodopsin and/or reduced activation of PDE6 by Gα_tG2A. Consistent with this idea, the average response generated by a single photoexcited rhodopsin was smaller for Gα_tG2A/Gα_t^−/− rods than wild-type rods (Fig. 5A, Table 1). The population average Gα_tG2A response deviated from that of the average wild-type response early on the rising phase of the response (Fig. 5B,C), reaching a twofold smaller peak slightly earlier. The early rising phase of the response time course, r(t), is well described by the activation theory of Pugh and Lamb (Lamb and Pugh, 1992; Pugh and Lamb, 1993):

where r_max is the saturating amplitude, A is the amplification constant, Ö is the average number of photoexcited rhodopsins, and t_eff is an effective delay, which we assumed to be 2 ms. This equation was fitted to the population average single-photon responses (φ = 1) to yield calculated A values of 12.4 s⁻² for wild-type and 4.10 s⁻² for Gα_tG2A rods (Fig. 5C).

Figure 4.

Gα_tG2A rods are less sensitive than normal. A, Representative families of responses from a wild-type rod (top) and a transgenic Gα_tG2A/Gα_t^−/− rod. Flashes (10 ms, 500 nm) were delivered at t = 0. B, The flash strengths needed to generate the same relative response amplitudes were higher for the Gα_tG2A/Gα_t^−/− rod. Data are from the same cells as in A. Curves are saturating exponential fits.

Table 1.

Characteristics of wild-type and N-acylation-deficient Gα _t rods

	Dark current (pA)	Ioa (photons/μm2)	Integration time (ms)	Dim flash τrec (ms)	Time to peak (ms)	Elementary amplitude (pA)	Bright flash τDb (ms)
G2A−/−	13.9 ± 0.7 (33)	341.1 ± 24.2 (28)	407 ± 30 (27)	487 ± 32 (26)	102 ± 6 (30)	0.23 ± 0.03 (11)	672 ± 98 (6)
G2A+/−	10.4 ± 1.1 (11)	65.6 ± 16.3 (7)	259 ± 32 (9)	245 ± 27 (10)	116 ± 9 (10)	0.58 ± 0.15 (5)	184 ± 14 (6)
Wild type	10.8 ± 0.9 (15)	53.0 ± 3.9 (14)	324 ± 30 (13)	239 ± 19 (13)	116 ± 7 (14)	0.51 ± 0.07 (10)	224+ 11 (10)

Numbers in parentheses represent n.

^aFlash strength that elicited a half-maximal response.

^bDominant time constant for recovery of saturating flash responses.

Figure 5.

Population average single-photon responses reveal decrease in amplification in Gα_tG2A/Gα_t^−/− rods. A, Un-normalized average single-photon response from 11 Gα_tG2A/Gα_t^−/− rods (bottom trace) and 10 wild-type rods (top trace). The average dark current for the wild-type rods was 10.3 pA, and that for the Gα_tG2A rods was 16.6 pA. B, Same responses as in A, on an expanded time scale. Error bars indicate SEM. C, Same responses as in A, on a further expanded time scale. The dotted curves are the best parabolic fits of Equation 23 of Pugh and Lamb (1993) (see Results, Physiology of Gα_tG2A rods, Eq. 2), yielding amplification constants of 12.4 s⁻² for the average wild-type response and 4.1 s⁻² for the average Gα_tG2A/Gα_t^−/− response.

The light responses of Gα_tG2A/Gα_t^−/− rods also showed abnormal recovery kinetics. Whereas the dim flash responses of wild-type rods showed a final recovery time course approximated by a single exponential function with a 240 ms time constant, dim flash responses of Gα_tG2A/Gα_t^−/− rods recovered more slowly, with a time constant of ∼500 ms. Likewise, for bright flash responses, the dominant time constant of recovery (Nikonov et al., 1998) was approximately threefold longer for Gα_tG2A rods than for wild-type rods (Table 1). The slower recovery in the transgenic rods is consistent with the prolonged lifetime of activated Gα_tG2A resulting from impaired GTPase activity (Fig. 3C).

Discussion

This work represents the first investigation of the functional significance of N-terminal acylation in transducin function. The immunofluorescence analysis of retinal sections from transgenic mice expressing the N-acylation-deficient Gα_t revealed a clear defect in Gα_tG2A localization. The major fraction of Gα_tG2A in the Gα_t knock-out background was found in the inner compartments of transgenic rods instead of the OS. The marked mislocalization of Gα_tG2A did not result from the disruption of the Gα_t–Gβ₁γ₁ interaction (Fig. 3A). Therefore, our results show for the first time the critical role of Gα_t N-acylation in transducin localization. Native holo-transducin carries two lipid anchors, N-acyl on Gα_t and C-terminal farnesyl on Gγ₁, which appear to be necessary and sufficient for trafficking Gα_tβ₁γ₁ to the OS in the dark-adapted retinas. Interestingly, Gβ₁γ₁ is relatively evenly distributed between the OS and the rod inner compartments in Gα_t knock-out mice (Zhang et al., 2003). Hence, the lipophilic nature of the transducin molecules determined by the lipid modifications decreases in the same order as the ability to correctly localize to the OS: Gα_tβ₁γ₁ > Gβ₁γ₁ > Gα_tG2Aβ₁γ₁. We surmise from this analysis that the lipid anchors on Gα_tβ₁γ₁, particularly the N-terminal acylation of Gα_t, represent key determinants for transducin compartmentalization to the OS.

The biochemical analysis of Gα_tG2A in transgenic ROS indicated that the lack of N-acylation of Gα_t does not block the activation of mutant transducin by R*, nor the GTPγS-induced activation of PDE6 by Gα_tG2A, although the extent of each of these reactions was severely reduced. This reduction is likely attributable to the ∼18-fold lower than normal concentration of Gα_tG2A. In light of the greatly reduced levels of activatable Gα_tG2A, it is remarkable that the sensitivity of Gα_tG2A/Gα_t^−/− rods was not correspondingly reduced. Instead, this 18-fold reduction in heterotrimer levels was associated with an amplification constant that was reduced only by a factor of three (Fig. 5).

From a theoretical analysis of the phototransduction cascade, the amplification constant is proportional, among other things, to the rate of G_t activation per R* (ν_RG), the coupling efficiency (c_GP; defined as the ratio of activated Gα_t to activated PDE6 subunits), and β_sub, the rate constant of cGMP hydrolysis by Gα_t-activated PDE subunit (Pugh and Lamb, 1993). Of these parameters, only ν_RG is expected to be dependent on expression level, because it is thought to be limited by the rate of diffusional encounter between a single R* and transducin molecules on the disc membrane (Lamb and Pugh, 1992; Pugh and Lamb, 1993; Calvert et al., 2001). For the mammalian rod at 37°C, the diffusion limit on ν_RG can be described by a simplified equation:

where D_R and D_G are the lateral diffusion coefficients of R* and transducin on the disc membrane, and C_G is the concentration (surface density) of transducin on the membrane (Lamb and Pugh, 1992; Pugh and Lamb, 1993). Thus, an ∼18-fold reduction of transducin concentration in the transgenic OS might be expected to cause an equivalent reduction in the amplification constant (and the rates of G_t and PDE6 activation per R*). Of course, if the G2A mutation also reduces c_GP and/or β_sub (which together define the ability of a given Gα_tG2A to activate a PDE6 subunit), then A would be expected to be even lower still.

How then do we explain the only threefold reduction in the amplification constant? One possibility is that the lack of N-acylation increases the lateral diffusion of G_t, thereby increasing its encounter rate with R* and the ν_RG rate. According to Equation 3 and taking the estimates D_R ∼ 1.5 and D_G ∼ 1.8 μm² · s⁻¹ at 37°C (Pugh and Lamb, 1993), an 18-fold reduction in C_G accompanied by a threefold reduction in A implies a coincident tenfold increase in D_G. On the other hand, Gα_tG2A may actually activate PDE6 more effectively than normal Gα_t (i.e., couple to PDE6 with higher c_GP and/or induce higher β_sub). Support for this comes in part from the results of Figure 3D, which showed a sevenfold decrease in GTPγS-dependent PDE6 activity, compared with the 18-fold decrease in Gα_tG2A expression. Still, the PDE6 activation results in vitro do not necessarily indicate greater ability of Gα_tG2A to activate PDE6 in intact rods, because the assay was conducted in diluted suspensions of ROS membranes. Finally, it may be that the amplification constant does not scale linearly with transducin expression. Although previous studies have demonstrated a clear correlation between A and outer segment G_t levels (Sokolov et al., 2002), the linearity of this relationship has not been explicitly studied. Because the surface density of transducin is comparable with the Michaelis constant for rhodopsin-transducin binding at 34°C (Heck and Hofmann, 2001), the rate of activation by transducin might be less than linearly dependent on transducin expression.

Another clear effect of the G2A mutation is the slower than normal recovery of flash responses of transgenic rods. The time constants of recovery of dim and bright flash responses of Gα_tG2A/Gα_t^−/− rods are approximately two and three times slower than normal, respectively (Table 1), consistent with the reduced GTPase activity of Gα_tG2A in ROS membranes (Fig. 3C). In mouse rods, the time constant of dim flash response recovery and the dominant time constant for bright flash response recovery are both sensitive to the expression levels of the RGS9-1/Gβ₅ complex (Krispel et al., 2006). The GAP activity of RGS9-1/Gβ₅ is greatly accelerated by formation of the membrane-bound complex with the membrane anchor, R9AP (Lishko et al., 2002; Hu et al., 2003). We propose that the N-acylation of Gα_t along with lipid modifications of PDE6 provides for tight tethering of the activated Gα_t/PDE6 complex to the membrane, which enables a more efficient deactivation of Gα_t by the GAP complex. Overall, this study demonstrates that in addition to the critical role in trafficking and subcellular localization, the N-acylation of transducin is essential for the normal recovery of rod photoresponses.