‘Rod’ cells in the eye enable us to see in starlight. Inside these cells, a protein called rhodopsin is activated by light, which leads to an electrical signal being produced that travels to the brain. The duration of the electrical signal depends on the time it takes for the rhodopsin to be deactivated. Rhodopsin is a member of a large class of receptor proteins known as G protein-coupled receptors that regulate many processes throughout the body.

Previous studies have shown that rhodopsin is deactivated by the attachment of phosphate groups to the protein. This allows another protein called arrestin to bind to rhodopsin. The phosphates can be attached to particular amino acids—the building blocks of proteins—at one end of rhodopsin. Three of these are a type of amino acid called serine. Previous work has shown that light increases the speed at which phosphate groups are added to these serines, suggesting that they are important for producing rapid electrical signals. The other three amino acids are of a different type—called threonine—but it is less clear what role they play in deactivating rhodopsin.

Here, Azevedo et al. studied mutant forms of rhodopsin that were missing the serines or threonines in mice. The experiments show that loss of the serines only slightly slowed the electrical signals. However, loss of the threonines resulted in much slower electrical signals that ended at random times. This was due to rhodopsin being less able to bind to arrestin.

Azevedo et al. propose a new model for how rhodopsin is deactivated. Once light activates the protein, phosphate groups are rapidly added to the serines, which begins to lower the activity of rhodopsin. However, it is the slower addition of phosphates to the threonines that is essential to promote arrestin binding and fully deactivate the protein. Other proteins belonging to the G protein-coupled receptor family also have these serines and threonines, and thus, may be regulated in a similar way.

G protein-coupled receptors (GPCRs) mediate an array of cellular responses with widely divergent kinetics (Lefkowitz and Shenoy, 2005). This diversity originates in part from differences in the processes controlling GPCR activity, which influence the duration and extent to which downstream pathways are activated (DeWire et al., 2007). Nonetheless, all but a few GPCRs desensitize through a common set of reactions: phosphorylation by G protein receptor kinases (GRKs) followed by the binding of arrestin. Interestingly, the >800 GPCRs expressed by humans are desensitized by only 7 GRKs and 4 arrestins (Wilden et al., 1986; Lohse et al., 1990; Pitcher et al., 1998; Gurevich and Gurevich, 2004). Given this apparent conservation of the desensitization reactions, how is diversity in GPCR lifetime achieved?

GPCR C-termini, where the desensitization reactions take place, typically have multiple possible phosphorylation sites (Krupnick and Benovic, 1998). Biochemical experiments indicate that some sites are more rapidly phosphorylated than others, for example, serine sites appear to be preferred in the case of the photopigment rhodopsin (Kennedy et al., 2001; Maeda, 2003). The pattern of phosphorylation sites can control the effect of arrestin binding on the receptor, that is, whether arrestin promotes receptor desensitization or internalization (Key et al., 2003; Kim et al., 2005; Ren et al., 2005; Butcher et al., 2011; Nobles et al., 2011; Shukla et al., 2011). The variety of patterns exhibited by different GPCRs has been compared to identification by different bar codes (Tobin et al., 2008; Nobles et al., 2011). This suggests that, similarly, the pattern of phosphorylation sites may determine the active receptor lifetime, and hence the duration of the resulting signals.

Desensitization of the photopigment rhodopsin provides a unique opportunity to study GPCR desensitization because the signal produced by the activation of a single GPCR can be measured directly (Baylor et al., 1979). Furthermore, past work characterizing responses produced by the activation of single rhodopsin molecules has identified an unexpected property of how this GPCR desensitizes: the signals produced by single rhodopsin activations vary much less than expected from other familiar signals controlled by single molecules (Baylor et al., 1979; Rieke and Baylor, 1998; Whitlock and Lamb, 1999; Field and Rieke, 2002; Doan et al., 2006). This low variability requires precise control of the timing of rhodopsin desensitization (Mendez et al., 2000; Doan et al., 2006, 2009). In particular, phosphorylation and arrestin binding need to work in a coordinated fashion so as to minimize brief rhodopsin activations produced by too rapid arrestin binding and long rhodopsin activations produced by too slow arrestin binding. How the required coordination between phosphorylation and arrestin binding works is unclear.

Here, we compare the roles of native serine and threonine sites in rhodopsin desensitization. Our experiments reveal an unexpected specificity of these reactions that underlies the tight coordination of phosphorylation and arrestin binding. Namely, native serine sites are phosphorylated rapidly and begin to lower rhodopsin's catalytic activity (Ohguro et al., 1993, 1994, 1995; Gibson et al., 2000; Mendez et al., 2000; Kennedy et al., 2001), but serine phosphorylation only weakly promotes arrestin binding; native threonine sites are phosphorylated more slowly but are essential to promote rapid arrestin binding. This coordination produces an orderly sequence of desensitization events and contributes to the precise control of rhodopsin's active lifetime. A similar interplay of serine and threonine sites may be an important general mechanism tuning the kinetics of arrestin binding and GPCR desensitization.

The results are organized as follows. First, we show that single-photon responses (SPRs) from wild-type (WT), serine-only, and threonine-only rhodopsin differ substantially. Second, we show that, despite the large differences in responses, arrestin binding and receptor phosphorylation remain the critical reactions that desensitize the mutant receptors. Third, we show that the slow step-like responses generated by serine-only rhodopsin can be attributed to slow arrestin binding. Finally, we use the prolonged responses of serine-only rhodopsin to compare the kinetics of downstream steps in the transduction cascade with those of rhodopsin desensitization.

Serine-only rhodopsin produces prolonged SPRs

Figure 1A depicts the canonical molecules and reactions responsible for GPCR desensitization. The native C-terminal sequence of mouse rhodopsin and the mutations examined in this study are also depicted: WT rhodopsin (Rho), rhodopsin with C-terminal serine residues replaced with alanines (S → A), and rhodopsin with C-terminal threonine residues replaced with alanines (T → A). Transgenic mouse strains were backcrossed with Rho^−/− mice so that these rods expressed only a single type of rhodopsin (Lem et al., 1999).

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We recorded responses to brief flashes of light in individual rod outer segments using suction electrodes (Yau et al., 1977). The flash strength was adjusted such that rods failed to respond to a majority of the flashes (‘failures’) but occasionally produced a quantized response. We used quantal analysis to isolate responses produced by the absorption of single photons (Figure 1B–G; see ‘Materials and methods’ and Figure 1—figure supplement 1 for isolation procedure). Average SPRs elicited by each type of rhodopsin differed, particularly in recovery kinetics (Figure 1H, bottom). Time-to-peak and peak amplitude were not significantly different across rhodopsin types (p > 0.14, Kruskal–Wallis test, corrected for multiple comparisons).

Individual responses to single rhodopsin activations (Figure 1B–G) reveal several features not apparent in the averages. In particular, T → A rhodopsin (Figure 1D) produced responses that plateau and then terminate abruptly. Overlaying all SPRs gathered from a rod illustrates the extent of this effect (Figure 1G); whereas responses elicited by Rho or S → A rhodopsin follow similar trajectories, the T → A responses plateau to a common level before terminating at random times. This gives rise to two bands, an upper one that thins over time as responses decay, and a lower one that collects the responses as they terminate. Thus, removal of serine vs threonine sites has quite different effects on SPR variability. Below, we characterize these differences to gain insight into the underlying mechanisms.

Removing serine and threonine sites increases response variability

We first estimated the duration of each response by determining the time at which the response terminated (Figure 2A, insets). Termination times were estimated using an algorithm to identify state transitions, in this case the return of the current to baseline (see ‘Materials and methods’). Figure 2A (top panel) plots the cumulative distributions of response durations from each recorded cell expressing a given rhodopsin. The pooled distributions are plotted below. Spread in the measurement is dominated by real variation in response duration across strains, as shown by two measures. First, the distributions are wider than the jitter produced by noise in the algorithm (see ‘Materials and methods’, Figure 2A-bottom, gray dots). Second, the distributions of response durations for individual cells of a given strain more closely resemble the pooled distributions for that strain than those of a different strain (Figure 2A). The response duration distributions do illustrate an overlap in response properties across strains, for example, the briefest T → A responses are similar to Rho responses both in kinetics (Figure 1B–D) and duration (Figure 2A). Likewise, Rho responses can occasionally last quite long (Baylor et al., 1984).

Figure 2

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The time course of response variability was measured from the ensemble variance at each point in time (variance of SPRs minus variance of failures). SPRs followed a consistent initial trajectory and then began to diverge near the time at which the mean response peaked. For all rhodopsin types, the variance peaked after the peak of the mean response (Figure 2C,D). This late peak of the variance reflects the fact that the rising phase and peak amplitude of the response vary little compared to the falling phase. The T → A mutant is an extreme case of this effect: the peak in the variance of the T → A responses coincides with the point at which approximately half of the responses have returned to baseline (2.9 s, compare to median duration 2.7 s in Figure 2B). This correspondence between the median response duration and the time at which the variance peaks is expected for a first-order decay process in which response duration is determined by a single stochastic step. Thus, the time course of the variance suggests that a single desensitization step is greatly prolonged for T → A rhodopsin.

Total response variability was quantified using the coefficient of variation (SD/mean) of the integrated SPRs, that is, the coefficient of variation of the response areas (CV_area). This measure is insensitive to whether rhodopsin desensitization is fast or slow relative to the kinetics of downstream steps in the transduction cascade, whereas the variability of the response at a given time (e.g., the peak) depends on the underlying kinetics (Hamer et al., 2003). The variance of the areas of the SPRs was corrected for the variance of the failures and then used to compute CV_area (Hamer et al., 2003; Doan et al., 2006, 2009). CV_area for WT rods (0.37 ± 0.02) was comparable to that found previously (Doan et al., 2006). Responses of S → A rods had increased variability (CV_area = 0.51 ± 0.02, Figure 2E); this increase in CV_area is quantitatively consistent with a model in which total WT rhodopsin activity is controlled by a series of ∼7 kinetic steps, 3 of which have been removed in S → A rhodopsin (Doan et al., 2006). By comparison, the measured increase in variability is greater than that predicted by models that posit that non-linear amplification of the receptor activity contributes substantially to low-response variability (Caruso et al., 2011, their table S6).

The CV_area for the T → A responses (0.78 ± 0.03, Figure 2E) was much higher, consistent with a strong disruption of the normally reproducible timing of rhodopsin desensitization. The different effects of removing threonines and serines were not predicted by previous models of rhodopsin desensitization; instead each kinetic step was assumed to represent identical or at least equivalent biochemical events (Rieke and Baylor, 1998; Hamer et al., 2003; Doan et al., 2006).

Thus, transgenic expression of phosphorylation site mutants produces SPRs with several salient features. S → A rods have longer and more variable single photon-responses than WT rods. T → A rods have much longer and dramatically more variable responses, which exhibit a characteristic plateau followed by an abrupt recovery; variability in the duration of this plateau suggests that a single step dominates the time course of desensitization of T → A rhodopsin. The dependence of the kinetics and variability of the response on both serine and threonine sites indicates that they play distinct roles in controlling rhodopsin desensitization.

Desensitization of rhodopsin mutants requires phosphorylation and arrestin-1 binding

Do changes in the time course of responses produced by the mutant rhodopsins indicate desensitization through non-canonical mechanisms? As described below, a combination of biochemical and physiological experiments indicates that this is not the case.

Isoelectric focusing uses an immobilized pH gradient in a polyacrylamide gel to separate proteins according to their isoelectric point (pI), which changes with phosphorylation level (Figure 3A). We first exposed retinae to a bright 470-nm light to activate all of the rhodopsin (>98%), then froze the retinae to stop all reactions at different time points (0 min, 10 min). Immunoblots of rhodopsin from solubilized WT retina samples showed eight bands, indicating unphosphorylated rhodopsin (0), opsin (0*) plus six differently phosphorylated species, consistent with previous results (Mendez et al., 2000). Blots of both T → A and S → A samples showed five bands, corresponding to opsin (0*) and unphosphorylated (0), singly (1), doubly (2), and triply (3) phosphorylated rhodopsin, showing that T → A and S → A mutants can be phosphorylated at all sites (Figure 3A,B, Mendez et al., 2000). Quantification of the intensity of the bands showed that the T → A mutant had higher levels of doubly and triply phosphorylated rhodopsin than the S → A mutants (Figure 3B, Mendez et al., 2000), a result consistent with previous observations that serine sites are more rapidly and efficiently phosphorylated than threonine sites (Ohguro et al., 1993, 1994, 1995; Kennedy et al., 2001).

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In addition to the finding that mutant rhodopsins can be phosphorylated, arrestin-1 was able to bind to T → A and S → A rhodopsin in purified outer segment disk membranes that had been phosphorylated by recombinant GRK1 (Figure 3C). This finding is also consistent with less direct assays of bovine arrestin-1 binding to rhodopsin mutants expressed and isolated from cultured cells (Brannock et al., 1999). As expected from the biochemical assay, the expression of mutant receptors on an arrestin-1 knockout background (Arr^−/−) prolonged responses (Figure 3D); the similarity of these prolonged responses to those of Rho/Arr^−/− rods (Xu et al., 1997) indicates that termination of the mutant activity requires arrestin-1.

Response differences are not due to alterations in the transduction cascade

Transgenic protein expression can generate compensatory changes in rod outer segments, such as changes in the concentration of other transduction cascade components or changes in outer segment volume or disk size. Such changes can in turn alter response kinetics (Calvert et al., 2001; Wen et al., 2009; Makino et al., 2012). Several analyses, however, indicate that the differences in SPRs illustrated in Figures 1, 2 are attributable to the mutations in rhodopsin rather than other factors. We consider first S → A and then T → A rods.

Alterations in rhodopsin expression level affect the ability of a rod to absorb incident light. Such changes can be measured from the ‘collecting area’, which relates the number of photoactivated rhodopsin molecules produced by a light flash to the photon density of the flash (Baylor et al., 1979). The collecting area is proportional to the total amount of rhodopsin, that is, the product of concentration and volume. We measured the collecting area of S → A rods from the probability that a flash of a known photon density failed to produce a response (Figure 4A; Baylor et al., 1979). The collecting area for S → A rods was 0.63 ± 0.04 μm² or 1.3× the collecting area of WT rods (0.5 μm², Azevedo and Rieke, 2011).

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A larger collecting area could be due to an increase in rod volume, rhodopsin concentration, or both. 3D reconstruction of rods from serial electron micrographs showed that transgenic expression of T → A and S → A rhodopsin changed the size of rod outer segments (Figure 4D, Figure 4—figure supplement 1). The volume of S → A rods was 1.3× larger than WT rods (T → A rods had ∼0.75× the volume). Thus, S → A rods express ∼1.3× as much rhodopsin as WT rods, at the same concentration. A quantitatively similar increase in rod volume has been shown previously to slow the SPR, though not to the extent that S → A SPRs are slowed (Calvert et al., 2001; Wen et al., 2009; Makino et al., 2012). Thus, differences between S → A and WT responses cannot be explained by changes in volume or rhodopsin expression level. As described below (Figure 8—figure supplement 1), analysis of the kinetics of response recovery indicates minimal changes in the transduction cascade of S → A rods. Together, these results indicate that differences in S → A and WT responses are due to alterations in the time course of activity of S → A compared to WT rhodopsin.

The differences between Rho and S → A responses here are more subtle than those previously reported (Mendez et al., 2000). Rod responses, however, depend substantially on recording conditions (Azevedo and Rieke, 2011); indeed, the differences between Rho and S → A were much closer to those observed previously when we replicated the recording conditions used in earlier work (Figure 4—figure supplement 2). The rationale for the choice of recording conditions that we used for the bulk of the work here is described in Azevedo and Rieke (2011).

To check for compensatory factors in T → A rods, Rho and T → A rhodopsin were expressed in the same rod (T → A/Rho^+/−, called ‘Combo’ rods). Analogous experiments using S → A/Rho^+/− rods were not possible as crossing S → A/Rho^−/− with either WT or T → A/Rho^−/− mice resulted in very few visible rods, likely because of rhodopsin overexpression. Combo rods produced both short- and long-lasting SPRs (Figure 5A). Several aspects of these responses could be explained as a mixture of unaltered responses from WT and T → A rhodopsin expressed in roughly equal numbers. (1) The normalized average response of the Combo rods could be fit by a weighted sum of the Rho (51%) and T → A responses (49%) (Figure 5B), indicating that a near-equal mixture of the pure responses can explain the time course of the Combo rod responses. (2) The distribution of response durations from the Combo rods (gray crosses in Figure 5C) was best fit by an equal mixture of the duration distributions from Rho and T → A rhodopsin (black line in Figure 5C). (3) The kinetics of identified T → A-like and Rho-like responses from Combo rods agreed closely with the kinetics of the pure responses (Figure 5D,E; see Figure 5—figure supplement 1 and ‘Materials and methods’ for response identification).

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In summary, Figure 5 shows that T → A rhodopsin produces responses with similar kinetics in either T → A or Combo rods, and WT rhodopsin produces responses with similar kinetics in either WT or Combo rods. These findings indicate that WT, T → A, and Combo rods have similar transduction cascade kinetics, and thus, that replacement of threonines causes the prolonged SPRs in T → A rods.

Rhodopsin C-terminal threonine residues are essential for rapid arrestin-1 binding

The experiments described above indicate that the deletion of serine or threonine phosphorylation sites differentially affects rhodopsin desensitization, and that these effects cannot be attributed to either non-canonical desensitization pathways or compensatory changes in the transduction cascade. We next probed the mechanisms responsible for the slowed response kinetics in S → A rods and the abrupt cessation of T → A responses.

We hypothesized that the step-like T → A responses are terminated by the sudden desensitization of the GPCR via either abrupt phosphorylation or arrestin-1 binding. Phosphorylation consists of GRK1 binding and phosphate attachment, both of which can contribute to the rate of desensitization (Figure 1A; Doan et al., 2009). Arrestin-1 serves both to quench rhodopsin activity and to modify the GRK1-binding rate by competing with GRK1 (Doan et al., 2009). The long duration of the T → A responses suggests that GRK1 binding, phosphate attachment, or arrestin-1 binding is substantially slowed and limits the desensitization rate. We distinguished among these possibilities in both T → A and S → A rods by lowering separately the GRK1 or arrestin-1 concentration. If GRK1 binding is rate limiting, lowering the GRK1 but not the arrestin-1 concentration should further slow the responses. If arrestin-1 binding is rate limiting, lowering the arrestin-1 but not the GRK1 concentration should slow the responses. If phosphate attachment by bound GRK1 is rate limiting, the responses should be insensitive to lowering GRK1 and arrestin-1 concentrations.

We genetically lowered the concentrations of arrestin-1 or GRK1 in S → A rods by crossing S → A mice with either GRK1^−/−/Rho^−/− mice or Arr1^−/−/Rho^−/− mice to produce heterozygous strains that express less GRK1 or arrestin-1. Western blots confirmed reduced concentrations of arrestin-1 and GRK1 (data not shown). Reduced arrestin-1 expression had little or no effect on S → A SPR kinetics (Figure 6A,B, cyan trace) or durations (Figure 6C, cyan crosses). Reduced GRK1 expression similarly did not noticeably change SPR kinetics or durations (Figure 6D–F). This result differs from reduced GRK1 expression in Rho rods (GRK1^+/−/Rho^+/+), which clearly exhibit slowed responses (Doan et al., 2009; Chen et al., 2010). The smaller effect in S → A rods suggests that phosphate attachment by bound GRK1 is much slower than GRK1 binding in these rods—likely a consequence of having only threonine residues remaining (see ‘Discussion’). Slow phosphorylation of threonine residues is consistent with previous biochemical findings that serine sites are preferentially phosphorylated (Ohguro et al., 1993, 1994, 1995; Kennedy et al., 2001) and with the relatively low levels of doubly and triply phosphorylated rhodopsin in S → A isoelectric-focusing measurements (Mendez et al., 2000, Figure 3A).

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Reduced expression of arrestin-1 in T → A rods led to SPRs that qualitatively resemble those with native arrestin-1 concentrations, that is, a plateau of variable length, followed by a rapid recovery to baseline (Figure 7A). The trajectory of the abrupt recovery from the plateau was similar to that of T → A rods, suggesting that the cascade downstream of rhodopsin was unchanged (data not shown). Reducing the arrestin-1 concentration, however, slowed the kinetics of the average SPR (Figure 7B, cyan trace). Correspondingly, the duration distribution in T → A/Arr1^+/− rods shifted to longer times (Figure 7C): the median ΔT (50% point on the y-axis) increased by a factor of ∼1.5, from 2.7 s to 4.3 s (Figure 7C,G). Thus, desensitization of T → A rhodopsin is slowed by reducing the arrestin-1 concentration.

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Slow phosphorylation of the remaining serine residues by GRK1 could also contribute to the duration of the T → A responses. Reducing the GRK1 concentration, however, produced modest changes in the average SPR (Figure 7E) and response duration (Figure 7F). The cumulative probability distribution of T → A/GRK1^+/− response durations (Figure 7F) shows fewer short responses (<1 s), which accounts for the steepening of the curve. This suggests that GRK1 binding is slowed, as predicted by the results of previous experiments with GRK1^+/− mice (Doan et al., 2009). Long responses, and thus the median duration, are little affected (Figure 7F,G). The sensitivity of T → A responses to reducing arrestin-1 and the relative insensitivity to reducing GRK1 implicates arrestin-1 binding as the slow step in desensitization of T → A rhodopsin. This in turn suggests that threonine residues are required for timely rhodopsin desensitization.

Rhodopsin desensitization and the transduction cascade have similar kinetics

The kinetics of rhodopsin desensitization relative to those of downstream events in the transduction cascade is an important factor shaping SPRs. However, separating the two is difficult (Pepperberg et al., 1992; Rieke and Baylor, 1998; Krispel et al., 2006; Burns and Pugh, 2010; Gross and Burns, 2010). In one hypothetical regime, rhodopsin activates a collection of G proteins and then quickly desensitizes. In this case, the kinetics of the response recovery reflects the rate at which the active G protein population decays. In another hypothetical regime, rhodopsin desensitizes slowly compared to the rate of G protein decay such that the kinetics of the response recovery reflects the rate at which rhodopsin desensitizes. More generally, the response recovery kinetics reflect both G protein decay and rhodopsin desensitization. Indeed, the analysis described below indicates that the kinetics of brief SPRs are shaped almost entirely by the transduction cascade, while longer-lasting responses also reflect the kinetics of rhodopsin desensitization.

We grouped WT, T → A, and Combo rod responses according to their measured durations (Figure 5C), binning them into 1-s wide bins centered every 0.5 s (e.g., responses with durations <1 s, durations between 0.5 s and 1.5 s, etc). We then aligned the binned responses either to their termination points (Figure 8A) or to the time of the flash (Figure 8C) and measured the time constants of decay of the average responses in each bin. We plot the termination-aligned decay time constants (τ_term; Figure 8B) and flash-aligned decay time constants (τ_flash; Figure 8D) as functions of the response duration for WT, T → A, and Combo responses. The Combo responses, as might be expected from Figure 5, are consistent with a combination of WT and T → A responses.

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This analysis highlights several response properties. First, the fastest response decay has a time constant ∼200 ms, similar to previous measurements of the rate of the G protein's GTPase activity (Krispel et al., 2006). Three types of responses share this fast decay: (1) all termination-aligned responses (Figure 8B); (2) the briefest flash-aligned responses (Figure 8C; 0.5 s bin); and (3) the longest flash-aligned responses (Figure 8C; 4 s bin). Flash-aligned traces of intermediate kinetics have substantially slower decay time constants (Figure 8D, τ_flash; bins between 1 s and 3 s). This effect was consistent for responses of Rho and T → A rhodopsin, including when both were in the same cell.

Termination-aligned S → A responses also exhibited recovery time constants near 200 ms (Figure 8—figure supplement 1), matching those of termination-aligned responses from Rho and T → A rhodopsin. The similarity of these time constant supports our conclusion that S → A transduction is normal.

The analysis of Figure 8 suggests that G protein decay dominates the kinetics of the fastest WT responses, when, stochastically, rhodopsin is short-lived. The slower recovery of flash-aligned, intermediate-duration WT responses is consistent with a combination of slower rhodopsin desensitization and G protein decay. This effect is clear in the T → A responses (Figure 1G), where the slow decay of the average response reflects the long and variable response plateau; the analysis here indicates a similar picture that applies to WT responses.

To noticeably impact the recovery kinetics, rhodopsin must desensitize with comparable or slower kinetics than G protein decay. Consistent with this suggestion, the dashed lines in Figure 8B,D plot the decay time constant when all WT SPRs are averaged, the typical measure of the recovery time constant. The time constant of this average response is considerably slower than the fastest recovery time constants of the binned responses. Thus, our data support a model in which stochastic rhodopsin desensitization occasionally occurs quickly, such that the transduction cascade kinetics dominate the response kinetics, but desensitization often occurs more slowly and influences response kinetics.

The importance of phosphorylation and arrestin binding for GPCR desensitization has been appreciated for several decades (Stadel et al., 1982, 1983; Wilden and Kühn, 1982; Kühn et al., 1984; Maeda, 2003). Nonetheless, we lack a clear understanding of how these mechanisms support the observed diversity in GPCR signaling. Here, we investigated the role of different C-terminus phosphorylation sites in rhodopsin desensitization. We found that phosphorylation of native serines is rapid but ineffective in promoting rapid arrestin-1 binding. Phosphorylation of native threonines is slow but effective in promoting rapid arrestin-1 binding. Together, these properties ensure not only that several phosphorylation steps will complete before arrestin-1 will bind but also that after these steps complete, arrestin-1 will bind rapidly. These results challenge assumptions about the relative importance of serine and threonine sites and show how phosphorylation and arrestin binding can act in concert to tightly control the timing of rhodopsin desensitization.

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Author details

1. Anthony W Azevedo

   Department of Physiology and Biophysics, University of Washington, Seattle, United States

   Contribution

   AWA, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Thuy Doan

   Department of Ophthalmology, University of Washington, Seattle, United States

   Contribution

   TD, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

3. Hormoz Moaven

   Departments of Cell & Neurobiology and Ophthalmology, Zilkha Neurogenetic Institute, Keck School of Medicine of University of Southern California, Los Angeles, United States

   Contribution

   HM, Conception and design, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

4. Iza Sokal

   Department of Physiology and Biophysics, University of Washington, Seattle, United States

   Contribution

   IS, Isolated rhodopsin outer segments, Acquisition of data

   Competing interests

   The authors declare that no competing interests exist.

5. Faiza Baameur

   Department of Pharmacology, Vanderbilt University School of Medicine, Nashville, United States

   Contribution

   FB, Conception and design, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

6. Sergey A Vishnivetskiy

   Department of Pharmacology, Vanderbilt University School of Medicine, Nashville, United States

   Contribution

   SAV, Conception and design, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

7. Kristoff T Homan

   Life Sciences Institute, Departments of Pharmacology and Biological Chemistry, University of Michigan, Ann Arbor, United States

   Contribution

   KTH, Expressed and purified GRK1, Acquisition of data

   Competing interests

   The authors declare that no competing interests exist.

8. John JG Tesmer

   Life Sciences Institute, Departments of Pharmacology and Biological Chemistry, University of Michigan, Ann Arbor, United States

   Contribution

   JJGT, Expressed and purified GRK1, Acquisition of data

   Competing interests

   The authors declare that no competing interests exist.

9. Vsevolod V Gurevich

   Department of Pharmacology, Vanderbilt University School of Medicine, Nashville, United States

   Contribution

   VVG, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-3950-5351

10. Jeannie Chen

    Departments of Cell & Neurobiology and Ophthalmology, Zilkha Neurogenetic Institute, Keck School of Medicine of University of Southern California, Los Angeles, United States

    Contribution

    JC, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

    Competing interests

    The authors declare that no competing interests exist.

11. Fred Rieke

    1. Department of Physiology and Biophysics, University of Washington, Seattle, United States
    2. Howard Hughes Medical Institute, University of Washington, Seattle, United States

    Contribution

    FR, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    rieke@u.washington.edu

    Competing interests

    The authors declare that no competing interests exist.

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