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Functional Characterization of Phytochrome Autophosphorylation in Plant Light Signaling

Functional Characterization of Phytochrome Autophosphorylation in Plant Light Signaling Abstract Plant phytochromes, molecular light switches that regulate various aspects of plant growth and development, are phosphoproteins that are also known to be autophosphorylating serine/threonine kinases. Although a few protein phosphatases that directly interact with and dephosphorylate phytochromes have been identified, no protein kinase that acts on phytochromes has been reported thus far, and the exact site of phytochrome autophosphorylation has not been identified. In this study, we investigated the functional role of phytochrome autophosphorylation. We first mapped precisely the autophosphorylation sites of oat phytochrome A (phyA), and identified Ser8 and Ser18 in the 65 amino acid N-terminal extension (NTE) region as being the autophosphorylation sites. The in vivo functional roles of phytochrome autophosphorylation were examined by introducing autophosphorylation site mutants into phyA-deficient Arabidopsis thaliana. We found that all the transgenic plants expressing the autophosphorylation site mutants exhibited hypersensitive light responses, indicating an increase in phyA activity. Further analysis showed that these phyA mutant proteins were degraded at a significantly slower rate than wild-type phyA under light conditions, which suggests that the increased phyA activity of the mutants is related to their increased protein stability. In addition, protoplast transfection analyses with green fluorescent protein (GFP)-fused phyA constructs showed that the autophosphorylation site mutants formed sequestered areas of phytochrome (SAPs) in the cytosol much more slowly than did wild-type phyA. These results suggest that the autophosphorylation of phyA plays an important role in the regulation of plant phytochrome signaling through the control of phyA protein stability. Introduction Phytochromes are red/far-red (R/FR) photoreceptors that regulate many aspects of plant growth and development in response to informational light signals from the environment (Rockwell et al. 2006, Bae and Choi 2008). They are dimeric chromoproteins with covalently linked tetrapyrrole chromophore phytochromobilin, and exist in two photointerconvertible species, red-light-absorbing Pr and FR-light absorbing Pfr forms. Phytochromes are biosynthesized as the Pr form in the dark, which can be phototransformed into the Pfr form upon exposure to red light. This photoactivation of phytochromes from the Pr form to the Pfr form induces the highly regulated signaling network for photomorphogenesis in plants (Chen et al. 2004, Han et al. 2007). Recent studies on phytochrome-mediated light signaling revealed that phytochrome photoactivation has a major impact on the control of protein subcellular localization (Nagatani 2004, Fankhauser and Chen 2008), transcription (Jiao et al. 2007), protein stability (Henriques et al. 2009) and protein phosphorylation (Kim et al. 2005). However, despite intensive studies on phytochromes involving a broad range of experimental approaches since their first discovery in the 1950s, the exact biochemical mechanisms underlying the phytochrome regulation of photoresponses in plants have not been fully elucidated. Phytochromes are known to be phosphoproteins by phosphate analysis on purified phytochrome preparations (Hunt and Pratt 1980), and the sites of phytochrome phosphorylation have been investigated with purified phytochrome A (phyA) from oat seedlings (Lapko et al. 1996, Lapko et al. 1997, Lapko et al. 1999). There are two phosphorylation sites (Ser8 and Ser18) in the N-terminal extension (NTE) region of the phyA molecule, and one site (Ser599) in the hinge region between the N- and C-terminal domains. In previous reports, the NTE was shown to be required for full biological activity of phyA (Cherry et al. 1992), and serine to alanine substitution in the NTE region including Ser8 and Ser18 has resulted in increased biological activity, showing that transgenic plants expressing this mutant form of phyA were hypersensitive to light and had a dwarf phenotype (Stockhaus et al. 1992). These results suggest that phosphorylation in the NTE region is a mechanism of signal attenuation or desensitization of phytochrome signal transduction (Emmler et al. 1995, Jordan et al. 1996, Jordan et al. 1997, Casal et al. 2002). However, it remains unclear how this substitution generates an attenuation signal. In addition, functional characterization has shown that phosphorylation at Ser599 prevents the interaction of phyA with its signal transducers such as nucleoside di-phosphate kinase 2 (NDPK2) and phytochrome-interacting factor 3 (PIF3), which suggests that the hinge region of phytochromes serves as a phosphorylatable signal-modulating site that regulates protein–protein interactions between phytochrome and its signal transducers (Kim et al. 2004). These data suggest the importance of phytochrome phosphorylation in the light signaling activities of phytochromes. The observation that phytochromes are phosphoproteins suggests the existence of protein kinase(s) that phosphorylate the phytochromes and protein phosphatase(s) that dephosphorylate them. However, despite extensive studies of phytochrome-interacting proteins, there is no report thus far of a protein kinase that can phosphorylate phytochromes. On the other hand, a few protein phosphatases have been reported as being able to interact with and dephosphorylate phytochromes, including flower-specific phytochrome-associated protein phosphatase (FyPP) (Kim et al. 2002), phytochrome-associated protein phosphatase 5 (PAPP5) (Ryu et al. 2005) and phytochrome-associated protein phosphatase type 2C (PAPP2C) (Phee et al., 2008). The FyPP-overexpressing transgenic plants had enhanced phytochrome activity during flowering and hypocotyl shortening, whereas the antisense repression of FyPP resulted in transgenic plants with reduced phytochrome activity (Kim et al. 2002). PAPP5, which is involved in the regulation of de-etiolation, positively influences the protein stability of phytochrome and the interaction of phytochrome with a downstream transducer NDPK2 (Ryu et al. 2005). PAPP2C also positively regulates the light responses of plants (Phee et al. 2008). These observations suggest that phytochrome phosphorylation and dephosphorylation play important roles in phytochrome-mediated light signaling. Since no protein kinase is known to act on phytochromes, it is possible that phytochromes are phosphorylated by means of autophosphorylation because phytochromes are known to be autophosphorylating serine/threonine protein kinases (Yeh and Lagarias 1998). Because phytochrome phosphorylation and dephosphorylation have been known to play an important role in the regulation of phytochrome functions (Kim et al. 2005), phytochrome phosphorylation would be controlled by autophosphorylation and protein phosphatases. However, the in vivo functional role of phytochrome phosphorylation remains to be explored. Thus, several important aspects of the role of phosphorylation in phytochrome signaling remain to be resolved, including the determination of autophosphorylation site(s) and the in vivo functional role of phytochrome autophosphorylation. In this study, we determined the autophosphorylation sites of oat phyA and investigated the functional role of phyA autophosphorylation using phyA-deficient Arabidopsis plants transformed with oat phyA autophosphorylation site mutants. We demonstrated that oat phyA was autophosphorylated on Ser8 and Ser18 in the NTE region and that the transgenic plants of autophosphorylation site-deleted mutants exhibited hypersensitive phenotypes in response to FR light, indicating that the mutants are hyperactive in plants. Further studies demonstrated that the phyA mutant proteins degraded slowly compared with wild-type phyA, which could account for the enhanced photoresponse of transgenic plants bearing the phyA mutant. Therefore, our results provide direct evidence for the functional role of phyA autophosphorylation. Results Oat phyA contains autophosphorylation activity that is stimulated by the addition of histone H1 Before investigating the phytochrome autophosphorylation site(s), we first examined the autophosphorylation of purified recombinant oat phyA. Since the expression of full-length plant phytochromes in a recombinant system is known to be very difficult, we expressed and purified recombinant phyA proteins using the Pichia protein expression system and streptavidin affinity chromatography, as we previously reported (Kim et al. 2004). Spectroscopic analyses of the purified serine to alanine phyA mutants, namely S8A, S18A and S8/18A, showed that the absorbance and R/FR difference spectra of the mutants were identical to those of wild-type phyA (Supplementary Table S1 and Supplementary Data), indicating that the protein integrity of the mutants is similar to that of the wild type. We then performed phytochrome autophosphorylation experiments with the purified phyA proteins. The phyA protein is known to be autophosphorylated, and the autophosphorylation is stimulated in the presence of polycations, such as histone H1 (Wong et al. 1986, Wong et al. 1989, Yeh and Lagarias 1998). Thus, we investigated the autophosphorylation of purified recombinant phyA in the presence or absence of histone H1 (Fig. 1A). These experiments confirmed that phyA possessed autophosphorylation activity that was stimulated by histone H1. Zinc fluorescence and SDS–PAGE also confirmed the formation of chromophore-ligated phyA proteins and the protein purity used in the reactions, respectively. Under our experimental conditions, Pr and Pfr forms of oat phyA were similarly autophosphorylated, and the addition of histone H1 preferentially stimulated phyA autophosphorylation of the Pr form, consistent with findings of a previous report (Yeh and Lagarias 1998). When the autoradiogram bands were quantified with a densitometer (Molecular Dynamics), the intensities of autophosphorylated Pr and Pfr forms of phyA in the presence of histone H1 were 3.5 and 1.7, respectively, with the Pr intensity in the absence of histone H1 being set at 1.0. In addition, histone H1 was phosphorylated by phyA, indicating the phosphotransfer activity of phyA. Unlike Pr-preferential stimulated autophosphorylation of phyA, histone H1 phosphorylation was not light dependent. Fig. 1 View largeDownload slide Autophosphorylation analysis of recombinant oat phyA. (A) Autophosphorylation of purified recombinant oat phyA in the absence or presence of histone H1. FL-phyA, full-length wild-type oat phyA; H1, histone H1; S, protein size standards. Autoradiogram (Autorad), zinc fluorescence (Zinc) and SDS–PAGE are shown. A 1.5 μg aliquot of the Pr or Pfr form of phyA was used in the reactions with or without 0.75 μg of histone H1. (B) Time-dependent autophosphorylation of the Pr and Pfr forms of oat phyA in the absence or presence of histone H1. In these reactions, 4 pmol of the Pr or Pfr form of phyA was used with or without 2 μg of histone H1. (C) 32P incorporation in oat phyA by autophosphorylation shown in (B). Each analysis was repeated three times. Error bar = SD (n = 3). Fig. 1 View largeDownload slide Autophosphorylation analysis of recombinant oat phyA. (A) Autophosphorylation of purified recombinant oat phyA in the absence or presence of histone H1. FL-phyA, full-length wild-type oat phyA; H1, histone H1; S, protein size standards. Autoradiogram (Autorad), zinc fluorescence (Zinc) and SDS–PAGE are shown. A 1.5 μg aliquot of the Pr or Pfr form of phyA was used in the reactions with or without 0.75 μg of histone H1. (B) Time-dependent autophosphorylation of the Pr and Pfr forms of oat phyA in the absence or presence of histone H1. In these reactions, 4 pmol of the Pr or Pfr form of phyA was used with or without 2 μg of histone H1. (C) 32P incorporation in oat phyA by autophosphorylation shown in (B). Each analysis was repeated three times. Error bar = SD (n = 3). We further investigated phyA autophosphorylation over a time course (Fig. 1B, C). The results of these experiments showed that the phyA autophosphorylation was slightly higher in the Pr form than in the Pfr form, but the difference was not significant. In the presence of histone H1, 32P incorporation was stimulated >2-fold, in a Pr-specific manner, whereas 32P incorporation of the Pfr form increased about 1.2-fold. The analysis of these time course experiments revealed that autophosphorylation increased with time and reached a maximum value after 60 min in the absence of histone H1 and after approximately 120 min in the presence of histone H1. Therefore, our results showed that oat phyA possessed autophosphorylation activity and that the autophosphorylation could be stimulated in the presence of histone H1, and thereby confirmed phyA as an autophosphorylating protein kinase. The sites of PhyA autophosphorylation reside in the NTE region Since three phosphorylation sites were known to exist in oat phyA (Lapko et al. 1999), we sought to identify the autophosphorylation site(s) of oat phyA by generating serine to alanine mutants of the known phosphorylation sites. Previously, we reported that S599 is not the site of autophosphorylation (Kim et al. 2004), and the serine-rich NTE region has been proposed to be phosphorylated by phytochrome itself or by a phytochrome-associated kinase (McMichael and Lagarias 1990, Lapko et al. 1999, Kim et al. 2005). Thus, we first examined the autophosphorylation of NTE-deleted Δ65-phyA, in which two known phosphorylation sites, Ser8 and Ser18, were deleted (Fig. 2A). We found that phyA autophosphorylation was not detected in the NTE-deleted Δ65-phyA mutant, even in the presence of histone H1, which suggests that the autophosphorylation site(s) of phyA reside in the NTE region. Although Δ65-phyA did not exhibit any autophosphorylation, it was still able to show the phosphotransfer activity onto histone H1. This result indicates that the NTE is necessary for the phyA autophosphorylation, but not for the phyA kinase activity. Fig. 2 View largeDownload slide Determination of autophosphorylation sites in oat phyA. (A) Autophosphorylation of full-length (FL, amino acids 1–1,129) and NTE-deleted (Δ65, amino acids 66–1,129) phyA in the absence or presence of histone H1. (B) Autophosphorylation of serine to alanine mutants of three known phosphorylation sites. (C) Autophosphorylation of a double site mutant S8/18A. Fig. 2 View largeDownload slide Determination of autophosphorylation sites in oat phyA. (A) Autophosphorylation of full-length (FL, amino acids 1–1,129) and NTE-deleted (Δ65, amino acids 66–1,129) phyA in the absence or presence of histone H1. (B) Autophosphorylation of serine to alanine mutants of three known phosphorylation sites. (C) Autophosphorylation of a double site mutant S8/18A. We next investigated the autophosphorylation of the serine to alanine mutants (Fig. 2B, C). The results showed that the autophosphorylation of the S8A and S18A mutants was significantly reduced, but that the S599A mutant exhibited a similar level of autophosphorylation to wild-type phyA (Fig. 2B). When the autoradiogram bands were quantified with a phosphoimage analyzer, we found that the intensities of both autophosphorylated S8A and S18A were reduced to approximately half the levels seen in wild-type phyA. Furthermore, the S8/18A double mutant showed little autophosphorylation (Fig. 2C). Thus, our results showed that Ser8 and Ser18, which are located in the NTE, are the autophosphorylation sites of oat phyA. To our knowledge, this is the first direct determination of the exact autophosphorylation sites of phyA. Autophosphorylation of PhyA is involved in the regulation of light responses To investigate the functional role of phyA autophosphorylation, we produced homozygous lines of transgenic phyA- deficient Arabidopsis thaliana (phyA-201) plants that expressed the autophosphorylation site mutants as well as wild-type oat phyA. Since it is known that phytochrome function has strong dependency on the amounts of photoreceptor (Boylan and Quail 1991, Whitelam et al. 1993, Wagner and Quail 1995), we first selected transgenic lines overexpressing wild-type phyA whose expression level of oat phyA is similar to that of transgenic lines overexpressing autophosphorylation site mutants for the proper comparison of light responses. To do this, we performed reverse transcription–PCR (RT–PCR) and Western blot analyses to detect the transcript and protein levels in the transgenic lines overexpressing wild-type phyA (Supplementary Fig. S2). For the Western blot analysis, we included the proteasome inhibitor MG-132 in the seedling growth medium and protein extraction buffer to prevent any possible phyA protein degradation during the preparation of plant crude extracts. We found that the Wt-OX6 line showed similar transcript and protein levels of oat phyA to the transgenic lines overexpressing autophosphorylation site mutants (Fig. 3A, B). Among the transgenic plants with wild-type oat phyA, the Wt-OX47 line expressed the highest amount of phyA because of strong transcriptional expression (Supplementary Fig. S2). Thus, we used the Wt-OX6 line for the comparison of light responses with the transgenic plants of autophosphorylation site mutants in the present study. In addition, we included Wt-OX47 as a strong phyA expressor in the experiments of light responsiveness and protein degradation analysis. Fig. 3 View largeDownload slide Seedling de-etiolation responses of transgenic Arabidopsis plants expressing the autophosphorylation site mutants under FR light. (A) RT–PCR analysis to show the transcript levels of oat phyA in transgenic plants. phyA-201, phyA-deficient Arabidopsis (Ler ecotype); Ler, wild-type Arabidopsis; Wt-OX6, transgenic Arabidopsis transformed with wild-type oat phyA; S8A, S18S and S8/18A, transgenic Arabidopsis transformed with the corresponding autophosphorylation site mutant. Numbers represent independent homozygous lines. Actin (ACT2) is shown as a loading control. (B) Western blot analysis to show the protein levels of oat phyA in transgenic plants. For this analysis, seedlings were cultured on MG-132-containing media, and crude proteins were extracted with an MG-132-containing buffer. Oat phyA-specific oat25 antibody was used for the detection of oat phyA, and β-tubulin (TUB) is shown as a loading control. S, protein size standards. (C) Hypocotyl de-etiolation of representative seedlings grown under continuous FR light conditions. 0.5 and 5.0 in parenthesis are the fluence rates (μmol m−2 s−1) used in the analyses. Bar = 5.0 mm. (D) The average hypocotyl lengths of seedlings in (C). Seedlings (n ≥ 29) were grown for 4 d in 1/2 MS medium under fluence rates of 0.5 or 5.0 μmol m−2 s−1, or in darkness. Data are the means ± SD. (E) FR fluence-rate response curves for inhibition of hypocotyl growth. Data are the means (n ≥ 29) ± SD. Fig. 3 View largeDownload slide Seedling de-etiolation responses of transgenic Arabidopsis plants expressing the autophosphorylation site mutants under FR light. (A) RT–PCR analysis to show the transcript levels of oat phyA in transgenic plants. phyA-201, phyA-deficient Arabidopsis (Ler ecotype); Ler, wild-type Arabidopsis; Wt-OX6, transgenic Arabidopsis transformed with wild-type oat phyA; S8A, S18S and S8/18A, transgenic Arabidopsis transformed with the corresponding autophosphorylation site mutant. Numbers represent independent homozygous lines. Actin (ACT2) is shown as a loading control. (B) Western blot analysis to show the protein levels of oat phyA in transgenic plants. For this analysis, seedlings were cultured on MG-132-containing media, and crude proteins were extracted with an MG-132-containing buffer. Oat phyA-specific oat25 antibody was used for the detection of oat phyA, and β-tubulin (TUB) is shown as a loading control. S, protein size standards. (C) Hypocotyl de-etiolation of representative seedlings grown under continuous FR light conditions. 0.5 and 5.0 in parenthesis are the fluence rates (μmol m−2 s−1) used in the analyses. Bar = 5.0 mm. (D) The average hypocotyl lengths of seedlings in (C). Seedlings (n ≥ 29) were grown for 4 d in 1/2 MS medium under fluence rates of 0.5 or 5.0 μmol m−2 s−1, or in darkness. Data are the means ± SD. (E) FR fluence-rate response curves for inhibition of hypocotyl growth. Data are the means (n ≥ 29) ± SD. After selecting two independent homozygous lines of each autophosphorylation site mutant transgenic plant that showed a level of phyA transcript and protein expression comparable with the Wt-OX6 transgenic line, we investigated the light responses of the transgenic plants. Since phyA is known to participate exclusively in the FR-induced inhibition of hypocotyl elongation (Fankhauser and Casal 2004), we first investigated the seedling de-etiolation response of the homozygous transgenic lines in continuous FR (cFR) light with fluence rates of 0.5 and 5.0 μmol m−2 s−1, or in darkness. We measured the hypocotyl lengths of the transgenic lines and wild-type seedlings, and found that all of the transgenic lines (i.e. S8A, S18A and S8/18A) had shorter hypocotyls than non-transformed wild-type Arabidopsis (Ler), or the Wt-OX6 (Fig. 3C, D). When we compared the hypocotyl lengths of the transgenic lines with the strong phyA-overexpressing line Wt-OX47, the transgenic lines still had slightly shorter hypocotyls than Wt-OX47. In particular, the S8/S18 transgenic lines showed much shorter hypocotyls than Wt-OX47. These results suggest that the autophosphorylation site mutants of phyA were hyperactive, and therefore increased the sensitivity of the seedlings to the FR light. Since the hypocotyls of the S8/18A lines were much shorter than those of S8A and S18A, our results suggest that S8 and S18 phosphorylation might have an additive effect on the function of phyA. The observations that the autophosphorylation activity of S8A and S18A was reduced to half that of wild-type phyA, and that of S8/18A was barely detectable, suggest that there is a negative relationship between the magnitude of autophosphorylation and photoresponses in de-etiolation. Furthermore, FR fluence-rate response curves for inhibition of hypocotyl growth confirmed that transgenic seedlings of autophosphorylation site mutants were more sensitive to FR than were those of control plants (i.e. Ler or Wt-OXs) (Fig. 3E). Again, the S8/18A transgenic lines showed the greatest hypersensitive photoresponses among the tested seedlings including Wt-OX47. These results suggest that reduction of autophosphorylation augments the photoresponse in plants, and thus that phyA autophosphorylation might play a role in inhibiting the function of phyA. This is in agreement with the previous proposal that phytochrome phosphorylation is a mechanism of signal attenuation or desensitization for phytochrome- mediated light signaling (Jordan et al. 1996, Casal et al. 2002, Ryu et al. 2005). PhyA mediates a series of photoresponses to cFR, including inhibition of hypocotyl growth, unfolding of the cotyledons, accumulation of anthocyanin and blocking of subsequent greening under white light (Chen et al. 2004). PhyA also mediates two distinct photobiological responses in plants, the very low fluence responses (VLFRs) and the high irradiance responses (HIRs). The VLFR can be achieved by short intermittent pulses of FR [i.e. pulsed FR (pFR)], while the HIR involves the sustained activation of phyA in response to higher fluences of FR (Casal et al. 2002). Since the VLFRs and FR-HIRs are regulated by phyA activity in plants, we analyzed the VLFRs of transgenic seedlings by hourly treatment with FR (i.e. pFR, 24 μmol m−2 s−1 FR : dark = 5 : 55 min), and also the HIRs with continuous FR (2 μmol m−2 s−1) (Fig. 4A). The results showed that all the transgenic lines expressing autophosphorylation site mutants had increased inhibition of hypocotyl growth compared with Ler or Wt-OX6, and that the S8/18A double mutant exhibits the greatest inhibition of hypocotyl elongation in response to cFR and pFR. Therefore, phyA autophosphorylation is involved in the regulation of both VLFRs and HIRs. Fig. 4 View largeDownload slide Photoresponse analysis of transgenic Arabidopsis seedlings expressing the autophosphorylation site mutants. (A) Inhibition of hypocotyl growth under cFR or pFR (i.e. hourly FR). Four-day-old seedlings were exposed to continuous FR (2 μmol m−2 s−1) or hourly FR (24 μmol m−2 s−1 FR : dark = 5 : 55 min). Data are means (n ≥ 29) ± SD. (B) Accumulation of anthocyanin in seedlings grown in the dark, with cFR or pFR light. This assay was repeated three times. Data are the means ± SD (n = 45). (C) Expression of the light-inducible CHS gene in transgenic plants. Seedlings were cultured for 3 d in the dark and transferred to FR (10 μmol m−2 s−1) for 18 h. Total RNA was extracted and RT–PCR analyses were performed using CHS-specific primers. The relative intensities of bands (Irel) are shown beneath the gel, where the band intensity of Ler was set at 1.0. (D) Blocking of greening under FR light. The amount of chlorophyll was measured in 30 seedlings of each line. Data are means (n ≥ 30) ± SD. Seedlings were cultured in the dark, under cFR or pFR light for 3 d, and then transferred to white light (150 μmol m−2 s−1) for 1 d before analyzing the chlorophyll content. Fig. 4 View largeDownload slide Photoresponse analysis of transgenic Arabidopsis seedlings expressing the autophosphorylation site mutants. (A) Inhibition of hypocotyl growth under cFR or pFR (i.e. hourly FR). Four-day-old seedlings were exposed to continuous FR (2 μmol m−2 s−1) or hourly FR (24 μmol m−2 s−1 FR : dark = 5 : 55 min). Data are means (n ≥ 29) ± SD. (B) Accumulation of anthocyanin in seedlings grown in the dark, with cFR or pFR light. This assay was repeated three times. Data are the means ± SD (n = 45). (C) Expression of the light-inducible CHS gene in transgenic plants. Seedlings were cultured for 3 d in the dark and transferred to FR (10 μmol m−2 s−1) for 18 h. Total RNA was extracted and RT–PCR analyses were performed using CHS-specific primers. The relative intensities of bands (Irel) are shown beneath the gel, where the band intensity of Ler was set at 1.0. (D) Blocking of greening under FR light. The amount of chlorophyll was measured in 30 seedlings of each line. Data are means (n ≥ 30) ± SD. Seedlings were cultured in the dark, under cFR or pFR light for 3 d, and then transferred to white light (150 μmol m−2 s−1) for 1 d before analyzing the chlorophyll content. Accumulation of anthocyanins in response to light is also mediated by phyA, which up-regulates the expression of the chalcone synthase gene (CHS) upon irradiation with FR light. We investigated the relative amounts of anthocyanin in seedlings cultured for 3 d under dark, cFR and pulsed FR (FR : dark = 5 : 55 min), and found that anthocyanin accumulation was elevated in response to both cFR and pFR in the transgenic lines expressing autophosphorylation site mutants (Fig. 4B). The extent of induction of CHS was positively correlated with the results of anthocyanin accumulation (Fig. 4C). Furthermore, the results of another HIR to phyA, the blocking of greening under white light, showed that the autophosphorylation site mutants inhibited white light-induced greening significantly more than did Ler or Wt-OX6 in the same light conditions (Fig. 4D). Together, these results suggest that the autophosphorylation site mutants are hyperactive in plants. In addition, the S8/18A mutant exhibited a synergistic increase in the intensity of light responses compared with S8A or S18A. Thus, phosphorylation of both S8 and S18 sites is important for mediating phyA-regulated light responses, including VLFR and FR-HIR. The increased phyA activity in these mutants is related to the decreased extent of phyA autophosphorylation. Overall, our results indicate that phyA autophosphorylation plays an inhibitory role in phytochrome signaling. Autophosphorylation controls the stability of the PhyA protein Previous studies with PAPP5, a phytochrome-interacting protein phosphatase in Arabidopsis, showed that phytochrome stability is increased in PAPP5-overexpression lines, and decreased in papp5 knockout lines, suggesting that phytochrome phosphorylation is involved in regulating the stability of the phytochrome (Ryu et al. 2005). This indicates that dephosphorylated phytochrome is more stable than phosphorylated phytochrome. Furthermore, since phosphorylation at Ser599 does not affect the stability of phyA (Kim et al. 2004), phosphorylation at Ser8 and Ser18 in the NTE have been proposed to be important for protein stability (Kim et al. 2005). Thus, we investigated whether protein stability of the autophosphorylation site mutants was increased by analyzing in vivo protein degradation of phyA proteins upon red light irradiation (28 μmol m−2 s−1) in a light-emitting diode (LED) growth chamber. We found that the autophosphorylation site mutants were more stable than wild-type phyA (Fig. 5). The phyA protein is known to be degraded quickly upon red light irradiation (Henriques et al. 2009), and our results showed that most of the wild-type phyA proteins were degraded within 1 h of red light irradiation, even in the strong phyA-overexpressing Wt-OX47 line. However, the autophosphorylation mutant phyA proteins were degraded much more slowly, and remained even after 3 h of red light irradiation. In addition, S8/18A phyA proteins were more stable than S8A and S18A, indicating the close relationship between autophosphorylation and protein stability of phyA. Interestingly, during our analysis of the amount of phyA in seedlings, we found that the autophosphorylation mutant phyA proteins were not completely degraded in seedlings grown even in white light conditions (Supplementary Fig. S3). We examined the phyA protein levels in dark-grown or white light-grown transgenic seedlings, and found that the autophosphorylation mutant phyA was not completely degraded in seedlings grown for 2 weeks under white light conditions in that wild-type phyA was not detected. Taken together, the loss of autophosphorylation appears to increase the protein stability of phyA, even under conditions of prolonged light irradiation. Fig. 5 View largeDownload slide PhyA protein stability in transgenic plants expressing the autophosphorylation site mutants. (A) Time-dependent degradation of phyA proteins. Three-day-old dark-grown seedlings of transgenic plants were irradiated with continuous red light (28 μmol m−2 s−1) for the indicated periods of time. A 40 μg aliquot of crude extract was loaded onto SDS–polyacrylamide gels and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. (B) Protein degradation graphs of phyA shown in (A). The time-dependent degradation experiments were performed three times, and the averages are shown with the SD. Fig. 5 View largeDownload slide PhyA protein stability in transgenic plants expressing the autophosphorylation site mutants. (A) Time-dependent degradation of phyA proteins. Three-day-old dark-grown seedlings of transgenic plants were irradiated with continuous red light (28 μmol m−2 s−1) for the indicated periods of time. A 40 μg aliquot of crude extract was loaded onto SDS–polyacrylamide gels and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. (B) Protein degradation graphs of phyA shown in (A). The time-dependent degradation experiments were performed three times, and the averages are shown with the SD. To examine further the stability of the phyA protein, green fluorescent protein (GFP)-fused phyA constructs of autophosphorylation site mutants were transiently expressed in mesophyll cell protoplasts, and degradation of GFP-fused phyA was examined under red light irradiation. During light-induced degradation of phyA, phyA exhibits rapid light-dependent aggregation, forming sequestered areas of phytochrome (SAPs) before its degradation via the ubiquitin–26S proteasome pathway (Eichenberg et al. 1999). We also observed the formation of SAPs upon red light irradiation (Fig. 6). In the case of wild-type phyA, SAPs were observed even in the dark, and their size transiently increased and then rapidly disappeared upon red light irradiation. On the other hand, the formation of SAPs was delayed in the autophosphorylation mutants, and they disappeared more slowly than those derived from wild-type phyA. In particular, S8/18A generated SAPs much more slowly than the other phyA mutants, suggesting that the retarded degradation of autophosphorylation mutant phyA proteins in the plant cells is due to the delayed formation of SAPs. Collectively, our results demonstrate that phyA autophosphorylation plays an important role in the regulation of plant phytochrome signaling via the control of phyA protein stability. Fig. 6 View largeDownload slide Analysis of formation of SAPs in mesophyll protoplasts transfected with the autophosphorylation site mutants. GFP-fused phyA constructs were transfected into Arabidopsis mesophyll protoplasts, and phyA proteins were detected with a Leica TCS SP5 confocal microscope upon red light (10 μmol m−2 s−1) irradiation for the indicated periods of time. Bar = 10 μm. Fig. 6 View largeDownload slide Analysis of formation of SAPs in mesophyll protoplasts transfected with the autophosphorylation site mutants. GFP-fused phyA constructs were transfected into Arabidopsis mesophyll protoplasts, and phyA proteins were detected with a Leica TCS SP5 confocal microscope upon red light (10 μmol m−2 s−1) irradiation for the indicated periods of time. Bar = 10 μm. Discussion Autophosphorylation sites of oat PhyA The phytochrome molecule is known to consist of two structural domains, the globular N-terminal chromophore-binding domain (∼65 kDa) and the conformationally open or extended C-terminal domain (∼55 kDa) (Rockwell et al. 2006). The two domains are connected via a flexible hinge region. The N-terminal domain is necessary and sufficient for photoperception and possesses a few subdomains, including the NTE and a bilin lyase domain, while the C-terminal domain contains Per-Arnt-Sim (PAS)-related domains and a histidine kinase-related domain (HKRD), and is necessary for phytochrome dimerization and nuclear localization (Kim et al. 2005, Han et al. 2007). The present study shows that the autophosphorylation sites of phyA reside in the NTE (Fig. 2). The first ∼65 amino acids of the protein (i.e. the NTE) are known to be dispensable for chromophore binding, but necessary for biological activity (Cherry et al. 1992, Jordan et al. 1997). In addition, serine to alanine substitutions in the first 10 serine residues (including Ser8 and Ser18) of rice phyA and deletion of amino acids 6–12 (including Ser8) of oat phyA produce hyperactive phyA, which has higher biological activity than the corresponding wild-type phyA, suggesting that phosphorylation in this region is involved in the down-regulation of phyA activity, such as desensitization (Stockhaus et al. 1992, Jordan et al. 1996). Therefore, our results of the autophosphorylation sites in the NTE are in good agreement with these previous reports. When we compared the autophosphorylation between the Pr and Pfr forms in the absence of histone H1, we could not detect any significant difference between the two forms. However, the Pr form is slightly more phosphorylated than the Pfr form (Fig. 1A). These results are somewhat different from those of a previous report, in that phyA is autophosphorylated in a Pfr-preferential manner (Yeh and Lagarias 1998). This discrepancy is probably due to the low protein kinase activity of phyA compared with other known kinases, such as protein kinase A. This weak autophosphorylation activity of phytochromes may account for the difference between our results and previous results, and may also underlie the long controversy regarding whether or not phytochrome is a protein kinase (Fankhauser 2000, Kim et al. 2005). The weak protein kinase activity of phyA might also indicate the existence of phyA kinase-stimulating molecules, such as polycationic proteins, which include histones. Indeed, phytochrome autophosphorylation and kinase activity are stimulated by the addition of histone H1 (Fig. 1B). Interestingly, the Pr form of phyA, which is considered as the biologically inactive form of phytochrome, is preferentially stimulated by histone H1. Previous studies also reported Pr-specific phytochrome phosphorylation (McMichael and Lagarias 1990, Biermann et al. 1994). Therefore, phytochrome kinase activity might be a Pr-specific kinase that could be stimulated by other factors such as polycationic proteins including histone H1. However, the significance of the stimulation of phytochrome kinase activity in the presence of polycationic proteins needs to be investigated in the future. In addition, we still do not know about the kinase domain of phytochrome, although phytochrome exhibits autophosphorylation and phosphotransfer activities. In an earlier study of affinity labeling of oat phytochrome with ATP analogs, two polypeptide sequences, Glu-Leu-Glu-Lys-Gln-Leu-Arg-Glu-Lys-Asn-Ile-Leu-Lys (residues 403–415) and Asp-Leu-Lys-Leu-Asp-Gly-Leu-Ala (residues 606–613), were suggested to be similar to peptide sequences found within the nucleotide-binding sites of known protein kinases (Wong and Lagarias 1989). In the C-terminal domain of phytochrome, a HKRD exists, but it has been suggested that this HKRD is not a functional kinase domain because the key conserved residues within the histidine kinase domain (HKD) are absent in the phytochrome HKRD (Quail 1997). A recent study of the structure of cyanobacterial phytochrome revealed that the structure of the N-terminal domain is similar to adenylyl cyclase, consisting of PAS (Per-Arnt-Sim)/GAF (cGMP phosphodiesterase/adenylyl cyclase/FhlA)/GAF domains (Essen et al. 2008). Together with an earlier study (Wong and Lagarias 1989), this result suggest that the N-terminal domain of phytochrome might be able to bind to cAMP or possibly to ATP. However, further studies are necessary to determine the phytochrome kinase domain and the catalytic sites. From the present studies, the autophosphorylation sites have been clearly determined as being Ser8 and Ser18 in the NTE (Fig. 2). The sites determined are consistent with the previous results of serine to alanine mutants in the NTE and the studies of in vivo and in vitro phosphorylation sites of phyA (Stockhaus et al. 1992, Lapko et al. 1999). There are three known phosphorylation sites on oat phyA. When we analyzed the amino acid sequences of the sites, a pattern of amino acids, –RXXS–, can be found from Ser8 (–RPAS–), Ser18 (–RQSS–) and Ser599 (–REAS–), in which the phosphorylated serines are underlined. Therefore, the phosphorylation sites of oat phyA contain a conserved pattern of amino acid sequences. Recently, it was reported that Arabidopsis phyA, with a deletion of amino acids 6–12, showed hypoactive responses to cFR light, because of enhanced destruction of the phyA mutant protein (Trupkin et al. 2007), which contrasts with our results. In this report, the deleted amino acid sequence is at 6–12 in Arabidopsis phyA [–(R)PTQSSEG–], which does not include the –RXXS– sequence, whereas the corresponding phosphorylation sequence of oat phyA at 6–12 is –(R)PASSSSS–. This indicates that there might be no autophosphorylation site in amino acids 6–12 of Arabidopsis phyA, and that the difference in phyA sequences of oat and Arabidopsis may account for the discrepancy in the results. Therefore, it appears that the hypoactivity and enhanced degradation of the 6–12 deletion mutant phyA is not due to autophosphorylation, but rather due to another reason, such as a deletion of seven amino acids changing the conformation of the protein. However, we cannot rule out the possibility that the amino acid sequence that mediates autophosphorylation of the dicot (i.e. Arabidopsis) phyA is different from that of the monocot (i.e. oat) phyA. Many studies of the NTE of phyA have suggested that the NTE is important for the biological activity of phyA (Cherry et al. 1992, Stockhaus et al. 1992, Jordan et al. 1996, Jordan et al. 1997, Casal et al. 2002, Trupkin et al. 2007). Most of these studies suggest that the NTE might have a role in signal attenuation or desensitization in the regulation of photoresponses. Furthermore, modifications in the NTE, including the 6–12 deleted Arabidopsis phyA (Trupkin et al. 2007), influence the degradation of phyA. The present study provides the first direct evidence for the existence of autophosphorylation sites in the NTE. Both Ser8 and Ser18 in the NTE of oat phyA are sites of autophosphorylation. The autophosphorylation in S8A or S18A was each reduced to approximately half of wild-type phyA autophosphorylation levels (Fig. 2B), suggesting that there might be no autophosphorylation preference between the sites. It is noteworthy that the S8/18A double mutant showed almost no autophosphorylation (Fig. 2C), indicating that only two autophosphorylation sites exist on oat phyA. To our knowledge, our report is the first to have determined the exact sites of phyA autophosphorylation, and also confirmed that phyA contained two autophosphorylation sites. Therefore, the NTE of phyA might play a role in the regulation of phytochrome signaling through autophosphorylation, and it is possible that an initial event in phytochrome-mediated signaling involves a post-translational modification of the photoreceptor itself. Functional roles of PhyA autophosphorylation We investigated the functional roles of phyA autophosphorylation by using transgenic plants expressing the autophosphorylation site mutants, S8A, S18A and S8/18A. Our results showed that all of the autophosphorylation site mutants were hyperactive compared with wild-type oat phyA, in terms of FR light sensitivity (Fig. 3), inhibition of hypocotyl growth under cFR and pFR (Fig. 4A), accumulation of anthocyanin (Fig. 4B, C) and blocking of greening (Fig. 4D). The S8/18A mutant was more hyperactive than S8A or S18A, suggesting that Ser8 and Ser18 phosphorylation had cumulative effects. Our results are consistent with previous results from serine to alanine substitution and Δ6–12 deletion mutants, in which both mutants were hyperactive in transgenic plants (Stockhaus et al. 1992, Casal et al. 2002). Therefore, these results suggest that phytochrome autophosphorylation plays a role in the negative regulation of phyA signaling through signal attenuation or desensitization. Previously, phytochrome phosphorylation was suggested to be important for degradation of the phytochrome protein. For example, PAPP5 knockout plants showed a decrease in photoresponsiveness and a decline in the stability of the phyA protein (Ryu et al. 2005). On the other hand, the protein stability of phyA increased in plants overexpressing PAPP5. These results suggest that phosphorylation of phyA accelerates the degradation of phyA, whereas the dephosphorylation of phyA slows the degradation of phyA. Our results confirmed that the protein stability of the autophosphorylation site mutants increased, and thus slowly degraded in the transgenic plants (Fig. 5). The S8/18A mutant was degraded much more slowly than the single site mutants and the wild-type phyA, which could explain the increased photoresponsiveness in the transgenic plants. Even in the white light conditions, in which wild-type phyA was completely degraded, the autophosphorylation site mutants were not completely degraded, but remained in small amounts (Supplementary Fig. S3). Therefore, phyA autophosphorylation is necessary not only for the rapid degradation of phyA, but also for complete degradation under light conditions. Recently, it has been reported that phosphorylated phyA preferentially associates with the COP1–SPA1 complex, the E3 ligase involved in the degradation of phyA (Saijo et al. 2008). This study suggests that phyA phosphorylation is important for its interaction with the COP1–SPA1 complex and shows that the phosphorylated form of phyA is enriched during co-immunoprecipitation with COP1. Therefore, our results are in good agreement with this report, in that the autophosphorylation site mutants of phyA are not efficiently degraded, possibly because of decreased interaction of the mutants with the COP1–SPA1 complex. PhyA degradation has been further investigated using protoplast transfection assays (Fig. 6). PhyA is biosynthesized in the cytosol as the Pr form in the dark and accumulates continuously under illumination. Upon illumination, most phyA is rapidly degraded in the cytosol by forming SAPs, and some phyA is translocated into the nucleus to initiate signaling cascades. The light-induced formation of SAPs is considered to be a molecular property of phyA, although the exact function of the formation SAPs is not fully known (Eichenberg et al. 1999). Thus, we observed the formation of SAPs by the autophosphorylation site mutants and found that these mutants formed SAPs more slowly than wild-type phyA, which exactly matched the results of the protein degradation assays. Since the SAPs are considered to be the functional complex for phyA degradation, the slow formation of SAPs is correlated with the slow degradation of phyA. Since the phyA accumulated in the dark for the sensitive light signaling may need to be degraded rapidly for their desensitization, the SAPs might be formed for the quick and efficient degradation of phyA, in which autophosphorylation is necessary for the formation of SAPs. Therefore, underphosphorylation of phyA in the mutants might delay the formation of SAPs and thus slow the degradation of phyA, which could sustain the activity of phyA and impart increased photoresponses to FR light in the plants. When we consider the findings that phytochrome autophosphorylation occurs similarly in the Pr and Pfr forms and the phosphorylated phyA in the Pfr form is degraded in vivo, phyA might be equally autophosphorylated in vivo between the Pr and Pfr forms and the photoconversion from Pr to Pfr might not be significantly affected by the autophosphorylation. The finding that phyA is down-regulated by phosphorylation might be similar to the case for G-protein-coupled receptors. As an example, rhodopsin is desensitized by phosphorylation through rhodopsin kinase (Arshavsky 2002, Sokal et al. 2002). The desensitization of rhodopsin is important for vision. If the activation of rhodopsin is not attenuated efficiently, blindness will occur. However, the signal attenuation mechanism of rhodopsin might be different from that of phytochrome: activation of rhodopsin results in its phosphorylation and subsequent binding to arrestin-like proteins, whereas phytochrome phosphorylation and its activation results in acceleration of its degradation through binding with an E3 ligase complex (i.e. COP1–SPA1). Overall, our results provide a plausible model for the functional role of phyA autophosphorylation. In the dark, phyA proteins are synthesized and accumulate as the Pr form in the cytosol, consisting of autophosphorylated phyA and dephosphorylated phyA, due to the action of protein phosphatases. Upon illumination, the Pr form is photoactivated to the Pfr form, which can be degraded via the ubiquitin–26S proteasome protein degradation pathway. At this point, the phosphorylated Pfr form is rapidly degraded, compared with the unphosphorylated phyA species. The balance between the autophosphorylated and dephosphorylated phyA proteins is probably shifted to the autophosphorylated phyA, because the phosphorylated phyA proteins are efficiently degraded by the ubiquitin–26S proteasome through the COP1–SPA1 complex. This rapid phyA degradation mechanism that involves autophosphorylation might facilitate the efficient desensitization of the phyA signal. Otherwise, phytochrome protein phosphatases might increase the phyA protein stability with enhanced light responses by decreasing the amounts of the autophosphorylated phyA. If phyA activity is not attenuated upon illumination, plants could over-respond to light by eliciting too many photomorphogenic signaling events and/or could become desensitized to subsequent changes in light quality or quantity. Therefore, this regulation may provide an efficient means of rapidly controlling the number of active phyA photoreceptors available to initiate signaling events, and thereby improve the response of plants to fluctuating light environments. Materials and Methods Phytochrome constructs Full-length cDNA (amino acids 1–1,129) of Avena sativa (oat) phyA was cloned into pGEM®-11zf(+) (Promega) from pFY122 (Boylan and Quail 1989) by digestion with BamHI and EcoRI, and was used as a template for mutagenesis and PCRs. Each of the two known phosphorylation sites on oat phyA (i.e. Ser8 and Ser18) was mutagenized to alanine by site-directed mutagenesis using the GeneEditor™ in vitro Site-Directed Mutagenesis System (Promega). The mutagenic primers used were phosphorylated-5′-CCTCAAGGCCTGCAGCCAGTTCTTCCA GC-3′ for Ser8Ala (S8A), and phosphorylated-5′-GGAACCG CCAGAGTGCGCAGGCAAGGGTG-3′ for Ser18Ala (S18A). After the mutagenesis of S8A and S18A, the second mutagenesis was carried out to make a combination mutant Ser8Ala/Ser18Ala (S8/18A). After mutagenesis, all the constructs were confirmed by DNA sequencing. In addition to these S8A and S18A mutants, the NTE was deleted to generate the Δ65 mutant (amino acids 66–1,129), and the S599A mutant was obtained as previously described (Kim et al. 2004). All the constructs were then cloned into pPIC3.5K for the preparation of recombinant proteins using the Pichia Protein Expression System (Invitrogen) or into the pBI121 binary plasmid (Clontech) for the generation of transgenic plants by transformation into phyA-deficient A. thaliana (phyA-201, Ler ecotype), as described (Kim et al. 2004). The constructs in pPIC3.5K contained the streptavidin affinity tag (strep-tag), consisting of 10 amino acids at the C-terminus for efficient purification by streptavidin affinity chromatography. Plant materials and growth conditions The constructs S8A, S18A, S8/18A and wild-type PHYA in pBI121 were each introduced into phyA-201 using the Agrobacterium tumefaciens (strain GV3101)-mediated floral dip method, as described (Clough and Bent 1998). Transgenic lines segregating ∼3 : 1 for antibiotic resistance in the T2 generation were selected, and the T3 or T4 homozygous generation was used for subsequent analyses. Arabidopsis plants were grown on soil in a culture room (22°C with a 16 h photoperiod), following routine procedures. For seedling assays, seeds were surface-sterilized, incubated at 4°C for 3 d in the dark, and placed on 0.8% phytoagar (w/v) medium containing half-strength MS salts and vitamins. The plates were then transferred to a growth chamber (22°C with a 16 h photoperiod). Expression, reconstitution and purification of PhyA–chromophore holoproteins The pPIC3.5K constructs bearing PHYA genes were transformed into Pichia cells by means of a Micropulser™ Electroporation apparatus (Bio-Rad). Recombinant phytochrome proteins were expressed in the Pichia expression system (Invitrogen) and purified by streptavidin affinity chromatography (IBA), according to the procedure described by the manufacturer. Phycocyanobilin (PCB) was prepared from Spirulina platensis extracts (Sigma) by methanolysis as described (Park et al. 2000), and used as a chromophore for the holo-phytochrome assembly. The concentration of purified PCB was quantified by absorption spectroscopy in HCl (2%)/methanol, using an extinction coefficient (ε) of 37,900 M−1 cm−1 at 690 nm. From the harvested Pichia cells, crude extracts were prepared by breaking cells in liquid nitrogen using a homogenizer (Nihonseiki Kaisha). The phytochrome samples were then precipitated by adding 0.23 g l−1 ammonium sulfate, resuspended in a buffer [100 mM Tris (pH 7.8), 1 mM EDTA], the PCB chromophores in dimethylsulfoxide (DMSO) were added to the samples at a final concentration of 10 μM and the mixture was incubated for in vitro reconstitution on ice for 1 h. After dialysis to remove free chromophores, the samples were loaded onto streptavidin affinity chromatography columns and holo-phytochromes without free chromophores were purified. Zn2+ fluorescence and spectroscopic analysis For Zn2+ fluorescence assays to assess the chromophore ligation, the protein samples were analyzed on a 10% SDS–polyacrylamide gel and the gel was then soaked in 20 mM zinc acetate/150 mM Tris–HCl (pH 7.0) for 5–30 min at room temperature with gentle shaking. Zinc fluorescence of holo-phytochromes was visualized under UV light (312 nm). For the spectroscopic analysis of recombinant phyA, absorption spectra were recorded by a diode array UV/VIS spectrophotometer (Cary) after red or FR light irradiation. All the spectroscopic experiments were carried out under the green safety light condition, which consisted of a white fluorescent lamp equipped with a plastic filter (Rosco) with a maximal transmittance at 500 nm, and a fiberoptic illuminator system (Cole-Parmer) equipped with 656 and 730 nm interference filters (Oriel) was used as a light source. The light intensity was 8 W m−2 for red light and 6 W m−2 for FR light. The samples were illuminated with red or FR light for 5 min. A differential spectrum was obtained by subtracting the Pfr spectrum from the Pr spectrum. The concentrations of the holo-phytochrome samples were calculated from the λmax of the Pr peak with an extinction coefficient of 132,000 M−1 cm−1. The concentrations of other protein samples were determined using the Bradford Protein Assay Kit (Bio-Rad), with bovine serum albumin as a standard. Autophosphorylation assay Phytochrome phosphorylation experiments were performed as described (Yeh and Lagarias 1998, Kim et al. 2004) with minor modifications. The reaction mixtures (total volume of 20 μl) contained kinase buffer [50 mM Tris–HCl (pH 7.8), 0.2 mM EDTA, 4 mM dithiothreitol (DTT) and 5 mM MgCl2] and purified recombinant phytochromes (0.5–2 μg), in either Pr or Pfr. Phytochrome samples were irradiated with red or FR light for 5 min before the start of a reaction. Autophosphorylation reactions were started by adding 0.1 mM ATP containing 10 μCi of [γ-32P]ATP (Perkin-Elmer, 3,000 Ci mmol−1) and were incubated at 30°C for 1 h. Reactions were stopped by the addition of 5 μl of 5× SDS sample buffer. Proteins were resolved on 10% (w/v) SDS–polyacrylamide gels, dried under vacuum and exposed to X-ray films for autoradiography. Coomassie Blue protein staining and zinc fluorescence assays were performed before drying. Because polycations, such as histone H1 (Roche), are known to increase the phytochrome autophosphorylation activity, histone H1 (0.5–2 μg) was added to phytochrome samples in the indicated experiments. For a quantitative analysis of phytochrome phosphorylation, 32P radiolabel incorporation was determined by scintillation counting of the excised protein bands from SDS–polyacrylamide gels as described (Wong et al. 1989). Using a curve of the relationship between the molar concentration of 32P and scintillation counting, 4 pmol of phyA was reacted with 10 μCi of [γ-32P]ATP, and the samples were loaded on SDS–polyacrylamide gels. The molar incorporation of 32P was determined by measuring the 32P concentration in each excised protein band from SDS–polyacrylamide gels by a liquid scintillation counter (Beckman Coulter, Inc.). RT–PCR and Western blot analysis To compare transcript levels of oat phyA from transgenic plants, total RNAs were extracted using a RNeasy® plant kit (Qiagen), and then a PrimeScript™ first-strand cDNA synthesis kit (TAKARA) was used for cDNA synthesis. PCR amplification for oat PHYA was performed with a forward primer at 2,704 bp, 5′-TACATGAGACATGCGATCAAC-3′, and a reverse primer at 3,264 bp, 5′-GCTCAAGCCCTCCTCTGACTG-3′. As a control, ACT2 was amplified with a forward primer, 5′-TTGACCTTG CTGGACGTG-3′, and a reverse primer, 5′-GGAAGCAAGAAT GGAACCAC-3′. To investigate phyA protein levels from transgenic plants, each transgenic line was cultured on half-strength MS medium with vitamins including 0.8% phytoagar and 50 μM MG-132 (A.G. Scientific, Inc.) for 2.5 d in darkness. Approximately 100 dark-grown seedlings were ground and extracted with an extraction buffer [70 mM Tris–HCl, pH 8.3, 7 mM EDTA, 35% ethylene glycol, 98 mM ammonium sulfate, 14 mM sodium metabisulfite, 0.07% polyethyleneimine, 2.8 mM phenylmethylsulfonyl fluoride (PMSF), and 10 μM MG-132]. The extracted protein samples were centrifuged at 14,000 r.p.m. at 4°C for 15 min, and 30 μg of protein samples were used for Western blot analysis. Photoresponse analyses of transgenic Arabidopsis plants To assess the expression of oat phyA and serine to alanine mutant proteins in transgenic Arabidopsis plants, Western blot analysis was performed on 4-day-old dark-grown seedlings as described (Kim et al. 2004). A 40 μg aliquot of crude extract was separated by 10% SDS–PAGE and transferred to a polyvinylidene difluoride (PVDF) membrane (Hybond-P, GE Healthcare). The membrane was then incubated with oat phyA-specific oat25 monoclonal antibody and developed using an ECL Advanced™ Western Blotting Analysis System (GE Healthcare). Hypocotyl lengths were measured in response to light treatments as described (Fankhauser and Casal 2004). The seeds were sown on half-strength MS medium and cold treated for 3 d in the dark. The seeds were then exposed to white light for 4 h to promote germination, returned to darkness (22°C) for 24 h, and then grown for 3 d under cFR or pFR light with various fluence rates, using an LED growth chamber (Vision, Korea). The hypocotyls were photographed with a digital camera (Nikon), and then measured with image analysis software (Scion Image, Frederick, MD, USA). The anthocyanin content and total chlorophyll content of seedlings were determined as described (Fankhauser and Casal 2004). Relative anthocyanin levels were determined by collecting 15 seedlings from each of the light treatments/line and incubating them overnight in 500 μl of methanol acidified with 1% HCl, with shaking, in the dark. The next day, 500 μl of chloroform was added, and the sample was vortexed and briefly centrifuged to separate the anthocyanins from chlorophyll. The total anthocyanin content was determined by measuring the A530 and A657 of the aqueous phase using a UV/VIS spectrophotometer (Cary). The relative amount of anthocyanin per seedling was calculated by subtracting the A657 from the A530. Total chlorophyll was determined from samples containing 15 seedlings. Seedlings were extracted by incubating in 1 ml of 80% acetone, with shaking, overnight in the dark. Chlorophyll levels were measured spectroscopically and the amount of chlorophyll was determined using the equation: chlorophylla + b = 7.15 × OD660 nm + 18.71 × OD647 nm. In vivo phytochrome degradation assay The transgenic plant seeds were germinated, and grown for 3.5 d in the dark. Seedlings were then illuminated with red light (28 μmol m−2 s−1) in the LED growth chamber (Vision Scientific Co., Ltd.) and harvested at the indicated times. The harvested seedlings were stored in liquid nitrogen and the protein samples were prepared as described (Kim et al. 2004). A 40 μg aliquot of crude extract was loaded onto SDS–polyarylamide gels, and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. To investigate the in vivo degradation of phyA using mesophyll cell protoplasts, transient expression of GFP-fused phyA mutants as well as wild-type phyA was examined. GFP–phyA constructs were cloned into the pCsVMV:GFP vector under the control of the CsVMV promoter (Verdaguer et al. 1996), using EcoRI and BamHI and primer set, 5′-CGGAATTCATGTCTTCCTCAAGGCCTGCTTCC-3′ (forward) and 5′-CGGGATCCTTGTCCCATTGCTGTTGGAGCGGAAGC-3′ (reverse) for the wild type and S18A phyA, and 5′-CGGAATTCATGTCTTCCTCAAGGCCTGCTGCAAGT-3′ (forward) and the same reverse primer for S8A and S8/18A phyA. Mesophyll cell protoplasts were isolated from phyA-deficient A. thaliana (phyA-211, Col-0 ecotype) and transfected as described (Hwang and Sheen 2001). Transfected protoplasts were incubated overnight in the dark, and then SAPs were observed during red light irradiation for the indicated periods of time using a confocal microscope. These confocal images were obtained using a laser scanning confocal microscope (Leica TCS SP5 AOBS/Tandem) at Korea Basic Science Institute, Gwangju Center. Funding This work was supported by the Plant Diversity Research Center of 21st Century Frontier Research Program [grant no. PF06302–02]; the MEST/NRF to the Environment Biotechnology National Core Research Center (NCRC) [grant no. 20090091493]; the Korea Research Foundation Grant funded by the Korean Government (MOEHRD) [grant no. KRF-2006-312-C00671]. References Arshavsky VY.  Rhodopsin phosphorylation: from terminating single photon responses to photoreceptor dark adaptation,  Trends Neurosci. ,  2002, vol.  25 (pg.  124- 126) Google Scholar CrossRef Search ADS PubMed  Bae G,  Choi G.  Decoding of light signals by plant phytochromes and their interacting proteins,  Annu. Rev. Plant Biol. ,  2008, vol.  59 (pg.  281- 311) Google Scholar CrossRef Search ADS PubMed  Biermann BJ,  Pao LI,  Feldman LJ.  Pr-specific phytochrome phosphorylation in vitro by a protein kinase present in anti-phytochrome maize immunoprecipitates,  Plant Physiol. ,  1994, vol.  105 (pg.  243- 251) Google Scholar CrossRef Search ADS PubMed  Boylan MT,  Quail PH.  Oat phytochrome is biologically active in transgenic tomatoes,  Plant Cell ,  1989, vol.  1 (pg.  765- 773) Google Scholar CrossRef Search ADS PubMed  Boylan MT,  Quail PH.  Phytochrome a overexpression inhibits hypocotyl elongation in transgenic Arabidopsis,  Proc. Natl Acad. Sci. USA ,  1991, vol.  88 (pg.  10806- 10810) Google Scholar CrossRef Search ADS   Casal JJ,  Davis SJ,  Kirchenbauer D,  Viczian A,  Yanovsky MJ,  Clough RC, et al.  The serine-rich N-terminal domain of oat phytochrome a helps regulate light responses and subnuclear localization of the photoreceptor,  Plant Physiol. ,  2002, vol.  129 (pg.  1127- 1137) Google Scholar CrossRef Search ADS PubMed  Chen M,  Chory J,  Fankhauser C.  Light signal transduction in higher plants,  Annu. Rev. Genet. ,  2004, vol.  38 (pg.  87- 117) Google Scholar CrossRef Search ADS PubMed  Cherry JR,  Hondred D,  Walker JM,  Vierstra RD.  Phytochrome requires the 6-kDa N-terminal domain for full biological activity,  Proc. Natl Acad. Sci. USA ,  1992, vol.  89 (pg.  5039- 5043) Google Scholar CrossRef Search ADS   Clough SJ,  Bent AF.  Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana,  Plant J. ,  1998, vol.  16 (pg.  735- 743) Google Scholar CrossRef Search ADS PubMed  Eichenberg K,  Kunkel T,  Kretsch T,  Speth V,  Schafer E.  In vivo characterization of chimeric phytochromes in yeast,  J. Biol. Chem. ,  1999, vol.  274 (pg.  354- 359) Google Scholar CrossRef Search ADS PubMed  Emmler K,  Stockhaus J,  Chua NH,  Schafer E.  An amino-terminal deletion of rice phytochrome A results in a dominant negative suppression of tobacco phytochrome A activity in transgenic tobacco seedlings,  Planta ,  1995, vol.  197 (pg.  103- 110) Google Scholar CrossRef Search ADS PubMed  Essen LO,  Mailliet J,  Hughes J.  The structure of a complete phytochrome sensory module in the Pr ground state,  Proc. Natl Acad. Sci. USA ,  2008, vol.  105 (pg.  14709- 14714) Google Scholar CrossRef Search ADS   Fankhauser C.  Phytochromes as light-modulated protein kinases,  Semin. Cell Dev. Biol. ,  2000, vol.  11 (pg.  467- 473) Google Scholar CrossRef Search ADS PubMed  Fankhauser C,  Casal JJ.  Phenotypic characterization of a photomorphogenic mutant,  Plant J. ,  2004, vol.  39 (pg.  747- 760) Google Scholar CrossRef Search ADS PubMed  Fankhauser C,  Chen M.  Transposing phytochrome into the nucleus,  Trends Plant Sci. ,  2008, vol.  13 (pg.  596- 601) Google Scholar CrossRef Search ADS PubMed  Han YJ,  Song PS,  Kim JI.  Phytochrome-mediated photomorphogenesis in plants,  J. Plant Biol. ,  2007, vol.  50 (pg.  230- 240) Google Scholar CrossRef Search ADS   Henriques R,  Jang IC,  Chua NH.  Regulated proteolysis in light-related signaling pathways,  Curr. Opin. Plant Biol. ,  2009, vol.  12 (pg.  49- 56) Google Scholar CrossRef Search ADS PubMed  Hunt RE,  Pratt LH.  Partial characterization of undegraded oat phytochrome,  Biochemistry ,  1980, vol.  19 (pg.  390- 394) Google Scholar CrossRef Search ADS PubMed  Hwang I,  Sheen J.  Two-component circuitry in Arabidopsis cytokinin signal transduction,  Nature ,  2001, vol.  413 (pg.  383- 389) Google Scholar CrossRef Search ADS PubMed  Jiao Y,  Lau OS,  Deng XW.  Light-regulated transcriptional networks in higher plants,  Nat. Rev. Genet. ,  2007, vol.  8 (pg.  217- 230) Google Scholar CrossRef Search ADS PubMed  Jordan ET,  Cherry JR,  Walker JM,  Vierstra RD.  The amino-terminus of phytochrome A contains two distinct functional domains,  Plant J. ,  1996, vol.  9 (pg.  243- 257) Google Scholar CrossRef Search ADS PubMed  Jordan ET,  Marita JM,  Clough RC,  Vierstra RD.  Characterization of regions within the N-terminal 6-kilodalton domain of phytochrome A that modulate its biological activity,  Plant Physiol. ,  1997, vol.  115 (pg.  693- 704) Google Scholar CrossRef Search ADS PubMed  Kim DH,  Kang JG,  Yang SS,  Chung KS,  Song PS,  Park CM.  A phytochrome-associated protein phosphatase 2A modulates light signals in flowering time control in Arabidopsis,  Plant Cell ,  2002, vol.  14 (pg.  3043- 3056) Google Scholar CrossRef Search ADS PubMed  Kim JI,  Park JE,  Zarate X,  Song PS.  Phytochrome phosphorylation in plant light signaling,  Photochem. Photobiol. Sci. ,  2005, vol.  4 (pg.  681- 687) Google Scholar CrossRef Search ADS PubMed  Kim JI,  Shen Y,  Han YJ,  Park JE,  Kirchenbauer D,  Soh MS, et al.  Phytochrome phosphorylation modulates light signaling by influencing the protein–protein interaction,  Plant Cell ,  2004, vol.  16 (pg.  2629- 2640) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Jiang XY,  Smith DL,  Song PS.  Posttranslational modification of oat phytochrome A: phosphorylation of a specific serine in a multiple serine cluster,  Biochemistry ,  1997, vol.  36 (pg.  10595- 10599) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Jiang XY,  Smith DL,  Song PS.  Mass spectrometric characterization of oat phytochrome A: isoforms and posttranslational modifications,  Protein Sci. ,  1999, vol.  8 (pg.  1032- 1044) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Wells TA,  Song PS.  Protein kinase A-catalyzed phosphorylation and its effect on conformation in phytochrome A,  Biochemistry ,  1996, vol.  35 (pg.  6585- 6594) Google Scholar CrossRef Search ADS PubMed  McMichael R.W.Jr.,  Lagarias,  J. C.  Phosphopeptide mapping of Avena phytochrome phosphorylated by protein kinases in vitro,  Biochemistry ,  1990, vol.  29 (pg.  3872- 3878) Google Scholar CrossRef Search ADS PubMed  Nagatani A.  Light-regulated nuclear localization of phytochromes,  Curr. Opin. Plant Biol. ,  2004, vol.  7 (pg.  708- 711) Google Scholar CrossRef Search ADS PubMed  Park CM,  Shim JY,  Yang SS,  Kang JG,  Kim JI,  Luka Z, et al.  Chromophore–apoprotein interactions in Synechocystis sp. PCC6803 phytochrome Cph1,  Biochemistry ,  2000, vol.  39 (pg.  6349- 6356) Google Scholar CrossRef Search ADS PubMed  Phee BK,  Kim JI,  Shin DH,  Yoo J,  Park KJ,  Han YJ, et al.  A novel protein phosphatase indirectly regulates phytochrome-interacting factor 3 via phytochrome,  Biochem. J. ,  2008, vol.  415 (pg.  247- 255) Google Scholar CrossRef Search ADS PubMed  Quail PH.  The phytochromes: a biochemical mechanism of signaling in sight?,  Bioessays ,  1997, vol.  19 (pg.  571- 579) Google Scholar CrossRef Search ADS PubMed  Rockwell NC,  Su YS,  Lagarias JC.  Phytochrome structure and signaling mechanisms,  Annu. Rev. Plant Biol. ,  2006, vol.  57 (pg.  837- 858) Google Scholar CrossRef Search ADS PubMed  Ryu JS,  Kim JI,  Kunkel T,  Kim BC,  Cho DS,  Hong SH, et al.  Phytochrome-specific type 5 phosphatase controls light signal flux by enhancing phytochrome stability and affinity for a signal transducer,  Cell ,  2005, vol.  120 (pg.  395- 406) Google Scholar CrossRef Search ADS PubMed  Saijo Y,  Zhu D,  Li J,  Rubio V,  Zhou Z,  Shen Y, et al.  Arabidopsis COP1/SPA1 complex and FHY1/FHY3 associate with distinct phosphorylated forms of phytochrome A in balancing light signaling,  Mol. Cell ,  2008, vol.  31 (pg.  607- 613) Google Scholar CrossRef Search ADS PubMed  Sokal I,  Pulvermuller A,  Buczylko J,  Hofmann KP,  Palczewski K.  Rhodopsin and its kinase,  Methods Enzymol. ,  2002, vol.  343 (pg.  578- 600) Google Scholar PubMed  Stockhaus J,  Nagatani A,  Halfter U,  Kay S,  Furuya M,  Chua NH.  Serine-to-alanine substitutions at the amino-terminal region of phytochrome A result in an increase in biological activity,  Genes Dev. ,  1992, vol.  6 (pg.  2364- 2372) Google Scholar CrossRef Search ADS PubMed  Trupkin SA,  Debrieux D,  Hiltbrunner A,  Fankhauser C,  Casal JJ.  The serine-rich N-terminal region of Arabidopsis phytochrome A is required for protein stability,  Plant Mol. Biol. ,  2007, vol.  63 (pg.  669- 678) Google Scholar CrossRef Search ADS PubMed  Verdaguer B,  de Kochko A,  Beachy RN,  Fauquet C.  Isolation and expression in transgenic tobacco and rice plants, of the cassava vein mosaic virus (CVMV) promoter,  Plant Mol. Biol. ,  1996, vol.  31 (pg.  1129- 1139) Google Scholar CrossRef Search ADS PubMed  Wagner D,  Quail PH.  Mutational analysis of phytochrome B identifies a small COOH-terminal-domain region critical for regulatory activity,  Proc. Natl Acad. Sci. USA ,  1995, vol.  92 (pg.  8596- 8600) Google Scholar CrossRef Search ADS   Whitelam GC,  Johnson E,  Peng J,  Carol P,  Anderson ML,  Cowl JS, et al.  Phytochrome A null mutants of Arabidopsis display a wild-type phenotype in white light,  Plant Cell ,  1993, vol.  5 (pg.  757- 768) Google Scholar CrossRef Search ADS PubMed  Wong YS,  Cheng HC,  Walsh DA,  Lagarias JC.  Phosphorylation of Avena phytochrome in vitro as a probe of light-induced conformational changes,  J. Biol. Chem. ,  1986, vol.  261 (pg.  12089- 12097) Google Scholar PubMed  Wong YS,  Lagarias JC.  Affinity labeling of Avena phytochrome with ATP analogs,  Proc. Natl Acad. Sci. USA ,  1989, vol.  86 (pg.  3469- 3473) Google Scholar CrossRef Search ADS   Wong YS,  McMichael RW,  Lagarias JC.  Properties of a polycation-stimulated protein kinase associated with purified Avena phytochrome,  Plant Physiol. ,  1989, vol.  91 (pg.  709- 718) Google Scholar CrossRef Search ADS PubMed  Yeh KC,  Lagarias JC.  Eukaryotic phytochromes: light-regulated serine/threonine protein kinases with histidine kinase ancestry,  Proc. Natl Acad. Sci. USA ,  1998, vol.  95 (pg.  13976- 13981) Google Scholar CrossRef Search ADS   Abbreviations Abbreviations ACT actin gene cFR continuous far-red CHS chalcone synthase gene FR far-red FyPP flower-specific phytochrome-associated protein phosphatase GFP green fluorescent protein HIR high-irradiance response HKRD histidine kinase-related domain LED light-emitting diode NDPK2 nucleoside di-phosphate kinase 2 NTE N-terminal extension PAPP5 phytochrome-associated protein phosphatase 5 PCB phycocyanobilin Pfr far-red light-absorbing form of phytochrome pFR pulsed far-red phyA phytochrome A PIF3 phytochrome-interacting factor 3 Pr red light-absorbing form of phytochrome RT–PCR reverse transcription–PCR SAPs sequestered areas of phytochrome VLFR very-low- fluence response. © The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Plant and Cell Physiology Oxford University Press

Functional Characterization of Phytochrome Autophosphorylation in Plant Light Signaling

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Oxford University Press
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© The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org
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0032-0781
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1471-9053
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10.1093/pcp/pcq025
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20495342
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Abstract

Abstract Plant phytochromes, molecular light switches that regulate various aspects of plant growth and development, are phosphoproteins that are also known to be autophosphorylating serine/threonine kinases. Although a few protein phosphatases that directly interact with and dephosphorylate phytochromes have been identified, no protein kinase that acts on phytochromes has been reported thus far, and the exact site of phytochrome autophosphorylation has not been identified. In this study, we investigated the functional role of phytochrome autophosphorylation. We first mapped precisely the autophosphorylation sites of oat phytochrome A (phyA), and identified Ser8 and Ser18 in the 65 amino acid N-terminal extension (NTE) region as being the autophosphorylation sites. The in vivo functional roles of phytochrome autophosphorylation were examined by introducing autophosphorylation site mutants into phyA-deficient Arabidopsis thaliana. We found that all the transgenic plants expressing the autophosphorylation site mutants exhibited hypersensitive light responses, indicating an increase in phyA activity. Further analysis showed that these phyA mutant proteins were degraded at a significantly slower rate than wild-type phyA under light conditions, which suggests that the increased phyA activity of the mutants is related to their increased protein stability. In addition, protoplast transfection analyses with green fluorescent protein (GFP)-fused phyA constructs showed that the autophosphorylation site mutants formed sequestered areas of phytochrome (SAPs) in the cytosol much more slowly than did wild-type phyA. These results suggest that the autophosphorylation of phyA plays an important role in the regulation of plant phytochrome signaling through the control of phyA protein stability. Introduction Phytochromes are red/far-red (R/FR) photoreceptors that regulate many aspects of plant growth and development in response to informational light signals from the environment (Rockwell et al. 2006, Bae and Choi 2008). They are dimeric chromoproteins with covalently linked tetrapyrrole chromophore phytochromobilin, and exist in two photointerconvertible species, red-light-absorbing Pr and FR-light absorbing Pfr forms. Phytochromes are biosynthesized as the Pr form in the dark, which can be phototransformed into the Pfr form upon exposure to red light. This photoactivation of phytochromes from the Pr form to the Pfr form induces the highly regulated signaling network for photomorphogenesis in plants (Chen et al. 2004, Han et al. 2007). Recent studies on phytochrome-mediated light signaling revealed that phytochrome photoactivation has a major impact on the control of protein subcellular localization (Nagatani 2004, Fankhauser and Chen 2008), transcription (Jiao et al. 2007), protein stability (Henriques et al. 2009) and protein phosphorylation (Kim et al. 2005). However, despite intensive studies on phytochromes involving a broad range of experimental approaches since their first discovery in the 1950s, the exact biochemical mechanisms underlying the phytochrome regulation of photoresponses in plants have not been fully elucidated. Phytochromes are known to be phosphoproteins by phosphate analysis on purified phytochrome preparations (Hunt and Pratt 1980), and the sites of phytochrome phosphorylation have been investigated with purified phytochrome A (phyA) from oat seedlings (Lapko et al. 1996, Lapko et al. 1997, Lapko et al. 1999). There are two phosphorylation sites (Ser8 and Ser18) in the N-terminal extension (NTE) region of the phyA molecule, and one site (Ser599) in the hinge region between the N- and C-terminal domains. In previous reports, the NTE was shown to be required for full biological activity of phyA (Cherry et al. 1992), and serine to alanine substitution in the NTE region including Ser8 and Ser18 has resulted in increased biological activity, showing that transgenic plants expressing this mutant form of phyA were hypersensitive to light and had a dwarf phenotype (Stockhaus et al. 1992). These results suggest that phosphorylation in the NTE region is a mechanism of signal attenuation or desensitization of phytochrome signal transduction (Emmler et al. 1995, Jordan et al. 1996, Jordan et al. 1997, Casal et al. 2002). However, it remains unclear how this substitution generates an attenuation signal. In addition, functional characterization has shown that phosphorylation at Ser599 prevents the interaction of phyA with its signal transducers such as nucleoside di-phosphate kinase 2 (NDPK2) and phytochrome-interacting factor 3 (PIF3), which suggests that the hinge region of phytochromes serves as a phosphorylatable signal-modulating site that regulates protein–protein interactions between phytochrome and its signal transducers (Kim et al. 2004). These data suggest the importance of phytochrome phosphorylation in the light signaling activities of phytochromes. The observation that phytochromes are phosphoproteins suggests the existence of protein kinase(s) that phosphorylate the phytochromes and protein phosphatase(s) that dephosphorylate them. However, despite extensive studies of phytochrome-interacting proteins, there is no report thus far of a protein kinase that can phosphorylate phytochromes. On the other hand, a few protein phosphatases have been reported as being able to interact with and dephosphorylate phytochromes, including flower-specific phytochrome-associated protein phosphatase (FyPP) (Kim et al. 2002), phytochrome-associated protein phosphatase 5 (PAPP5) (Ryu et al. 2005) and phytochrome-associated protein phosphatase type 2C (PAPP2C) (Phee et al., 2008). The FyPP-overexpressing transgenic plants had enhanced phytochrome activity during flowering and hypocotyl shortening, whereas the antisense repression of FyPP resulted in transgenic plants with reduced phytochrome activity (Kim et al. 2002). PAPP5, which is involved in the regulation of de-etiolation, positively influences the protein stability of phytochrome and the interaction of phytochrome with a downstream transducer NDPK2 (Ryu et al. 2005). PAPP2C also positively regulates the light responses of plants (Phee et al. 2008). These observations suggest that phytochrome phosphorylation and dephosphorylation play important roles in phytochrome-mediated light signaling. Since no protein kinase is known to act on phytochromes, it is possible that phytochromes are phosphorylated by means of autophosphorylation because phytochromes are known to be autophosphorylating serine/threonine protein kinases (Yeh and Lagarias 1998). Because phytochrome phosphorylation and dephosphorylation have been known to play an important role in the regulation of phytochrome functions (Kim et al. 2005), phytochrome phosphorylation would be controlled by autophosphorylation and protein phosphatases. However, the in vivo functional role of phytochrome phosphorylation remains to be explored. Thus, several important aspects of the role of phosphorylation in phytochrome signaling remain to be resolved, including the determination of autophosphorylation site(s) and the in vivo functional role of phytochrome autophosphorylation. In this study, we determined the autophosphorylation sites of oat phyA and investigated the functional role of phyA autophosphorylation using phyA-deficient Arabidopsis plants transformed with oat phyA autophosphorylation site mutants. We demonstrated that oat phyA was autophosphorylated on Ser8 and Ser18 in the NTE region and that the transgenic plants of autophosphorylation site-deleted mutants exhibited hypersensitive phenotypes in response to FR light, indicating that the mutants are hyperactive in plants. Further studies demonstrated that the phyA mutant proteins degraded slowly compared with wild-type phyA, which could account for the enhanced photoresponse of transgenic plants bearing the phyA mutant. Therefore, our results provide direct evidence for the functional role of phyA autophosphorylation. Results Oat phyA contains autophosphorylation activity that is stimulated by the addition of histone H1 Before investigating the phytochrome autophosphorylation site(s), we first examined the autophosphorylation of purified recombinant oat phyA. Since the expression of full-length plant phytochromes in a recombinant system is known to be very difficult, we expressed and purified recombinant phyA proteins using the Pichia protein expression system and streptavidin affinity chromatography, as we previously reported (Kim et al. 2004). Spectroscopic analyses of the purified serine to alanine phyA mutants, namely S8A, S18A and S8/18A, showed that the absorbance and R/FR difference spectra of the mutants were identical to those of wild-type phyA (Supplementary Table S1 and Supplementary Data), indicating that the protein integrity of the mutants is similar to that of the wild type. We then performed phytochrome autophosphorylation experiments with the purified phyA proteins. The phyA protein is known to be autophosphorylated, and the autophosphorylation is stimulated in the presence of polycations, such as histone H1 (Wong et al. 1986, Wong et al. 1989, Yeh and Lagarias 1998). Thus, we investigated the autophosphorylation of purified recombinant phyA in the presence or absence of histone H1 (Fig. 1A). These experiments confirmed that phyA possessed autophosphorylation activity that was stimulated by histone H1. Zinc fluorescence and SDS–PAGE also confirmed the formation of chromophore-ligated phyA proteins and the protein purity used in the reactions, respectively. Under our experimental conditions, Pr and Pfr forms of oat phyA were similarly autophosphorylated, and the addition of histone H1 preferentially stimulated phyA autophosphorylation of the Pr form, consistent with findings of a previous report (Yeh and Lagarias 1998). When the autoradiogram bands were quantified with a densitometer (Molecular Dynamics), the intensities of autophosphorylated Pr and Pfr forms of phyA in the presence of histone H1 were 3.5 and 1.7, respectively, with the Pr intensity in the absence of histone H1 being set at 1.0. In addition, histone H1 was phosphorylated by phyA, indicating the phosphotransfer activity of phyA. Unlike Pr-preferential stimulated autophosphorylation of phyA, histone H1 phosphorylation was not light dependent. Fig. 1 View largeDownload slide Autophosphorylation analysis of recombinant oat phyA. (A) Autophosphorylation of purified recombinant oat phyA in the absence or presence of histone H1. FL-phyA, full-length wild-type oat phyA; H1, histone H1; S, protein size standards. Autoradiogram (Autorad), zinc fluorescence (Zinc) and SDS–PAGE are shown. A 1.5 μg aliquot of the Pr or Pfr form of phyA was used in the reactions with or without 0.75 μg of histone H1. (B) Time-dependent autophosphorylation of the Pr and Pfr forms of oat phyA in the absence or presence of histone H1. In these reactions, 4 pmol of the Pr or Pfr form of phyA was used with or without 2 μg of histone H1. (C) 32P incorporation in oat phyA by autophosphorylation shown in (B). Each analysis was repeated three times. Error bar = SD (n = 3). Fig. 1 View largeDownload slide Autophosphorylation analysis of recombinant oat phyA. (A) Autophosphorylation of purified recombinant oat phyA in the absence or presence of histone H1. FL-phyA, full-length wild-type oat phyA; H1, histone H1; S, protein size standards. Autoradiogram (Autorad), zinc fluorescence (Zinc) and SDS–PAGE are shown. A 1.5 μg aliquot of the Pr or Pfr form of phyA was used in the reactions with or without 0.75 μg of histone H1. (B) Time-dependent autophosphorylation of the Pr and Pfr forms of oat phyA in the absence or presence of histone H1. In these reactions, 4 pmol of the Pr or Pfr form of phyA was used with or without 2 μg of histone H1. (C) 32P incorporation in oat phyA by autophosphorylation shown in (B). Each analysis was repeated three times. Error bar = SD (n = 3). We further investigated phyA autophosphorylation over a time course (Fig. 1B, C). The results of these experiments showed that the phyA autophosphorylation was slightly higher in the Pr form than in the Pfr form, but the difference was not significant. In the presence of histone H1, 32P incorporation was stimulated >2-fold, in a Pr-specific manner, whereas 32P incorporation of the Pfr form increased about 1.2-fold. The analysis of these time course experiments revealed that autophosphorylation increased with time and reached a maximum value after 60 min in the absence of histone H1 and after approximately 120 min in the presence of histone H1. Therefore, our results showed that oat phyA possessed autophosphorylation activity and that the autophosphorylation could be stimulated in the presence of histone H1, and thereby confirmed phyA as an autophosphorylating protein kinase. The sites of PhyA autophosphorylation reside in the NTE region Since three phosphorylation sites were known to exist in oat phyA (Lapko et al. 1999), we sought to identify the autophosphorylation site(s) of oat phyA by generating serine to alanine mutants of the known phosphorylation sites. Previously, we reported that S599 is not the site of autophosphorylation (Kim et al. 2004), and the serine-rich NTE region has been proposed to be phosphorylated by phytochrome itself or by a phytochrome-associated kinase (McMichael and Lagarias 1990, Lapko et al. 1999, Kim et al. 2005). Thus, we first examined the autophosphorylation of NTE-deleted Δ65-phyA, in which two known phosphorylation sites, Ser8 and Ser18, were deleted (Fig. 2A). We found that phyA autophosphorylation was not detected in the NTE-deleted Δ65-phyA mutant, even in the presence of histone H1, which suggests that the autophosphorylation site(s) of phyA reside in the NTE region. Although Δ65-phyA did not exhibit any autophosphorylation, it was still able to show the phosphotransfer activity onto histone H1. This result indicates that the NTE is necessary for the phyA autophosphorylation, but not for the phyA kinase activity. Fig. 2 View largeDownload slide Determination of autophosphorylation sites in oat phyA. (A) Autophosphorylation of full-length (FL, amino acids 1–1,129) and NTE-deleted (Δ65, amino acids 66–1,129) phyA in the absence or presence of histone H1. (B) Autophosphorylation of serine to alanine mutants of three known phosphorylation sites. (C) Autophosphorylation of a double site mutant S8/18A. Fig. 2 View largeDownload slide Determination of autophosphorylation sites in oat phyA. (A) Autophosphorylation of full-length (FL, amino acids 1–1,129) and NTE-deleted (Δ65, amino acids 66–1,129) phyA in the absence or presence of histone H1. (B) Autophosphorylation of serine to alanine mutants of three known phosphorylation sites. (C) Autophosphorylation of a double site mutant S8/18A. We next investigated the autophosphorylation of the serine to alanine mutants (Fig. 2B, C). The results showed that the autophosphorylation of the S8A and S18A mutants was significantly reduced, but that the S599A mutant exhibited a similar level of autophosphorylation to wild-type phyA (Fig. 2B). When the autoradiogram bands were quantified with a phosphoimage analyzer, we found that the intensities of both autophosphorylated S8A and S18A were reduced to approximately half the levels seen in wild-type phyA. Furthermore, the S8/18A double mutant showed little autophosphorylation (Fig. 2C). Thus, our results showed that Ser8 and Ser18, which are located in the NTE, are the autophosphorylation sites of oat phyA. To our knowledge, this is the first direct determination of the exact autophosphorylation sites of phyA. Autophosphorylation of PhyA is involved in the regulation of light responses To investigate the functional role of phyA autophosphorylation, we produced homozygous lines of transgenic phyA- deficient Arabidopsis thaliana (phyA-201) plants that expressed the autophosphorylation site mutants as well as wild-type oat phyA. Since it is known that phytochrome function has strong dependency on the amounts of photoreceptor (Boylan and Quail 1991, Whitelam et al. 1993, Wagner and Quail 1995), we first selected transgenic lines overexpressing wild-type phyA whose expression level of oat phyA is similar to that of transgenic lines overexpressing autophosphorylation site mutants for the proper comparison of light responses. To do this, we performed reverse transcription–PCR (RT–PCR) and Western blot analyses to detect the transcript and protein levels in the transgenic lines overexpressing wild-type phyA (Supplementary Fig. S2). For the Western blot analysis, we included the proteasome inhibitor MG-132 in the seedling growth medium and protein extraction buffer to prevent any possible phyA protein degradation during the preparation of plant crude extracts. We found that the Wt-OX6 line showed similar transcript and protein levels of oat phyA to the transgenic lines overexpressing autophosphorylation site mutants (Fig. 3A, B). Among the transgenic plants with wild-type oat phyA, the Wt-OX47 line expressed the highest amount of phyA because of strong transcriptional expression (Supplementary Fig. S2). Thus, we used the Wt-OX6 line for the comparison of light responses with the transgenic plants of autophosphorylation site mutants in the present study. In addition, we included Wt-OX47 as a strong phyA expressor in the experiments of light responsiveness and protein degradation analysis. Fig. 3 View largeDownload slide Seedling de-etiolation responses of transgenic Arabidopsis plants expressing the autophosphorylation site mutants under FR light. (A) RT–PCR analysis to show the transcript levels of oat phyA in transgenic plants. phyA-201, phyA-deficient Arabidopsis (Ler ecotype); Ler, wild-type Arabidopsis; Wt-OX6, transgenic Arabidopsis transformed with wild-type oat phyA; S8A, S18S and S8/18A, transgenic Arabidopsis transformed with the corresponding autophosphorylation site mutant. Numbers represent independent homozygous lines. Actin (ACT2) is shown as a loading control. (B) Western blot analysis to show the protein levels of oat phyA in transgenic plants. For this analysis, seedlings were cultured on MG-132-containing media, and crude proteins were extracted with an MG-132-containing buffer. Oat phyA-specific oat25 antibody was used for the detection of oat phyA, and β-tubulin (TUB) is shown as a loading control. S, protein size standards. (C) Hypocotyl de-etiolation of representative seedlings grown under continuous FR light conditions. 0.5 and 5.0 in parenthesis are the fluence rates (μmol m−2 s−1) used in the analyses. Bar = 5.0 mm. (D) The average hypocotyl lengths of seedlings in (C). Seedlings (n ≥ 29) were grown for 4 d in 1/2 MS medium under fluence rates of 0.5 or 5.0 μmol m−2 s−1, or in darkness. Data are the means ± SD. (E) FR fluence-rate response curves for inhibition of hypocotyl growth. Data are the means (n ≥ 29) ± SD. Fig. 3 View largeDownload slide Seedling de-etiolation responses of transgenic Arabidopsis plants expressing the autophosphorylation site mutants under FR light. (A) RT–PCR analysis to show the transcript levels of oat phyA in transgenic plants. phyA-201, phyA-deficient Arabidopsis (Ler ecotype); Ler, wild-type Arabidopsis; Wt-OX6, transgenic Arabidopsis transformed with wild-type oat phyA; S8A, S18S and S8/18A, transgenic Arabidopsis transformed with the corresponding autophosphorylation site mutant. Numbers represent independent homozygous lines. Actin (ACT2) is shown as a loading control. (B) Western blot analysis to show the protein levels of oat phyA in transgenic plants. For this analysis, seedlings were cultured on MG-132-containing media, and crude proteins were extracted with an MG-132-containing buffer. Oat phyA-specific oat25 antibody was used for the detection of oat phyA, and β-tubulin (TUB) is shown as a loading control. S, protein size standards. (C) Hypocotyl de-etiolation of representative seedlings grown under continuous FR light conditions. 0.5 and 5.0 in parenthesis are the fluence rates (μmol m−2 s−1) used in the analyses. Bar = 5.0 mm. (D) The average hypocotyl lengths of seedlings in (C). Seedlings (n ≥ 29) were grown for 4 d in 1/2 MS medium under fluence rates of 0.5 or 5.0 μmol m−2 s−1, or in darkness. Data are the means ± SD. (E) FR fluence-rate response curves for inhibition of hypocotyl growth. Data are the means (n ≥ 29) ± SD. After selecting two independent homozygous lines of each autophosphorylation site mutant transgenic plant that showed a level of phyA transcript and protein expression comparable with the Wt-OX6 transgenic line, we investigated the light responses of the transgenic plants. Since phyA is known to participate exclusively in the FR-induced inhibition of hypocotyl elongation (Fankhauser and Casal 2004), we first investigated the seedling de-etiolation response of the homozygous transgenic lines in continuous FR (cFR) light with fluence rates of 0.5 and 5.0 μmol m−2 s−1, or in darkness. We measured the hypocotyl lengths of the transgenic lines and wild-type seedlings, and found that all of the transgenic lines (i.e. S8A, S18A and S8/18A) had shorter hypocotyls than non-transformed wild-type Arabidopsis (Ler), or the Wt-OX6 (Fig. 3C, D). When we compared the hypocotyl lengths of the transgenic lines with the strong phyA-overexpressing line Wt-OX47, the transgenic lines still had slightly shorter hypocotyls than Wt-OX47. In particular, the S8/S18 transgenic lines showed much shorter hypocotyls than Wt-OX47. These results suggest that the autophosphorylation site mutants of phyA were hyperactive, and therefore increased the sensitivity of the seedlings to the FR light. Since the hypocotyls of the S8/18A lines were much shorter than those of S8A and S18A, our results suggest that S8 and S18 phosphorylation might have an additive effect on the function of phyA. The observations that the autophosphorylation activity of S8A and S18A was reduced to half that of wild-type phyA, and that of S8/18A was barely detectable, suggest that there is a negative relationship between the magnitude of autophosphorylation and photoresponses in de-etiolation. Furthermore, FR fluence-rate response curves for inhibition of hypocotyl growth confirmed that transgenic seedlings of autophosphorylation site mutants were more sensitive to FR than were those of control plants (i.e. Ler or Wt-OXs) (Fig. 3E). Again, the S8/18A transgenic lines showed the greatest hypersensitive photoresponses among the tested seedlings including Wt-OX47. These results suggest that reduction of autophosphorylation augments the photoresponse in plants, and thus that phyA autophosphorylation might play a role in inhibiting the function of phyA. This is in agreement with the previous proposal that phytochrome phosphorylation is a mechanism of signal attenuation or desensitization for phytochrome- mediated light signaling (Jordan et al. 1996, Casal et al. 2002, Ryu et al. 2005). PhyA mediates a series of photoresponses to cFR, including inhibition of hypocotyl growth, unfolding of the cotyledons, accumulation of anthocyanin and blocking of subsequent greening under white light (Chen et al. 2004). PhyA also mediates two distinct photobiological responses in plants, the very low fluence responses (VLFRs) and the high irradiance responses (HIRs). The VLFR can be achieved by short intermittent pulses of FR [i.e. pulsed FR (pFR)], while the HIR involves the sustained activation of phyA in response to higher fluences of FR (Casal et al. 2002). Since the VLFRs and FR-HIRs are regulated by phyA activity in plants, we analyzed the VLFRs of transgenic seedlings by hourly treatment with FR (i.e. pFR, 24 μmol m−2 s−1 FR : dark = 5 : 55 min), and also the HIRs with continuous FR (2 μmol m−2 s−1) (Fig. 4A). The results showed that all the transgenic lines expressing autophosphorylation site mutants had increased inhibition of hypocotyl growth compared with Ler or Wt-OX6, and that the S8/18A double mutant exhibits the greatest inhibition of hypocotyl elongation in response to cFR and pFR. Therefore, phyA autophosphorylation is involved in the regulation of both VLFRs and HIRs. Fig. 4 View largeDownload slide Photoresponse analysis of transgenic Arabidopsis seedlings expressing the autophosphorylation site mutants. (A) Inhibition of hypocotyl growth under cFR or pFR (i.e. hourly FR). Four-day-old seedlings were exposed to continuous FR (2 μmol m−2 s−1) or hourly FR (24 μmol m−2 s−1 FR : dark = 5 : 55 min). Data are means (n ≥ 29) ± SD. (B) Accumulation of anthocyanin in seedlings grown in the dark, with cFR or pFR light. This assay was repeated three times. Data are the means ± SD (n = 45). (C) Expression of the light-inducible CHS gene in transgenic plants. Seedlings were cultured for 3 d in the dark and transferred to FR (10 μmol m−2 s−1) for 18 h. Total RNA was extracted and RT–PCR analyses were performed using CHS-specific primers. The relative intensities of bands (Irel) are shown beneath the gel, where the band intensity of Ler was set at 1.0. (D) Blocking of greening under FR light. The amount of chlorophyll was measured in 30 seedlings of each line. Data are means (n ≥ 30) ± SD. Seedlings were cultured in the dark, under cFR or pFR light for 3 d, and then transferred to white light (150 μmol m−2 s−1) for 1 d before analyzing the chlorophyll content. Fig. 4 View largeDownload slide Photoresponse analysis of transgenic Arabidopsis seedlings expressing the autophosphorylation site mutants. (A) Inhibition of hypocotyl growth under cFR or pFR (i.e. hourly FR). Four-day-old seedlings were exposed to continuous FR (2 μmol m−2 s−1) or hourly FR (24 μmol m−2 s−1 FR : dark = 5 : 55 min). Data are means (n ≥ 29) ± SD. (B) Accumulation of anthocyanin in seedlings grown in the dark, with cFR or pFR light. This assay was repeated three times. Data are the means ± SD (n = 45). (C) Expression of the light-inducible CHS gene in transgenic plants. Seedlings were cultured for 3 d in the dark and transferred to FR (10 μmol m−2 s−1) for 18 h. Total RNA was extracted and RT–PCR analyses were performed using CHS-specific primers. The relative intensities of bands (Irel) are shown beneath the gel, where the band intensity of Ler was set at 1.0. (D) Blocking of greening under FR light. The amount of chlorophyll was measured in 30 seedlings of each line. Data are means (n ≥ 30) ± SD. Seedlings were cultured in the dark, under cFR or pFR light for 3 d, and then transferred to white light (150 μmol m−2 s−1) for 1 d before analyzing the chlorophyll content. Accumulation of anthocyanins in response to light is also mediated by phyA, which up-regulates the expression of the chalcone synthase gene (CHS) upon irradiation with FR light. We investigated the relative amounts of anthocyanin in seedlings cultured for 3 d under dark, cFR and pulsed FR (FR : dark = 5 : 55 min), and found that anthocyanin accumulation was elevated in response to both cFR and pFR in the transgenic lines expressing autophosphorylation site mutants (Fig. 4B). The extent of induction of CHS was positively correlated with the results of anthocyanin accumulation (Fig. 4C). Furthermore, the results of another HIR to phyA, the blocking of greening under white light, showed that the autophosphorylation site mutants inhibited white light-induced greening significantly more than did Ler or Wt-OX6 in the same light conditions (Fig. 4D). Together, these results suggest that the autophosphorylation site mutants are hyperactive in plants. In addition, the S8/18A mutant exhibited a synergistic increase in the intensity of light responses compared with S8A or S18A. Thus, phosphorylation of both S8 and S18 sites is important for mediating phyA-regulated light responses, including VLFR and FR-HIR. The increased phyA activity in these mutants is related to the decreased extent of phyA autophosphorylation. Overall, our results indicate that phyA autophosphorylation plays an inhibitory role in phytochrome signaling. Autophosphorylation controls the stability of the PhyA protein Previous studies with PAPP5, a phytochrome-interacting protein phosphatase in Arabidopsis, showed that phytochrome stability is increased in PAPP5-overexpression lines, and decreased in papp5 knockout lines, suggesting that phytochrome phosphorylation is involved in regulating the stability of the phytochrome (Ryu et al. 2005). This indicates that dephosphorylated phytochrome is more stable than phosphorylated phytochrome. Furthermore, since phosphorylation at Ser599 does not affect the stability of phyA (Kim et al. 2004), phosphorylation at Ser8 and Ser18 in the NTE have been proposed to be important for protein stability (Kim et al. 2005). Thus, we investigated whether protein stability of the autophosphorylation site mutants was increased by analyzing in vivo protein degradation of phyA proteins upon red light irradiation (28 μmol m−2 s−1) in a light-emitting diode (LED) growth chamber. We found that the autophosphorylation site mutants were more stable than wild-type phyA (Fig. 5). The phyA protein is known to be degraded quickly upon red light irradiation (Henriques et al. 2009), and our results showed that most of the wild-type phyA proteins were degraded within 1 h of red light irradiation, even in the strong phyA-overexpressing Wt-OX47 line. However, the autophosphorylation mutant phyA proteins were degraded much more slowly, and remained even after 3 h of red light irradiation. In addition, S8/18A phyA proteins were more stable than S8A and S18A, indicating the close relationship between autophosphorylation and protein stability of phyA. Interestingly, during our analysis of the amount of phyA in seedlings, we found that the autophosphorylation mutant phyA proteins were not completely degraded in seedlings grown even in white light conditions (Supplementary Fig. S3). We examined the phyA protein levels in dark-grown or white light-grown transgenic seedlings, and found that the autophosphorylation mutant phyA was not completely degraded in seedlings grown for 2 weeks under white light conditions in that wild-type phyA was not detected. Taken together, the loss of autophosphorylation appears to increase the protein stability of phyA, even under conditions of prolonged light irradiation. Fig. 5 View largeDownload slide PhyA protein stability in transgenic plants expressing the autophosphorylation site mutants. (A) Time-dependent degradation of phyA proteins. Three-day-old dark-grown seedlings of transgenic plants were irradiated with continuous red light (28 μmol m−2 s−1) for the indicated periods of time. A 40 μg aliquot of crude extract was loaded onto SDS–polyacrylamide gels and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. (B) Protein degradation graphs of phyA shown in (A). The time-dependent degradation experiments were performed three times, and the averages are shown with the SD. Fig. 5 View largeDownload slide PhyA protein stability in transgenic plants expressing the autophosphorylation site mutants. (A) Time-dependent degradation of phyA proteins. Three-day-old dark-grown seedlings of transgenic plants were irradiated with continuous red light (28 μmol m−2 s−1) for the indicated periods of time. A 40 μg aliquot of crude extract was loaded onto SDS–polyacrylamide gels and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. (B) Protein degradation graphs of phyA shown in (A). The time-dependent degradation experiments were performed three times, and the averages are shown with the SD. To examine further the stability of the phyA protein, green fluorescent protein (GFP)-fused phyA constructs of autophosphorylation site mutants were transiently expressed in mesophyll cell protoplasts, and degradation of GFP-fused phyA was examined under red light irradiation. During light-induced degradation of phyA, phyA exhibits rapid light-dependent aggregation, forming sequestered areas of phytochrome (SAPs) before its degradation via the ubiquitin–26S proteasome pathway (Eichenberg et al. 1999). We also observed the formation of SAPs upon red light irradiation (Fig. 6). In the case of wild-type phyA, SAPs were observed even in the dark, and their size transiently increased and then rapidly disappeared upon red light irradiation. On the other hand, the formation of SAPs was delayed in the autophosphorylation mutants, and they disappeared more slowly than those derived from wild-type phyA. In particular, S8/18A generated SAPs much more slowly than the other phyA mutants, suggesting that the retarded degradation of autophosphorylation mutant phyA proteins in the plant cells is due to the delayed formation of SAPs. Collectively, our results demonstrate that phyA autophosphorylation plays an important role in the regulation of plant phytochrome signaling via the control of phyA protein stability. Fig. 6 View largeDownload slide Analysis of formation of SAPs in mesophyll protoplasts transfected with the autophosphorylation site mutants. GFP-fused phyA constructs were transfected into Arabidopsis mesophyll protoplasts, and phyA proteins were detected with a Leica TCS SP5 confocal microscope upon red light (10 μmol m−2 s−1) irradiation for the indicated periods of time. Bar = 10 μm. Fig. 6 View largeDownload slide Analysis of formation of SAPs in mesophyll protoplasts transfected with the autophosphorylation site mutants. GFP-fused phyA constructs were transfected into Arabidopsis mesophyll protoplasts, and phyA proteins were detected with a Leica TCS SP5 confocal microscope upon red light (10 μmol m−2 s−1) irradiation for the indicated periods of time. Bar = 10 μm. Discussion Autophosphorylation sites of oat PhyA The phytochrome molecule is known to consist of two structural domains, the globular N-terminal chromophore-binding domain (∼65 kDa) and the conformationally open or extended C-terminal domain (∼55 kDa) (Rockwell et al. 2006). The two domains are connected via a flexible hinge region. The N-terminal domain is necessary and sufficient for photoperception and possesses a few subdomains, including the NTE and a bilin lyase domain, while the C-terminal domain contains Per-Arnt-Sim (PAS)-related domains and a histidine kinase-related domain (HKRD), and is necessary for phytochrome dimerization and nuclear localization (Kim et al. 2005, Han et al. 2007). The present study shows that the autophosphorylation sites of phyA reside in the NTE (Fig. 2). The first ∼65 amino acids of the protein (i.e. the NTE) are known to be dispensable for chromophore binding, but necessary for biological activity (Cherry et al. 1992, Jordan et al. 1997). In addition, serine to alanine substitutions in the first 10 serine residues (including Ser8 and Ser18) of rice phyA and deletion of amino acids 6–12 (including Ser8) of oat phyA produce hyperactive phyA, which has higher biological activity than the corresponding wild-type phyA, suggesting that phosphorylation in this region is involved in the down-regulation of phyA activity, such as desensitization (Stockhaus et al. 1992, Jordan et al. 1996). Therefore, our results of the autophosphorylation sites in the NTE are in good agreement with these previous reports. When we compared the autophosphorylation between the Pr and Pfr forms in the absence of histone H1, we could not detect any significant difference between the two forms. However, the Pr form is slightly more phosphorylated than the Pfr form (Fig. 1A). These results are somewhat different from those of a previous report, in that phyA is autophosphorylated in a Pfr-preferential manner (Yeh and Lagarias 1998). This discrepancy is probably due to the low protein kinase activity of phyA compared with other known kinases, such as protein kinase A. This weak autophosphorylation activity of phytochromes may account for the difference between our results and previous results, and may also underlie the long controversy regarding whether or not phytochrome is a protein kinase (Fankhauser 2000, Kim et al. 2005). The weak protein kinase activity of phyA might also indicate the existence of phyA kinase-stimulating molecules, such as polycationic proteins, which include histones. Indeed, phytochrome autophosphorylation and kinase activity are stimulated by the addition of histone H1 (Fig. 1B). Interestingly, the Pr form of phyA, which is considered as the biologically inactive form of phytochrome, is preferentially stimulated by histone H1. Previous studies also reported Pr-specific phytochrome phosphorylation (McMichael and Lagarias 1990, Biermann et al. 1994). Therefore, phytochrome kinase activity might be a Pr-specific kinase that could be stimulated by other factors such as polycationic proteins including histone H1. However, the significance of the stimulation of phytochrome kinase activity in the presence of polycationic proteins needs to be investigated in the future. In addition, we still do not know about the kinase domain of phytochrome, although phytochrome exhibits autophosphorylation and phosphotransfer activities. In an earlier study of affinity labeling of oat phytochrome with ATP analogs, two polypeptide sequences, Glu-Leu-Glu-Lys-Gln-Leu-Arg-Glu-Lys-Asn-Ile-Leu-Lys (residues 403–415) and Asp-Leu-Lys-Leu-Asp-Gly-Leu-Ala (residues 606–613), were suggested to be similar to peptide sequences found within the nucleotide-binding sites of known protein kinases (Wong and Lagarias 1989). In the C-terminal domain of phytochrome, a HKRD exists, but it has been suggested that this HKRD is not a functional kinase domain because the key conserved residues within the histidine kinase domain (HKD) are absent in the phytochrome HKRD (Quail 1997). A recent study of the structure of cyanobacterial phytochrome revealed that the structure of the N-terminal domain is similar to adenylyl cyclase, consisting of PAS (Per-Arnt-Sim)/GAF (cGMP phosphodiesterase/adenylyl cyclase/FhlA)/GAF domains (Essen et al. 2008). Together with an earlier study (Wong and Lagarias 1989), this result suggest that the N-terminal domain of phytochrome might be able to bind to cAMP or possibly to ATP. However, further studies are necessary to determine the phytochrome kinase domain and the catalytic sites. From the present studies, the autophosphorylation sites have been clearly determined as being Ser8 and Ser18 in the NTE (Fig. 2). The sites determined are consistent with the previous results of serine to alanine mutants in the NTE and the studies of in vivo and in vitro phosphorylation sites of phyA (Stockhaus et al. 1992, Lapko et al. 1999). There are three known phosphorylation sites on oat phyA. When we analyzed the amino acid sequences of the sites, a pattern of amino acids, –RXXS–, can be found from Ser8 (–RPAS–), Ser18 (–RQSS–) and Ser599 (–REAS–), in which the phosphorylated serines are underlined. Therefore, the phosphorylation sites of oat phyA contain a conserved pattern of amino acid sequences. Recently, it was reported that Arabidopsis phyA, with a deletion of amino acids 6–12, showed hypoactive responses to cFR light, because of enhanced destruction of the phyA mutant protein (Trupkin et al. 2007), which contrasts with our results. In this report, the deleted amino acid sequence is at 6–12 in Arabidopsis phyA [–(R)PTQSSEG–], which does not include the –RXXS– sequence, whereas the corresponding phosphorylation sequence of oat phyA at 6–12 is –(R)PASSSSS–. This indicates that there might be no autophosphorylation site in amino acids 6–12 of Arabidopsis phyA, and that the difference in phyA sequences of oat and Arabidopsis may account for the discrepancy in the results. Therefore, it appears that the hypoactivity and enhanced degradation of the 6–12 deletion mutant phyA is not due to autophosphorylation, but rather due to another reason, such as a deletion of seven amino acids changing the conformation of the protein. However, we cannot rule out the possibility that the amino acid sequence that mediates autophosphorylation of the dicot (i.e. Arabidopsis) phyA is different from that of the monocot (i.e. oat) phyA. Many studies of the NTE of phyA have suggested that the NTE is important for the biological activity of phyA (Cherry et al. 1992, Stockhaus et al. 1992, Jordan et al. 1996, Jordan et al. 1997, Casal et al. 2002, Trupkin et al. 2007). Most of these studies suggest that the NTE might have a role in signal attenuation or desensitization in the regulation of photoresponses. Furthermore, modifications in the NTE, including the 6–12 deleted Arabidopsis phyA (Trupkin et al. 2007), influence the degradation of phyA. The present study provides the first direct evidence for the existence of autophosphorylation sites in the NTE. Both Ser8 and Ser18 in the NTE of oat phyA are sites of autophosphorylation. The autophosphorylation in S8A or S18A was each reduced to approximately half of wild-type phyA autophosphorylation levels (Fig. 2B), suggesting that there might be no autophosphorylation preference between the sites. It is noteworthy that the S8/18A double mutant showed almost no autophosphorylation (Fig. 2C), indicating that only two autophosphorylation sites exist on oat phyA. To our knowledge, our report is the first to have determined the exact sites of phyA autophosphorylation, and also confirmed that phyA contained two autophosphorylation sites. Therefore, the NTE of phyA might play a role in the regulation of phytochrome signaling through autophosphorylation, and it is possible that an initial event in phytochrome-mediated signaling involves a post-translational modification of the photoreceptor itself. Functional roles of PhyA autophosphorylation We investigated the functional roles of phyA autophosphorylation by using transgenic plants expressing the autophosphorylation site mutants, S8A, S18A and S8/18A. Our results showed that all of the autophosphorylation site mutants were hyperactive compared with wild-type oat phyA, in terms of FR light sensitivity (Fig. 3), inhibition of hypocotyl growth under cFR and pFR (Fig. 4A), accumulation of anthocyanin (Fig. 4B, C) and blocking of greening (Fig. 4D). The S8/18A mutant was more hyperactive than S8A or S18A, suggesting that Ser8 and Ser18 phosphorylation had cumulative effects. Our results are consistent with previous results from serine to alanine substitution and Δ6–12 deletion mutants, in which both mutants were hyperactive in transgenic plants (Stockhaus et al. 1992, Casal et al. 2002). Therefore, these results suggest that phytochrome autophosphorylation plays a role in the negative regulation of phyA signaling through signal attenuation or desensitization. Previously, phytochrome phosphorylation was suggested to be important for degradation of the phytochrome protein. For example, PAPP5 knockout plants showed a decrease in photoresponsiveness and a decline in the stability of the phyA protein (Ryu et al. 2005). On the other hand, the protein stability of phyA increased in plants overexpressing PAPP5. These results suggest that phosphorylation of phyA accelerates the degradation of phyA, whereas the dephosphorylation of phyA slows the degradation of phyA. Our results confirmed that the protein stability of the autophosphorylation site mutants increased, and thus slowly degraded in the transgenic plants (Fig. 5). The S8/18A mutant was degraded much more slowly than the single site mutants and the wild-type phyA, which could explain the increased photoresponsiveness in the transgenic plants. Even in the white light conditions, in which wild-type phyA was completely degraded, the autophosphorylation site mutants were not completely degraded, but remained in small amounts (Supplementary Fig. S3). Therefore, phyA autophosphorylation is necessary not only for the rapid degradation of phyA, but also for complete degradation under light conditions. Recently, it has been reported that phosphorylated phyA preferentially associates with the COP1–SPA1 complex, the E3 ligase involved in the degradation of phyA (Saijo et al. 2008). This study suggests that phyA phosphorylation is important for its interaction with the COP1–SPA1 complex and shows that the phosphorylated form of phyA is enriched during co-immunoprecipitation with COP1. Therefore, our results are in good agreement with this report, in that the autophosphorylation site mutants of phyA are not efficiently degraded, possibly because of decreased interaction of the mutants with the COP1–SPA1 complex. PhyA degradation has been further investigated using protoplast transfection assays (Fig. 6). PhyA is biosynthesized in the cytosol as the Pr form in the dark and accumulates continuously under illumination. Upon illumination, most phyA is rapidly degraded in the cytosol by forming SAPs, and some phyA is translocated into the nucleus to initiate signaling cascades. The light-induced formation of SAPs is considered to be a molecular property of phyA, although the exact function of the formation SAPs is not fully known (Eichenberg et al. 1999). Thus, we observed the formation of SAPs by the autophosphorylation site mutants and found that these mutants formed SAPs more slowly than wild-type phyA, which exactly matched the results of the protein degradation assays. Since the SAPs are considered to be the functional complex for phyA degradation, the slow formation of SAPs is correlated with the slow degradation of phyA. Since the phyA accumulated in the dark for the sensitive light signaling may need to be degraded rapidly for their desensitization, the SAPs might be formed for the quick and efficient degradation of phyA, in which autophosphorylation is necessary for the formation of SAPs. Therefore, underphosphorylation of phyA in the mutants might delay the formation of SAPs and thus slow the degradation of phyA, which could sustain the activity of phyA and impart increased photoresponses to FR light in the plants. When we consider the findings that phytochrome autophosphorylation occurs similarly in the Pr and Pfr forms and the phosphorylated phyA in the Pfr form is degraded in vivo, phyA might be equally autophosphorylated in vivo between the Pr and Pfr forms and the photoconversion from Pr to Pfr might not be significantly affected by the autophosphorylation. The finding that phyA is down-regulated by phosphorylation might be similar to the case for G-protein-coupled receptors. As an example, rhodopsin is desensitized by phosphorylation through rhodopsin kinase (Arshavsky 2002, Sokal et al. 2002). The desensitization of rhodopsin is important for vision. If the activation of rhodopsin is not attenuated efficiently, blindness will occur. However, the signal attenuation mechanism of rhodopsin might be different from that of phytochrome: activation of rhodopsin results in its phosphorylation and subsequent binding to arrestin-like proteins, whereas phytochrome phosphorylation and its activation results in acceleration of its degradation through binding with an E3 ligase complex (i.e. COP1–SPA1). Overall, our results provide a plausible model for the functional role of phyA autophosphorylation. In the dark, phyA proteins are synthesized and accumulate as the Pr form in the cytosol, consisting of autophosphorylated phyA and dephosphorylated phyA, due to the action of protein phosphatases. Upon illumination, the Pr form is photoactivated to the Pfr form, which can be degraded via the ubiquitin–26S proteasome protein degradation pathway. At this point, the phosphorylated Pfr form is rapidly degraded, compared with the unphosphorylated phyA species. The balance between the autophosphorylated and dephosphorylated phyA proteins is probably shifted to the autophosphorylated phyA, because the phosphorylated phyA proteins are efficiently degraded by the ubiquitin–26S proteasome through the COP1–SPA1 complex. This rapid phyA degradation mechanism that involves autophosphorylation might facilitate the efficient desensitization of the phyA signal. Otherwise, phytochrome protein phosphatases might increase the phyA protein stability with enhanced light responses by decreasing the amounts of the autophosphorylated phyA. If phyA activity is not attenuated upon illumination, plants could over-respond to light by eliciting too many photomorphogenic signaling events and/or could become desensitized to subsequent changes in light quality or quantity. Therefore, this regulation may provide an efficient means of rapidly controlling the number of active phyA photoreceptors available to initiate signaling events, and thereby improve the response of plants to fluctuating light environments. Materials and Methods Phytochrome constructs Full-length cDNA (amino acids 1–1,129) of Avena sativa (oat) phyA was cloned into pGEM®-11zf(+) (Promega) from pFY122 (Boylan and Quail 1989) by digestion with BamHI and EcoRI, and was used as a template for mutagenesis and PCRs. Each of the two known phosphorylation sites on oat phyA (i.e. Ser8 and Ser18) was mutagenized to alanine by site-directed mutagenesis using the GeneEditor™ in vitro Site-Directed Mutagenesis System (Promega). The mutagenic primers used were phosphorylated-5′-CCTCAAGGCCTGCAGCCAGTTCTTCCA GC-3′ for Ser8Ala (S8A), and phosphorylated-5′-GGAACCG CCAGAGTGCGCAGGCAAGGGTG-3′ for Ser18Ala (S18A). After the mutagenesis of S8A and S18A, the second mutagenesis was carried out to make a combination mutant Ser8Ala/Ser18Ala (S8/18A). After mutagenesis, all the constructs were confirmed by DNA sequencing. In addition to these S8A and S18A mutants, the NTE was deleted to generate the Δ65 mutant (amino acids 66–1,129), and the S599A mutant was obtained as previously described (Kim et al. 2004). All the constructs were then cloned into pPIC3.5K for the preparation of recombinant proteins using the Pichia Protein Expression System (Invitrogen) or into the pBI121 binary plasmid (Clontech) for the generation of transgenic plants by transformation into phyA-deficient A. thaliana (phyA-201, Ler ecotype), as described (Kim et al. 2004). The constructs in pPIC3.5K contained the streptavidin affinity tag (strep-tag), consisting of 10 amino acids at the C-terminus for efficient purification by streptavidin affinity chromatography. Plant materials and growth conditions The constructs S8A, S18A, S8/18A and wild-type PHYA in pBI121 were each introduced into phyA-201 using the Agrobacterium tumefaciens (strain GV3101)-mediated floral dip method, as described (Clough and Bent 1998). Transgenic lines segregating ∼3 : 1 for antibiotic resistance in the T2 generation were selected, and the T3 or T4 homozygous generation was used for subsequent analyses. Arabidopsis plants were grown on soil in a culture room (22°C with a 16 h photoperiod), following routine procedures. For seedling assays, seeds were surface-sterilized, incubated at 4°C for 3 d in the dark, and placed on 0.8% phytoagar (w/v) medium containing half-strength MS salts and vitamins. The plates were then transferred to a growth chamber (22°C with a 16 h photoperiod). Expression, reconstitution and purification of PhyA–chromophore holoproteins The pPIC3.5K constructs bearing PHYA genes were transformed into Pichia cells by means of a Micropulser™ Electroporation apparatus (Bio-Rad). Recombinant phytochrome proteins were expressed in the Pichia expression system (Invitrogen) and purified by streptavidin affinity chromatography (IBA), according to the procedure described by the manufacturer. Phycocyanobilin (PCB) was prepared from Spirulina platensis extracts (Sigma) by methanolysis as described (Park et al. 2000), and used as a chromophore for the holo-phytochrome assembly. The concentration of purified PCB was quantified by absorption spectroscopy in HCl (2%)/methanol, using an extinction coefficient (ε) of 37,900 M−1 cm−1 at 690 nm. From the harvested Pichia cells, crude extracts were prepared by breaking cells in liquid nitrogen using a homogenizer (Nihonseiki Kaisha). The phytochrome samples were then precipitated by adding 0.23 g l−1 ammonium sulfate, resuspended in a buffer [100 mM Tris (pH 7.8), 1 mM EDTA], the PCB chromophores in dimethylsulfoxide (DMSO) were added to the samples at a final concentration of 10 μM and the mixture was incubated for in vitro reconstitution on ice for 1 h. After dialysis to remove free chromophores, the samples were loaded onto streptavidin affinity chromatography columns and holo-phytochromes without free chromophores were purified. Zn2+ fluorescence and spectroscopic analysis For Zn2+ fluorescence assays to assess the chromophore ligation, the protein samples were analyzed on a 10% SDS–polyacrylamide gel and the gel was then soaked in 20 mM zinc acetate/150 mM Tris–HCl (pH 7.0) for 5–30 min at room temperature with gentle shaking. Zinc fluorescence of holo-phytochromes was visualized under UV light (312 nm). For the spectroscopic analysis of recombinant phyA, absorption spectra were recorded by a diode array UV/VIS spectrophotometer (Cary) after red or FR light irradiation. All the spectroscopic experiments were carried out under the green safety light condition, which consisted of a white fluorescent lamp equipped with a plastic filter (Rosco) with a maximal transmittance at 500 nm, and a fiberoptic illuminator system (Cole-Parmer) equipped with 656 and 730 nm interference filters (Oriel) was used as a light source. The light intensity was 8 W m−2 for red light and 6 W m−2 for FR light. The samples were illuminated with red or FR light for 5 min. A differential spectrum was obtained by subtracting the Pfr spectrum from the Pr spectrum. The concentrations of the holo-phytochrome samples were calculated from the λmax of the Pr peak with an extinction coefficient of 132,000 M−1 cm−1. The concentrations of other protein samples were determined using the Bradford Protein Assay Kit (Bio-Rad), with bovine serum albumin as a standard. Autophosphorylation assay Phytochrome phosphorylation experiments were performed as described (Yeh and Lagarias 1998, Kim et al. 2004) with minor modifications. The reaction mixtures (total volume of 20 μl) contained kinase buffer [50 mM Tris–HCl (pH 7.8), 0.2 mM EDTA, 4 mM dithiothreitol (DTT) and 5 mM MgCl2] and purified recombinant phytochromes (0.5–2 μg), in either Pr or Pfr. Phytochrome samples were irradiated with red or FR light for 5 min before the start of a reaction. Autophosphorylation reactions were started by adding 0.1 mM ATP containing 10 μCi of [γ-32P]ATP (Perkin-Elmer, 3,000 Ci mmol−1) and were incubated at 30°C for 1 h. Reactions were stopped by the addition of 5 μl of 5× SDS sample buffer. Proteins were resolved on 10% (w/v) SDS–polyacrylamide gels, dried under vacuum and exposed to X-ray films for autoradiography. Coomassie Blue protein staining and zinc fluorescence assays were performed before drying. Because polycations, such as histone H1 (Roche), are known to increase the phytochrome autophosphorylation activity, histone H1 (0.5–2 μg) was added to phytochrome samples in the indicated experiments. For a quantitative analysis of phytochrome phosphorylation, 32P radiolabel incorporation was determined by scintillation counting of the excised protein bands from SDS–polyacrylamide gels as described (Wong et al. 1989). Using a curve of the relationship between the molar concentration of 32P and scintillation counting, 4 pmol of phyA was reacted with 10 μCi of [γ-32P]ATP, and the samples were loaded on SDS–polyacrylamide gels. The molar incorporation of 32P was determined by measuring the 32P concentration in each excised protein band from SDS–polyacrylamide gels by a liquid scintillation counter (Beckman Coulter, Inc.). RT–PCR and Western blot analysis To compare transcript levels of oat phyA from transgenic plants, total RNAs were extracted using a RNeasy® plant kit (Qiagen), and then a PrimeScript™ first-strand cDNA synthesis kit (TAKARA) was used for cDNA synthesis. PCR amplification for oat PHYA was performed with a forward primer at 2,704 bp, 5′-TACATGAGACATGCGATCAAC-3′, and a reverse primer at 3,264 bp, 5′-GCTCAAGCCCTCCTCTGACTG-3′. As a control, ACT2 was amplified with a forward primer, 5′-TTGACCTTG CTGGACGTG-3′, and a reverse primer, 5′-GGAAGCAAGAAT GGAACCAC-3′. To investigate phyA protein levels from transgenic plants, each transgenic line was cultured on half-strength MS medium with vitamins including 0.8% phytoagar and 50 μM MG-132 (A.G. Scientific, Inc.) for 2.5 d in darkness. Approximately 100 dark-grown seedlings were ground and extracted with an extraction buffer [70 mM Tris–HCl, pH 8.3, 7 mM EDTA, 35% ethylene glycol, 98 mM ammonium sulfate, 14 mM sodium metabisulfite, 0.07% polyethyleneimine, 2.8 mM phenylmethylsulfonyl fluoride (PMSF), and 10 μM MG-132]. The extracted protein samples were centrifuged at 14,000 r.p.m. at 4°C for 15 min, and 30 μg of protein samples were used for Western blot analysis. Photoresponse analyses of transgenic Arabidopsis plants To assess the expression of oat phyA and serine to alanine mutant proteins in transgenic Arabidopsis plants, Western blot analysis was performed on 4-day-old dark-grown seedlings as described (Kim et al. 2004). A 40 μg aliquot of crude extract was separated by 10% SDS–PAGE and transferred to a polyvinylidene difluoride (PVDF) membrane (Hybond-P, GE Healthcare). The membrane was then incubated with oat phyA-specific oat25 monoclonal antibody and developed using an ECL Advanced™ Western Blotting Analysis System (GE Healthcare). Hypocotyl lengths were measured in response to light treatments as described (Fankhauser and Casal 2004). The seeds were sown on half-strength MS medium and cold treated for 3 d in the dark. The seeds were then exposed to white light for 4 h to promote germination, returned to darkness (22°C) for 24 h, and then grown for 3 d under cFR or pFR light with various fluence rates, using an LED growth chamber (Vision, Korea). The hypocotyls were photographed with a digital camera (Nikon), and then measured with image analysis software (Scion Image, Frederick, MD, USA). The anthocyanin content and total chlorophyll content of seedlings were determined as described (Fankhauser and Casal 2004). Relative anthocyanin levels were determined by collecting 15 seedlings from each of the light treatments/line and incubating them overnight in 500 μl of methanol acidified with 1% HCl, with shaking, in the dark. The next day, 500 μl of chloroform was added, and the sample was vortexed and briefly centrifuged to separate the anthocyanins from chlorophyll. The total anthocyanin content was determined by measuring the A530 and A657 of the aqueous phase using a UV/VIS spectrophotometer (Cary). The relative amount of anthocyanin per seedling was calculated by subtracting the A657 from the A530. Total chlorophyll was determined from samples containing 15 seedlings. Seedlings were extracted by incubating in 1 ml of 80% acetone, with shaking, overnight in the dark. Chlorophyll levels were measured spectroscopically and the amount of chlorophyll was determined using the equation: chlorophylla + b = 7.15 × OD660 nm + 18.71 × OD647 nm. In vivo phytochrome degradation assay The transgenic plant seeds were germinated, and grown for 3.5 d in the dark. Seedlings were then illuminated with red light (28 μmol m−2 s−1) in the LED growth chamber (Vision Scientific Co., Ltd.) and harvested at the indicated times. The harvested seedlings were stored in liquid nitrogen and the protein samples were prepared as described (Kim et al. 2004). A 40 μg aliquot of crude extract was loaded onto SDS–polyarylamide gels, and Western blot analysis was performed to detect oat phyA, using the oat25 monoclonal antibody. To investigate the in vivo degradation of phyA using mesophyll cell protoplasts, transient expression of GFP-fused phyA mutants as well as wild-type phyA was examined. GFP–phyA constructs were cloned into the pCsVMV:GFP vector under the control of the CsVMV promoter (Verdaguer et al. 1996), using EcoRI and BamHI and primer set, 5′-CGGAATTCATGTCTTCCTCAAGGCCTGCTTCC-3′ (forward) and 5′-CGGGATCCTTGTCCCATTGCTGTTGGAGCGGAAGC-3′ (reverse) for the wild type and S18A phyA, and 5′-CGGAATTCATGTCTTCCTCAAGGCCTGCTGCAAGT-3′ (forward) and the same reverse primer for S8A and S8/18A phyA. Mesophyll cell protoplasts were isolated from phyA-deficient A. thaliana (phyA-211, Col-0 ecotype) and transfected as described (Hwang and Sheen 2001). Transfected protoplasts were incubated overnight in the dark, and then SAPs were observed during red light irradiation for the indicated periods of time using a confocal microscope. These confocal images were obtained using a laser scanning confocal microscope (Leica TCS SP5 AOBS/Tandem) at Korea Basic Science Institute, Gwangju Center. Funding This work was supported by the Plant Diversity Research Center of 21st Century Frontier Research Program [grant no. PF06302–02]; the MEST/NRF to the Environment Biotechnology National Core Research Center (NCRC) [grant no. 20090091493]; the Korea Research Foundation Grant funded by the Korean Government (MOEHRD) [grant no. KRF-2006-312-C00671]. References Arshavsky VY.  Rhodopsin phosphorylation: from terminating single photon responses to photoreceptor dark adaptation,  Trends Neurosci. ,  2002, vol.  25 (pg.  124- 126) Google Scholar CrossRef Search ADS PubMed  Bae G,  Choi G.  Decoding of light signals by plant phytochromes and their interacting proteins,  Annu. Rev. Plant Biol. ,  2008, vol.  59 (pg.  281- 311) Google Scholar CrossRef Search ADS PubMed  Biermann BJ,  Pao LI,  Feldman LJ.  Pr-specific phytochrome phosphorylation in vitro by a protein kinase present in anti-phytochrome maize immunoprecipitates,  Plant Physiol. ,  1994, vol.  105 (pg.  243- 251) Google Scholar CrossRef Search ADS PubMed  Boylan MT,  Quail PH.  Oat phytochrome is biologically active in transgenic tomatoes,  Plant Cell ,  1989, vol.  1 (pg.  765- 773) Google Scholar CrossRef Search ADS PubMed  Boylan MT,  Quail PH.  Phytochrome a overexpression inhibits hypocotyl elongation in transgenic Arabidopsis,  Proc. Natl Acad. Sci. USA ,  1991, vol.  88 (pg.  10806- 10810) Google Scholar CrossRef Search ADS   Casal JJ,  Davis SJ,  Kirchenbauer D,  Viczian A,  Yanovsky MJ,  Clough RC, et al.  The serine-rich N-terminal domain of oat phytochrome a helps regulate light responses and subnuclear localization of the photoreceptor,  Plant Physiol. ,  2002, vol.  129 (pg.  1127- 1137) Google Scholar CrossRef Search ADS PubMed  Chen M,  Chory J,  Fankhauser C.  Light signal transduction in higher plants,  Annu. Rev. Genet. ,  2004, vol.  38 (pg.  87- 117) Google Scholar CrossRef Search ADS PubMed  Cherry JR,  Hondred D,  Walker JM,  Vierstra RD.  Phytochrome requires the 6-kDa N-terminal domain for full biological activity,  Proc. Natl Acad. Sci. USA ,  1992, vol.  89 (pg.  5039- 5043) Google Scholar CrossRef Search ADS   Clough SJ,  Bent AF.  Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana,  Plant J. ,  1998, vol.  16 (pg.  735- 743) Google Scholar CrossRef Search ADS PubMed  Eichenberg K,  Kunkel T,  Kretsch T,  Speth V,  Schafer E.  In vivo characterization of chimeric phytochromes in yeast,  J. Biol. Chem. ,  1999, vol.  274 (pg.  354- 359) Google Scholar CrossRef Search ADS PubMed  Emmler K,  Stockhaus J,  Chua NH,  Schafer E.  An amino-terminal deletion of rice phytochrome A results in a dominant negative suppression of tobacco phytochrome A activity in transgenic tobacco seedlings,  Planta ,  1995, vol.  197 (pg.  103- 110) Google Scholar CrossRef Search ADS PubMed  Essen LO,  Mailliet J,  Hughes J.  The structure of a complete phytochrome sensory module in the Pr ground state,  Proc. Natl Acad. Sci. USA ,  2008, vol.  105 (pg.  14709- 14714) Google Scholar CrossRef Search ADS   Fankhauser C.  Phytochromes as light-modulated protein kinases,  Semin. Cell Dev. Biol. ,  2000, vol.  11 (pg.  467- 473) Google Scholar CrossRef Search ADS PubMed  Fankhauser C,  Casal JJ.  Phenotypic characterization of a photomorphogenic mutant,  Plant J. ,  2004, vol.  39 (pg.  747- 760) Google Scholar CrossRef Search ADS PubMed  Fankhauser C,  Chen M.  Transposing phytochrome into the nucleus,  Trends Plant Sci. ,  2008, vol.  13 (pg.  596- 601) Google Scholar CrossRef Search ADS PubMed  Han YJ,  Song PS,  Kim JI.  Phytochrome-mediated photomorphogenesis in plants,  J. Plant Biol. ,  2007, vol.  50 (pg.  230- 240) Google Scholar CrossRef Search ADS   Henriques R,  Jang IC,  Chua NH.  Regulated proteolysis in light-related signaling pathways,  Curr. Opin. Plant Biol. ,  2009, vol.  12 (pg.  49- 56) Google Scholar CrossRef Search ADS PubMed  Hunt RE,  Pratt LH.  Partial characterization of undegraded oat phytochrome,  Biochemistry ,  1980, vol.  19 (pg.  390- 394) Google Scholar CrossRef Search ADS PubMed  Hwang I,  Sheen J.  Two-component circuitry in Arabidopsis cytokinin signal transduction,  Nature ,  2001, vol.  413 (pg.  383- 389) Google Scholar CrossRef Search ADS PubMed  Jiao Y,  Lau OS,  Deng XW.  Light-regulated transcriptional networks in higher plants,  Nat. Rev. Genet. ,  2007, vol.  8 (pg.  217- 230) Google Scholar CrossRef Search ADS PubMed  Jordan ET,  Cherry JR,  Walker JM,  Vierstra RD.  The amino-terminus of phytochrome A contains two distinct functional domains,  Plant J. ,  1996, vol.  9 (pg.  243- 257) Google Scholar CrossRef Search ADS PubMed  Jordan ET,  Marita JM,  Clough RC,  Vierstra RD.  Characterization of regions within the N-terminal 6-kilodalton domain of phytochrome A that modulate its biological activity,  Plant Physiol. ,  1997, vol.  115 (pg.  693- 704) Google Scholar CrossRef Search ADS PubMed  Kim DH,  Kang JG,  Yang SS,  Chung KS,  Song PS,  Park CM.  A phytochrome-associated protein phosphatase 2A modulates light signals in flowering time control in Arabidopsis,  Plant Cell ,  2002, vol.  14 (pg.  3043- 3056) Google Scholar CrossRef Search ADS PubMed  Kim JI,  Park JE,  Zarate X,  Song PS.  Phytochrome phosphorylation in plant light signaling,  Photochem. Photobiol. Sci. ,  2005, vol.  4 (pg.  681- 687) Google Scholar CrossRef Search ADS PubMed  Kim JI,  Shen Y,  Han YJ,  Park JE,  Kirchenbauer D,  Soh MS, et al.  Phytochrome phosphorylation modulates light signaling by influencing the protein–protein interaction,  Plant Cell ,  2004, vol.  16 (pg.  2629- 2640) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Jiang XY,  Smith DL,  Song PS.  Posttranslational modification of oat phytochrome A: phosphorylation of a specific serine in a multiple serine cluster,  Biochemistry ,  1997, vol.  36 (pg.  10595- 10599) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Jiang XY,  Smith DL,  Song PS.  Mass spectrometric characterization of oat phytochrome A: isoforms and posttranslational modifications,  Protein Sci. ,  1999, vol.  8 (pg.  1032- 1044) Google Scholar CrossRef Search ADS PubMed  Lapko VN,  Wells TA,  Song PS.  Protein kinase A-catalyzed phosphorylation and its effect on conformation in phytochrome A,  Biochemistry ,  1996, vol.  35 (pg.  6585- 6594) Google Scholar CrossRef Search ADS PubMed  McMichael R.W.Jr.,  Lagarias,  J. C.  Phosphopeptide mapping of Avena phytochrome phosphorylated by protein kinases in vitro,  Biochemistry ,  1990, vol.  29 (pg.  3872- 3878) Google Scholar CrossRef Search ADS PubMed  Nagatani A.  Light-regulated nuclear localization of phytochromes,  Curr. Opin. Plant Biol. ,  2004, vol.  7 (pg.  708- 711) Google Scholar CrossRef Search ADS PubMed  Park CM,  Shim JY,  Yang SS,  Kang JG,  Kim JI,  Luka Z, et al.  Chromophore–apoprotein interactions in Synechocystis sp. PCC6803 phytochrome Cph1,  Biochemistry ,  2000, vol.  39 (pg.  6349- 6356) Google Scholar CrossRef Search ADS PubMed  Phee BK,  Kim JI,  Shin DH,  Yoo J,  Park KJ,  Han YJ, et al.  A novel protein phosphatase indirectly regulates phytochrome-interacting factor 3 via phytochrome,  Biochem. J. ,  2008, vol.  415 (pg.  247- 255) Google Scholar CrossRef Search ADS PubMed  Quail PH.  The phytochromes: a biochemical mechanism of signaling in sight?,  Bioessays ,  1997, vol.  19 (pg.  571- 579) Google Scholar CrossRef Search ADS PubMed  Rockwell NC,  Su YS,  Lagarias JC.  Phytochrome structure and signaling mechanisms,  Annu. Rev. Plant Biol. ,  2006, vol.  57 (pg.  837- 858) Google Scholar CrossRef Search ADS PubMed  Ryu JS,  Kim JI,  Kunkel T,  Kim BC,  Cho DS,  Hong SH, et al.  Phytochrome-specific type 5 phosphatase controls light signal flux by enhancing phytochrome stability and affinity for a signal transducer,  Cell ,  2005, vol.  120 (pg.  395- 406) Google Scholar CrossRef Search ADS PubMed  Saijo Y,  Zhu D,  Li J,  Rubio V,  Zhou Z,  Shen Y, et al.  Arabidopsis COP1/SPA1 complex and FHY1/FHY3 associate with distinct phosphorylated forms of phytochrome A in balancing light signaling,  Mol. Cell ,  2008, vol.  31 (pg.  607- 613) Google Scholar CrossRef Search ADS PubMed  Sokal I,  Pulvermuller A,  Buczylko J,  Hofmann KP,  Palczewski K.  Rhodopsin and its kinase,  Methods Enzymol. ,  2002, vol.  343 (pg.  578- 600) Google Scholar PubMed  Stockhaus J,  Nagatani A,  Halfter U,  Kay S,  Furuya M,  Chua NH.  Serine-to-alanine substitutions at the amino-terminal region of phytochrome A result in an increase in biological activity,  Genes Dev. ,  1992, vol.  6 (pg.  2364- 2372) Google Scholar CrossRef Search ADS PubMed  Trupkin SA,  Debrieux D,  Hiltbrunner A,  Fankhauser C,  Casal JJ.  The serine-rich N-terminal region of Arabidopsis phytochrome A is required for protein stability,  Plant Mol. Biol. ,  2007, vol.  63 (pg.  669- 678) Google Scholar CrossRef Search ADS PubMed  Verdaguer B,  de Kochko A,  Beachy RN,  Fauquet C.  Isolation and expression in transgenic tobacco and rice plants, of the cassava vein mosaic virus (CVMV) promoter,  Plant Mol. Biol. ,  1996, vol.  31 (pg.  1129- 1139) Google Scholar CrossRef Search ADS PubMed  Wagner D,  Quail PH.  Mutational analysis of phytochrome B identifies a small COOH-terminal-domain region critical for regulatory activity,  Proc. Natl Acad. Sci. USA ,  1995, vol.  92 (pg.  8596- 8600) Google Scholar CrossRef Search ADS   Whitelam GC,  Johnson E,  Peng J,  Carol P,  Anderson ML,  Cowl JS, et al.  Phytochrome A null mutants of Arabidopsis display a wild-type phenotype in white light,  Plant Cell ,  1993, vol.  5 (pg.  757- 768) Google Scholar CrossRef Search ADS PubMed  Wong YS,  Cheng HC,  Walsh DA,  Lagarias JC.  Phosphorylation of Avena phytochrome in vitro as a probe of light-induced conformational changes,  J. Biol. Chem. ,  1986, vol.  261 (pg.  12089- 12097) Google Scholar PubMed  Wong YS,  Lagarias JC.  Affinity labeling of Avena phytochrome with ATP analogs,  Proc. Natl Acad. Sci. USA ,  1989, vol.  86 (pg.  3469- 3473) Google Scholar CrossRef Search ADS   Wong YS,  McMichael RW,  Lagarias JC.  Properties of a polycation-stimulated protein kinase associated with purified Avena phytochrome,  Plant Physiol. ,  1989, vol.  91 (pg.  709- 718) Google Scholar CrossRef Search ADS PubMed  Yeh KC,  Lagarias JC.  Eukaryotic phytochromes: light-regulated serine/threonine protein kinases with histidine kinase ancestry,  Proc. Natl Acad. Sci. USA ,  1998, vol.  95 (pg.  13976- 13981) Google Scholar CrossRef Search ADS   Abbreviations Abbreviations ACT actin gene cFR continuous far-red CHS chalcone synthase gene FR far-red FyPP flower-specific phytochrome-associated protein phosphatase GFP green fluorescent protein HIR high-irradiance response HKRD histidine kinase-related domain LED light-emitting diode NDPK2 nucleoside di-phosphate kinase 2 NTE N-terminal extension PAPP5 phytochrome-associated protein phosphatase 5 PCB phycocyanobilin Pfr far-red light-absorbing form of phytochrome pFR pulsed far-red phyA phytochrome A PIF3 phytochrome-interacting factor 3 Pr red light-absorbing form of phytochrome RT–PCR reverse transcription–PCR SAPs sequestered areas of phytochrome VLFR very-low- fluence response. © The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org

Journal

Plant and Cell Physiology – Oxford University Press

Published: Apr 8, 2010

Keywords: Autophosphorylation Phytochrome Phosphorylation Protein degradation SAPs

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