Phytochromes are red and far-red light receptors in plants that mediate critical responses to light throughout the lifecycle. They achieve this in part by targeting negatively acting bHLH transcription factors called phytochrome-interacting factors (PIFs) for degradation within the nucleus. However, it is not known whether protein degradation is the primary mechanism by which phytochromes inhibit these repressors of photomorphogenesis. Here, we use chromatin immunoprecipitation to show that phyB inhibits the regulatory activity of PIF1 and PIF3 by releasing them from their DNA targets. The N-terminal fragment of phyB (NG-GUS-NLS; NGB) also inhibits binding of PIF3 to its target promoters. However, unlike full-length phyB, NGB does not promote PIF3 degradation, establishing the activity of NGB reflects its ability to inhibit PIF binding to DNA. We further show that Pfr forms of both full-length phyB and NGB inhibit DNA binding of PIF1 and PIF3 in vitro. Taken together, our results indicate that phyB inhibition of PIF function involves two separate processes: sequestration and protein degradation.
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
Phytochromes are red and far-red light photoreceptors that modulate key physiological and developmental processes of plants, including seed germination, seedling photomorphogenesis, shade avoidance and flowering (Rockwell et al., 2006). At least two branches of phytochromes have evolved to facilitate adaptation to the variation in the light spectrum found in common terrestrial environments (Mathews, 2006; Franklin and Quail, 2010). One branch, typified by Arabidopsis phytochrome B (phyB), mediates so-called low-fluence responses, i.e. those strongly activated by the red light common in open field plantings. The other branch, typified by Arabidopsis phytochrome A (phyA), mediate the very low-fluence response and the far-red high irradiance response: processes that are activated by the far-red light-enriched environments of deep canopy shade and dense crop plantings. Although the distinct response modes of phyA and phyB categories appear to be well conserved in flowering plants, phyA may also function as a low-fluence response sensor depending on the plant species and stage of development (Takano et al., 2005; Franklin et al., 2007). Despite this photosensory diversity, the regulatory gene networks targeted by all classes of plant phytochromes appear to significantly overlap and share common regulatory factors.
Phytochrome-interacting proteins play a key role in assisting all classes of phytochromes to regulate light responses. Some interacting proteins regulate specific phytochrome activity by modulating their subcellular localization, their stability or their light-independent thermal reversion (dark reversion) to inactive species, whereas others regulate light responses directly under phytochrome control (Bae and Choi, 2008). The most well-studied of these are the so-called phytochrome-interacting bHLH transcription factors, termed PIFs or PILs (PIF3-like proteins), which primarily function as negative regulators of photomorphogenesis singly and/or in combination (Leivar et al., 2008; Shin et al., 2009; Leivar and Quail, 2011). Seed germination is one example of a light response that is mainly regulated by a single PIF (Oh et al., 2004). By contrast, light-dependent chloroplast development, inhibition of hypocotyl elongation, and negative hypocotyl gravitropism are redundantly regulated by more than one PIF (Huq and Quail, 2002; Kim et al., 2003; Fujimori et al., 2004; Huq et al., 2004; Oh et al., 2004; Nozue et al., 2007; Lorrain et al., 2008, 2009; Shin et al., 2009; Stephenson et al., 2009). Consistent with the redundant negative roles of PIFs, the pif1 pif3 pif4 pif5 quadruple mutant displays a constitutive photomorphogenic phenotype (Leivar et al., 2008; Shin et al., 2009), expressing light-inducible genes even in the dark (Leivar et al., 2009; Shin et al., 2009). It has thus been argued that photomorphogenesis is the consequence of targeted removal of PIFs that regulate skotomorphogenetic development, i.e. the heterotrophic genetic pathway(s) that sustains seed and seedling viability/development using stored food reserves in the absence of light – a critical hallmark of seed plant development (Leivar and Quail, 2011).
Analysis of transgenic plants expressing truncated phytochromes suggested that N-terminal photosensory domains are themselves functional (Matsushita et al., 2003; Oka et al., 2004; Mateos et al., 2006), and mutated/truncated alleles of phytochromes that lack the ATP-binding motif in the C-terminal region retain photoregulatory activity (Krall and Reed, 2000). Further detailed analyses of the structural basis of PIF phytochrome interaction support the conclusion that N-terminal photosensory ‘input’ domains of plant phytochromes are wholly sufficient to sustain the light-dependent interaction with PIFs (Shimizu-Sato et al., 2002; Oka et al., 2008; Kikis et al., 2009). However, PIFs also bind to the C-terminal region of phytochromes, and loss-of-function point mutations in this region made it difficult to exclude a signaling role of the C-terminal region (Ni et al., 1998; Rockwell et al., 2006). It is therefore conceivable that these severely truncated hyperactive alleles mis-regulate PIF function in a light-dependent manner via a gain-of-function sequestration mechanism among other inhibitory mechanisms.
The present work was undertaken to address this sequestering mechanism. First, we addressed the question of whether PIF3 is destabilized by its interaction with the N-terminal domain of phyB (NG-GUS-NLS; NGB). These studies indicate that this interaction does not support PIF3 degradation. Second, we addressed whether DNA binding by PIFs is inhibited by phyB without PIFs being degraded. Chromatin immunoprecipitation (ChIP) analyses showed that binding of PIF1 or PIF3 to its target promoter is inhibited both by NGB and full-length phyB. In vitro binding assays further show that Pfr forms of full-length and N-terminally truncated phyB inhibit binding of PIFs to their target DNA. Thus, our results suggest that phyB inhibits PIFs in two ways: by releasing them from target promoters and by accelerating their turnover.
Previous reports have shown that truncated phytochromes lacking their histidine kinase related domain (HKRD) are still capable of inducing light responses (Krall and Reed, 2000; Matsushita et al., 2003; Mateos et al., 2006). Whether such phytochrome N-terminal domains induce light responses via the same molecular mechanism as their full-length counterparts is not known. Degradation of four negatively acting phytochrome-interacting bHLH transcription factors (PIFs) is one of the salient features of phytochrome signaling. We therefore investigated whether a previously described nuclear-targeted N-terminally truncated Arabidopsis phyB (NG-GUS-NLS; NGB) (Matsushita et al., 2003) induces degradation of PIF3. To obtain a phyB-deficient line that over-expresses both NGB and PIF3, the NGB/phyB line expressing the N-terminal 651 amino acid photosensory domain of phyB fused to GFP, GUS and a nuclear localization signal (NLS) (Matsushita et al., 2003), was crossed with the PIF3-OX/phyB and PIF3-OX/phyA lines expressing a recombinant myc-tagged PIF3 (Park et al., 2004).
A homozygous NGB/PIF3-OX/phyB line was then examined to test whether NGB induces PIF3 degradation. To minimize involvement of phyA in this process, plants were grown under continuous red light (Rc); these conditions promote phyA degradation and strongly repress PHYA transcription. The level of PIF3 was strongly reduced in Rc-grown PIF3-OX seedlings as expected for plants containing phyB, but, in contrast, PIF3 levels in Rc-grown PIF3-OX/phyB plants were unchanged when compared with their dark-grown counterparts (Figure 1a). These results show that phyB is the major phytochrome involved in targeted degradation of PIF3 under Rc. Unexpectedly, PIF3 levels in Rc-grown NGB/PIF3-OX/phyB line were the same as those found in plants grown in darkness, demonstrating that NGB does not promote PIF3 degradation. Taken together, these results indicate that the hyperactive phenotype of the NGB transgenic plants does not correlate with PIF3 turnover.
Phosphorylation of PIFs (Al-Sady et al., 2006; Lorrain et al., 2008; Shen et al., 2008) is a hallmark of phytochrome signaling. For this reason, we determined whether NGB supported light-induced phosphorylation of PIF3. To minimize the role of phyA, we generated NGB/PIF3-OX/phyA/phyB plants by crossing PIF3-OX/phyA and NGB/PIF3-OX/phyB. When etiolated PIF3-OX control seedlings were transferred to red light and examined for PIF3 bands using Western blot analysis, slower-migrating PIF3 bands were detected within 20 min (Figure 1b). These slower-migrating bands were not seen in samples treated with a protein phosphatase, indicating that they were the phosphorylated forms of PIF3. Similar phosphorylated PIF3 bands were not observed in red light-treated PIF3-OX/phyA/phyB, indicating that double mutations in PHYA and PHYB abolished the phosphorylation of PIF3. When NGB was introduced, phosphorylated PIF3 bands were detected. These results indicate that NGB restores phosphorylation of PIF3.
However, comparative analysis of the pattern of PIF3 phosphorylation revealed that full-length phytochrome mediates multiple phosphorylation of PIF3 compared with NGB (Figure 1b: compare PIF3-OX and NGB/PIF3-OX/phyA/phyB at 20 min red light). That all of these more slowly migrating species reflect phosphorylation events was ascertained by phosphatase treatment. These results indicate that NGB mediates light-dependent phosphorylation of PIF3, but multiple phosphorylation of PIF3 requires full-length phyB. As NGB does not activate the degradation of PIF3, our findings further correlate the multiple modification of PIFs with their turnover.
Previous studies have established that the N-terminal domain of phyB interacts with PIF3 in a light-dependent manner (Shimizu-Sato et al., 2002; Kikis et al., 2009). We hypothesize that such interaction may be responsible for the strong signaling activity of NGB in vivo. To test this hypothesis, co-immunoprecipitation experiments were performed using the NGB/PIF3-OX line. As shown in Figure 2(a) (left), NGB was co-precipitated with PIF3 in extracts from white light-grown tissue. This interaction was also observed in immunoprecipitates from R-treated extracts but not from FR-treated extracts (Figure 2a, right). These results are consistent with complex formation between PIF3 and the Pfr form of NGB in vivo.
Previous studies have established that formation of PIF complexes with the DELLA proteins HFR1 and PAR1 and 2 strongly inhibits PIF binding to target promoters (Feng et al., 2008; de Lucas et al., 2008; Hornitschek et al., 2009; Hao et al., 2012). We therefore assessed whether light-dependent PIF3–NGB complex formation inhibits PIF3 binding to DNA using a ChIP assay. Using promoter fragments for RGA and PIL1 genes as PIF target sequences (Oh et al., 2007; de Lucas et al., 2008; Hornitschek et al., 2009), we performed comparative PIF3 ChIP analyses on dark- and Rc-grown NGB/PIF3-OX/phyB lines. ChIP assays from dark-grown seedlings showed high enrichment of both RGA and PIL1 promoter target fragments compared to an rDNA control (Figure 2b). RGA and PIL1 promoter fragments were less abundant in immunoprecipitates from Rc-grown seedlings, supporting the conclusion that PIF3 binding to both promoters is inhibited by Rc. Western blot analyses indicate that this difference was not due to higher levels of precipitated PIF3 protein in ChIP assays from dark-grown seedlings (Figure 2c). We next examined the expression of PIL1, a gene known to be up-regulated by PIFs. As expected, expression of PIL1 was strongly repressed by red light – a response that was srongly dependent on functional phyB (Figure 2d). Red light also strongly decreased the expression of PIL1 in both the NGB/phyB and NGB/PIF3-OX/phyB lines (Figure 2d). Taken together, our results indicate that, unlike wild-type phyB, NGB inhibits PIF3 in the light without triggering its degradation. NGB may inhibit PIF3 DNA binding directly by binding to PIF3 or by modifying PIF3. Alternatively, as phytochrome is known to inhibit gibberellic acid (GA) biosynthesis in the seedling stage (Kamiya and Garcia-Martinez, 1999), NGB may inhibit PIF3 indirectly by increasing the DELLA level.
These observations led us to test whether full-length phyB also inhibits PIF3 binding to target promoters in a light-dependent manner. To do so, we performed ChIP analysis using dark- and Rc-grown PIF3-OX plant lines. Both PIF3 target promoters were greatly enriched in ChIPs from dark-grown seedlings, but not from Rc-grown seedlings (Figure 3a). To test whether PIF3 degradation in Rc was solely responsible for the result, we also performed ChIP experiments to compare promoter occupancy in dark continuous (Dc)- and Rc-grown PIF3-OX seedlings treated with the 26S proteasome inhibitor MG132. For Dc-grown seedlings, PIF3 was strongly associated with target promoters, and this association further increased following MG132 treatment (Figure 3b), reflecting the increased PIF3 level in MG132-treated plants. In contrast, under Rc, despite the enhanced PIF3 accumulation by MG132 treatment, we did not detect increased association of PIF3 with its target promoters. These data imply that PIF3 does not bind to its target promoters under Rc conditions that favor the phyB–PIF3 association.
An additional ChIP experiment was performed to test whether PIF1 binding to the PIL1 promoter was similarly repressed by Rc in a PIF1-OX line. Like PIF3, MG132 treatment increased the PIF1 protein level, but did not enhance its binding to the target promoter (Figure 3c). The results of pulse irradiation of Rc-grown seedlings with far-red light followed by a short incubation in darkness were also examined. These experiments showed that the amount of target promoter recognized by both PIFs was significantly increased (Figure 3d). Taken together, these results indicate that PIF binding to target promoters is inhibited by red light even in the absence of protein turnover.
To ascertain whether R-dependent inhibition of PIF3 binding to its target promoter is phyB-dependent, we performed ChIP analysis using the PIF3-OX/phyB mutant line. Unlike NGB/PIF3-OX/phyB plants, enrichment of the PIF3 target promoter was not reduced by R treatment in PIF3-OX/phyB mutant plants (Figure 4a). Based on these results, phyB is responsible for the R-dependent inhibition of PIF3 binding to its target promoter.
Finally, to establish whether phytochrome interaction with PIFs is required to inhibit DNA binding, we generated transgenic PIF3ΔN-OX lines expressing myc-tagged PIF3 lacking the N-terminal 300 amino acids. This PIF3 construct lacked both phy-interacting motifs (APA and APB) and its transcription activation domain, but retained the C-terminal 224 amino acids harboring the DNA-binding bHLH domain (Khanna et al., 2004; Al-Sady et al., 2006; Shen et al., 2008). Due to deletion of transcription activation domain, transgenic lines expressing PIF3ΔN displayed dominant-negative constitutive photomorphogenic phenotypes that included short hypocotyls in the dark, decreased greening of etiolated seedlings when transferred to light, and disrupted hypocotyl negative gravitropism in the dark (Figure S1). Similar constitutive photomorphogenic phenotypes were reported for an N-terminal deletion of PIF1 (Shen et al., 2008). PIF3ΔN was also not degraded by light (Figure S1), consistent with the essential role of phy-interacting motifs for light-dependent degradation. ChIP analysis showed that PIF3ΔN binds well to its target promoter, and this binding is not inhibited by red light (Figure 4b). This confirms that the Pfr form of phytochrome interacts with PIF3 to inhibit its binding to target promoters.
To further investigate whether phyB directly inhibits the DNA binding of PIFs, we performed in vitro inhibition assays using recombinant phyB, PIFs and a biotinylated PIL1 target promoter fragment. For the assay, we first incubated biotinylated target DNA with PIFs, and PIF-bound target DNAs were separated by streptavidin resin to remove unbound PIFs. The PIF–target DNA complex was then incubated with either the Pr or Pfr forms of recombinant phyB. After thorough washing, DNA-bound PIFs were eluted and quantified immunochemically. These experiments showed that smaller amounts of PIF1 and PIF3 proteins remained bound to target DNAs when incubated with the Pfr form of phyB than with the Pr form (Figure 5). We also performed the same experiment using the N-terminal domain of phyB (amino acids 1–650), which was previously shown to be functional even in the absence of the dimerization domain (Matsushita et al., 2003). Similar to our findings with full-length phyB, the Pfr form of the N-terminal domain of phyB also dissociated PIF1 from the target promoter fragment (Figure 5). Taken together, our results indicate that the Pfr forms of both full-length and N-terminal domain phyB directly inhibit binding of PIFs to their target promoters.
In this study, we show that the activity of the N-terminal domain of phyB (NG-GUS-NLS; NGB) (Matsushita et al., 2003) is not due to targeted degradation of PIF3 in response to red light. Instead, N-terminal phyB inhibits binding of PIF3 to its target promoters in a red light-dependent manner in vivo. Moreover, we also show that full-length phyB also inhibits the binding of PIFs to their target promoters under red light in vivo. Indeed, Pfr forms of both full-length and N-terminal phyB are capable of inhibiting the DNA binding of PIFs in vitro. Based on these observations, we propose that phyB inhibits negatively acting PIFs by two different modes of action: by releasing them from their target promoters and by mediating their degradation.
Our results also indicate that the NGB, although functionally similar to full-length phyB (Matsushita et al., 2003), lacks the full regulatory activity of full-length phyB. Indeed, although both N-terminal and full-length phyB inhibit DNA binding of PIFs both in vivo and in vitro, NGB fails to target PIFs for degradation. In addition, when etiolated seedlings were transferred to red light, NGB yielded only one slower-migrating phosphorylated PIF3 band. By contrast, the full-length phyB yielded multiple phosphorylated bands under the same conditions. This suggests that the light-dependent interaction between NGB and PIFs is sufficient to trigger partial PIF phosphorylation. We therefore conclude that the C-terminal domain, although dispensable for the induction of light responses (Krall and Reed, 2000; Matsushita et al., 2003), is required for multiple light-dependent phosphorylation of PIF3. As NGB does not mediate PIF3 degradation under red light, we also conclude that the C-terminal domain of phyB is necessary for targeted degradation of PIFs. We hypothesize that the C-terminal domain of phyB promotes PIF turnover by mediating multiple phosphotransfers to PIFs and/or by recruiting ubiquitin E3 ligases. Although phosphotransfer may be accomplished by phytochrome itself (Yeh and Lagarias, 1998), the C-terminal domain of phytochrome may be responsible for recruiting a protein kinase and/or a ubiquitin E3 ligase for these functions. A previous report showed that casein kinase II (CKII) phosphorylated PIF1 in vitro, and multiple mutations in CKII target residues (Ser/Thr residues including three serine residues at its C-terminus: S464, S465 and S466) partially inhibited light-induced degradation of PIF1 (Bu et al., 2011). Although such mutations did not completely abolish light-induced phosphorylation of PIF1, it would be interesting to determine whether CKII plays a role in multiple phosphorylation of PIFs in vivo.
Consistent with our data, previous EMSA-based studies showed that Pfr fails to form ternary complexes with the G-box element and PIF1 or PIF4 (Huq and Quail, 2002; Huq et al., 2004). By contrast, these reports also showed that the DNA-binding activities of PIF1 and PIF4 were not disrupted by the presence of Pfr. Our results also contrast with the reported formation of a ternary complex between the Pfr form of phyB, PIF3 and a G-box-containing oligonucleotide in an EMSA assay (Martinez-Garcia et al., 2000; Huq and Quail, 2002; Huq et al., 2004). Although it is presently unclear what is responsible for the difference between our results and previously reported results, we suspect that these differences may reflect the distinct protocols used. In this regard, in previous studies, phyB and PIFs were co-incubated in crude TnT reaction mixtures (Promega, http://www.promega.com), but we removed excess PIFs not bound to DNA prior to addition of phyB. When all three components (PIF1, phyB and the target DNA fragment) were incubated together without removing excess PIF1 prior to phyB addition, similar levels of PIF1 co-precipitated with the DNA fragment irrespective of Pr or Pfr. This suggests that free PIF1 should be removed to observe the effect of Pr and Pfr.
DNA binding by transcription factors is extensively regulated. Multiple modes of regulation have been reported for various transcription factors in various organisms. For example, DNA binding of a transcription factor may be regulated by controlling its nuclear localization. Among plant transcription factors, REPRESSION OF SHOOT GROWTH (RSG), a basic leucine zipper transcription factor regulating GA biosynthesis, is actively exported from the nucleus and is held by binding protein 14-3-3 in the cytosol in an inactive state (Igarashi et al., 2001). Similarly, BRASSINAZOLE RESISTANT1 (BZR1) and BRI1 EMS SUPPRESSOR1 (BES1), two positive brassinosteroid signaling transcription factors, are also actively exported from the nucleus upon brassinosteroid treatment (Bai et al., 2007; Gampala et al., 2007; Ryu et al., 2007, 2008). DNA binding of a transcription factor may also be regulated by controlling its ability to bind DNA. This may be accomplished by physical association with other proteins, e.g. disruption of DNA binding of the class III homeodomain/leucine zipper (HD-ZIPIII) by interaction with LITTLE ZIPPER (ZPR) protein (Wenkel et al., 2007), or by covalent modification such as phosphorylation, which has been shown to inhibit DNA binding for many transcription factors, i.e. ELONGATED HYPOCOTYL5 (HY5), BZR1, BZR2, AUXIN RESPONSE FACTOR2 and HEAT-SHOCK TRANSCRIPTION FACTOR1 (HSF1) (Reindl et al., 1997; Hardtke et al., 2000; Vert and Chory, 2006; Gampala et al., 2007; Vert et al., 2008).
Indeed, regulation of DNA binding by PIFs appears to be regulated by multiple factors. For example, it is well established that the DELLA repressors of GA signaling bind to bHLH motifs of PIF3 and PIF4 and prevent PIF binding to target promoters (Feng et al., 2008; de Lucas et al., 2008; Cheminant et al., 2011). This inhibition of PIF DNA binding by DELLA proteins has been suggested to play a key role in coordinating GA signaling and light signaling. HFR1, a positive far-red light signaling atypical bHLH protein, has been shown to heterodimerize with PIFs to inhibit their binding to target promoters (Hornitschek et al., 2009). Indeed, the positive role of HFR1 in far red-enriched shade conditions has been attributed to its ability to disrupt DNA binding of negatively acting PIFs. Similar to HFR1, other atypical HLH proteins, PAR1 and PAR2, heterodimerize with PIF4 to inhibit its binding to target promoters (Hao et al., 2012). Here, we show that light-activated phyB, like the DELLA proteins HFR1, PAR1, and PAR2, may also function as a PIF-binding protein to inhibit their DNA-binding activity. Our results support a regulatory model for phytochrome signaling in which the light-dependent interaction between PIFs and phyB inhibits binding of these bHLH transcription factors to their target promoters, effecting regulation of gene expression. It is interesting that the modes of interaction of each of the three classes appear distinct, i.e. phytochromes bind to N-terminal motifs of PIFs, DELLAs bind to the bHLH motif of PIFs (Feng et al., 2008; de Lucas et al., 2008), and HFR1, PAR1 and PAR2 heterodimerize with PIFs (Hornitschek et al., 2009; Hao et al., 2012). It is therefore conceivable that these factors function synergistically or additively in vivo.
Much work remains to elucidate the molecular mechanism underlying phytochrome inhibition of DNA binding of PIFs. We have shown that the inhibition requires a light-dependent interaction between phytochrome and PIFs and that it may occur in the absence of ATP. This indicates that the interaction itself is sufficient to disrupt DNA binding without phosphorylation. This implies that the interaction between PIFs and phyB-Pfr (the Pfr form of phyB) directly competes with DNA binding. We speculate that this association induces an allosteric change in PIFs to destabilize DNA binding. However, our data do not exclude the possibility that phosphorylation of PIFs and/or interaction with other proteins such as DELLA may further destabilize the interaction between PIFs and their target promoters in vivo. A recent report indicated that DNA binding of PIF7, which is phosphorylated but not destabilized by a high red:far red ratio, is also inhibited by high R:FR in vivo (Li et al., 2012). This suggests that the inhibition of PIF DNA binding by phytochrome may be a common mechanism for other PIFs. Further studies are required to determine the detailed molecular mechanism of how phytochrome disrupts DNA binding of PIFs.
Plant materials and growth conditions
Arabidopsis thaliana plants were maintained in a controlled growth facility with a 16 h light/8 h dark cycle at 22–24°C for general growth and seed harvesting. The NG-GUS-NLS (NGB) line NG-GUS-NLS 6-4 expressing the N-terminal domain of phyB fused to GFP, GUS, and NLS in the phyB-5 null background in the Ler ecotype has been described previously (Matsushita et al., 2003). PIF3-OX and PIF1-OX correspond to previously described PIF3-OX3 and PIL5-OX3 (Park et al., 2004; Oh et al., 2006). Two independent NGB/PIF3-OX/phyB lines were established by crossing PIF3-OX/phyB-9 with NGB/phyB-5. Four independent NGB/PIF3-OX/phyA/phyB lines were isolated by crossing PIF3-OX/phyA-211 with NGB/PIF3-OX/phyB-9. All showed similar partial phosphorylation and no degradation of PIF3. PIF3ΔN lines that express myc-tagged PIF3 lacking the N-terminal 300 amino acids under the control of the CaMV 35S promoter were generated by cloning N-terminally truncated PIF3 into the pHTM vector using a specific primer set (Table S1), a derivative of pBI121 vector (Clontech, http://www.clontech.com).
Chromatin immunoprecipitation (ChIP)
For the ChIP analysis, seedlings were grown under dark or red light (10 μmol m−2 sec−1) for 4 days. For some experiments, 4-day-old seedlings were treated with 80 μm MG132 before sampling for 8 h. Samples (1 g) from 4-day-old dark- or red light-grown seedlings were treated with 10 ml of 1% formaldehyde under vacuum infiltration conditions. ChIP assays were performed as described previously (Shin et al., 2007) with minor modification of the three wash buffers: low salt wash buffer (150 mm NaCl, 0.1% SDS, 1% Triton X-100, 2 mm EDTA, 20 mm Tris/HCl, pH 8), high-salt wash buffer (500 mm NaCl, 0.1% SDS, 1% Triton X-100, 2 mm EDTA, 20 mm Tris/HCl, pH 8), LiCl wash buffer (0.25 M LiCl, 1% v/v Nonidet P-40, 1% w/v sodium deoxycholate, 1 mm EDTA, 10 mm Tris/HCl, pH 8). The amount of each precipitated DNA fragment was determined by real-time PCR using RGA, PIL1 and rDNA primers (Table S1).
Phosphorylation and degradation of PIF3
For phosphatase treatments, 5-day-old dark grown seedlings expressing His- or myc-tagged PIF3 were ground in liquid nitrogen and homogenized in a denaturing buffer [100 mm NaH2PO4, pH 8.0, 10 mm NaCl, 8 m urea, 2 mm phenylmethanesulfonyl fluoride, 80 μm MG132, 1× complete protease inhibitor cocktail (Roche, http://www.roche.com)]. After centrifugation at 20676g for 10 min at 4°C, PIF3 was purified from supernatants using Ni-NTA beads (Qiagen, http://www.qiagen.com). Pellets were washed twice with PBS buffer (8 g l−1 NaCl, 0.2 g l−1 KCl, 1.44 g l−1 Na2HPO4, 0.24 g l−1 KH2PO4, pH 7.4) and once with CIP buffer (NEBuffer 3: 100 mm NaCl, 50 mm Tris/HCl, 10 mm MgCl2, 1 mm dithiothreitol, pH 7.9). The resuspended pellets were treated for 15 min at 37°C with no enzyme, 100 units calf intestine alkaline phosphatase (CIP) (NEB, http://www.neb.com), or comparable amounts of boiled CIP. Following these incubations, reaction mixtures were boiled in 2× SDS sample buffer (45 mm Tris/HCl pH 6.8, 10% v/v glycerol, 1% w/v SDS, 0.05% w/v bromophenol blue, 50 mm dithiothreitol) and subjected to Western blot analysis with anti-myc antibody. For assessment of PIF protein stability, 80 dark- and light-grown seedlings were frozen using liquid nitrogen and then homogenized in denaturing buffer (100 mm NaH2PO4, 10 mm Tris/HCl, 8 M urea, pH 8.0). The protein extraction and Western blotting were performed as described previously (Park et al., 2004).
For the NGB line, the pelleted nuclei fraction was resuspended in extraction buffer (without sucrose and with 0.5% Triton X-100) and immunoprecipitated with myc antibody. For co-immunoprecipitation of phyB and PIF3, total proteins were extracted with an immunoprecipitation buffer (100 mm NaH2PO4, pH 7.8, 100 mm NaCl, 0.1% v/v NP-40, 2 mm phenylmethanesulfonyl fluoride, 100 μm MG132, 1× complete protease inhibitor cocktail). After removing debris by centrifugation at 20676g for 10 min at 4°C, the supernatant was pre-cleared using Protein A beads (Thermo Scientific, http://www.thermoscientific.com) at 4°C for 1 h to remove non-specific binding proteins, and immunoprecipitated using anti-myc antibody (Santa Cruz Biotechnology, http://www.scbt.com). To test the reversible binding of NGB to PIF3, pre-cleared extracts from dark-grown seedlings were exposed to red light (10 μmol m−2 sec−1) or far-red light (3 μmol m−2 sec−1) for 20 min prior to co-immunoprecipitation.
DNA pull-down assay
Biotin-labeled double strand oligonucleotides corresponding to the PIF-binding site of the PIL1 promoter fragment (PIL1p, 300 ng) were first bound to 20 μl of streptavidin agarose resin (Thermo Scientific). The resin was then equilibrated by washing three times for one minute with 1 ml TKMG buffer (50 mm Tris/HCl, pH 7.5, 150 mm KCl, 1 mm EDTA, 5 mm MgCl2, 0.5% v/v NP-40, 10% v/v glycerol). His-tagged PIF1 (200 ng) or PIF3 (100 ng) proteins were incubated with the PIL1p resin in the presence of 1 μg poly dI-dC (deoxyinosinic-deoxycytidylic; Sigma, http://www.sigmaaldrich.com) and 10 μg BSA (NEB) at 4°C for 2 h. Unbound PIF1 or PIF3 proteins were removed by washing three times with 1 ml TKMG buffer. The PIF-bound PIL1p resin was incubated with 1 μg purified recombinant phyB at 4°C for 2 h in the dark. PhyB was irradiated with far-red light (3 μmol m−2 sec−1) for 10 min (Pr) or red light (10 μmol m−2 sec−1) for 10 min (Pfr) prior to incubation. After washing three times with 1 ml TKMG buffer under green light, bound PIFs were eluted by boiling in 2× SDS buffer. The eluted PIF1 or PIF3 proteins were resolved by SDS–PAGE, transblotted to nitrocellulose membrane, and detected by PIF1 or PIF3 antibody after SDS–PAGE. PIF3 antibody is a polyclonal antibody against PIF3–GST and PIF1 antibody is a polyclonal antibody against a PIF1 peptide (KTNVDDRKRKEREATT).
Recombinant full-length or N-terminal phytochrome B (amino acids 1–650) preparations were obtained by cloning into pBAD vector, co-transformation with pPL-PCB into Escherichia coli strain LMG 194, and purification as described previously (Gambetta and Lagarias, 2001). For purification, cells were collected by centrifugation (5590g, 4°C, 20 min), resuspended, and sonicated in a lysis buffer (50 mm NaH2PO4, pH 8.0, 300 mm NaCl, 10% glycerol, 20 mm imidazole, 0.05% Tween-20, 1 mm 2-mercaptoethanol, 1 mm phenylmethanesulfonyl fluoride, 1× complete protease inhibitor cocktail). Proteins were further purified using Ni-NTA agarose resin as described previously (Gambetta and Lagarias, 2001). Purified recombinant full-length phyB bound PIF3 light-dependently (Figure S2).
This work was supported in part by two grants from the National Research Foundation of Korea (2012R1A2A1A01003133 and ABC-0031339), and the Rural Development Administration (SSAC-PJ008120) to G.C, a grant from the US National Institutes of Health (GM068552) to J.C.L, and a Grant-in-Aid for Scientific Research on Innovative Areas from the Ministry of Education, Culture, Sports, Science and Technology, Japan (number 22120002) to A.N.