Translator Disclaimer
1 August 2011 Phytochrome Signaling Mechanisms
Author Affiliations +
Abstract

Phytochromes are red (R)/far-red (FR) light photoreceptors that play fundamental roles in photoperception of the light environment and the subsequent adaptation of plant growth and development. There are five distinct phytochromes in Arabidopsis thaliana, designated phytochrome A (phyA) to phyE. phyA is light-labile and is the primary photoreceptor responsible for mediating photomorphogenic responses in FR light, whereas phyB-phyE are light stable, and phyB is the predominant phytochrome regulating de-etiolation responses in R light. Phytochromes are synthesized in the cytosol in their inactive Pr form. Upon light irradiation, phytochromes are converted to the biologically active Pfr form, and translocate into the nucleus. phyB can enter the nucleus by itself in response to R light, whereas phyA nuclear import depends on two small plant-specific proteins FAR-RED ELONGATED HYPOCOTYL 1 (FHY1) and FHY1-LIKE (FHL). Phytochromes may function as light-regulated serine/threonine kinases, and can phosphorylate several substrates, including themselves in vitro. Phytochromes are phosphoproteins, and can be dephosphorylated by a few protein phosphatases. Photoactivated phytochromes rapidly change the expression of light-responsive genes by repressing the activity of CONSTITUTIVE PHOTOMORPHOGENIC 1 (COP1), an E3 ubiquitin ligase targeting several photomorphogenesis-promoting transcription factors for degradation, and by inducing rapid phosphorylation and degradation of Phytochrome-Interacting Factors (PIFs), a group of bHLH transcription factors repressing photomorphogenesis. Phytochromes are targeted by COP1 for degradation via the ubiquitin/26S proteasome pathway.

INTRODUCTION

As sessile organisms, plants have acquired a high degree of developmental plasticity to optimize their growth and reproduction in response to their ambient environment, such as light, temperature, humidity, and salinity. Plants utilize a wide range of sensory systems to perceive and transduce specific incoming environmental signals. Light is one of the key environmental signals that influences plant growth and development. In addition to being the primary energy source for plants, light also controls multiple developmental processes in the plant life cycle, including seed germination, seedling de-etiolation, leaf expansion, stem elongation, phototropism, stomata and chloroplast movement, shade avoidance, circadian rhythms, and flowering time (Deng and Quail, 1999; Wang and Deng, 2003; Jiao et al., 2007).

Plants can monitor almost all facets of light, such as direction, duration, quantity, and wavelength by using at least four major classes of photoreceptors: phytochromes (phys) primarily responsible for absorbing the red (R) and far-red (FR) wavelengths (600–750 nm), and three types of photoreceptors perceiving the blue (B)/ultraviolet-A (UV-A) region of the spectrum (320–500 nm): cryptochromes (crys), phototropins (phots), and three newly recognized LOV/F-box/Kelch-repeat proteins ZEITLUPE (ZTL), FLAVIN-BINDING KELCH REPEAT F-BOX (FKF), and LOV KELCH REPEAT PROTEIN 2 (LKP2). In addition, UV RESISTANCE LOCUS 8 (UVR8) was recently shown to be a UV-B (282–320 nm) photoreceptor (Rizzini et al., 2011). These photoreceptors perceive, interpret, and transduce light signals, via distinct intracellular signaling pathways, to modulate photoresponsive nuclear gene expression, and ultimately leading to adaptive changes at the cell and whole organism levels.

The past two decades have seen dramatic progress in molecular characterization and understanding of the photobiology and photochemistry of the phytochrome photoreceptors in higher plants. This chapter aims to highlight some of the most recent progress in elucidating the molecular, cellular and biochemical mechanisms of phytochrome signaling in Arabidopsis. Interested readers are encouraged to read the accompanying reviews on other related subjects, such as photomorphogenesis (Nemhauser and Chory, 2002), cryptochromes (Yu et al., 2010), phototropins (Pedmale et al., 2010), and the circadian clock (McClung et al., 2002).

PLANT PHYTOCHROMES

The Discovery and Action Modes of Phytochromes

The term phytochrome, meaning “plant color”, was originally coined to describe the proteinous pigment that controls photoperiod detection and floral induction of certain short-day plants (such as cocklebur and soybean) (Garner and Allard, 1920), and the reversible seed germination of lettuce (c.v. Grand Rapids) by R and FR light (Borthwick et al., 1952). R light promotes seed germination, whereas subsequent FR light treatment abolishes R light induction of seed germination. The germination response of lettuce seeds repeatedly treated with R/FR cycles is determined by the last light treatment. Thus R/FR photoreversibility is a characteristic feature of this response. In addition, the law of reciprocity applies to this response, i.e. the response is dependent on the total amount of photons received irrespective of the duration of light treatment.

Over the years, three action modes for phytochromes have been defined, i.e. low-fluence responses (LFRs), very-low-fluence responses (VLFRs) and high-irradiance responses (HIRs) (Table 1). The above-mentioned F/FR reversible response is characteristic of LFRs. LFRs also induce other transient responses, such as changes in ion flux, leaf movement, chloroplast rotation, and changes in gene expression (Haupt and Hader, 1994; Roux, 1994; Vince-Prue, 1994). VLFRs are activated by extremely low light intensities of different wavelengths (FR, R and B); examples include light-induced expression of the light-harvesting chlorophyll a/b-binding protein (LHCB) gene and light induction of seed germination. HIRs depend on prolonged exposure to relatively high light intensities, and are primarily responsible for the control of seedling de-etiolation (e.g. inhibition of hypocotyl elongation and promotion of cotyledon expansion) under all light qualities (Mustilli and Bowler, 1997; Casal et al., 1998; Neff et al., 2000; Table 1).

Chromophores and Two Reversible Forms of Phytochromes

Photoreversibility occurs because phytochromes exist as two distinct but photoreversible forms in vivo: the R light-absorbing form (Pr) and the FR light-absorbing form (Pfr). The Pr form absorbs maximally at 660 nm, whereas the Pfr form absorbs maximally at 730 nm (Quail, 1997a; Figure 1). The Pfr forms of phytochromes are generally considered to be the biologically active forms. It should be noted that in addition to their maximal absorptions of R and FR wavelengths, phytochromes also weakly absorb B light (Furuya and Song, 1994; Figure 1).

Phytochromes are soluble proteins and exist as homodimers. The molecular mass of the apoprotein monomer is approximately 125 kDa. Phytochrome apoproteins are synthesized in the cytosol, where they assemble autocatalytically with a linear tetrapyrrole chromophore, phytochromobillin (PΦB). The synthesis of PΦB is accomplished by a series of enzymatic reactions in the plastid that begins with 5-aminolevulinic acid (Figure 2A). The early steps in the PΦB pathway are shared with chlorophyll and heme biosynthesis. The committed step is the oxidative cleavage of heme by a ferredoxin-dependent heme oxygenase (HO) to form biliverdin IX (BV). BV is subsequently reduced to 3Z-PΦB by the enzyme PΦB synthase. Both 3Z-PΦB and its isomerized form 3E-PΦB can serve as functional precursors of the phytochrome chromophore. PΦB is then exported to the cytosol, where it binds to the newly synthesized apo-PHYs to form holo-PHYs (Terry, 1997; Figure 2A). The chromophore is attached via a thioether linkage to an invariant cysteine in a well-conserved domain among all phytochromes (see below).

The intrinsic photochemical activity of the chromophore prosthetic group allows phytochromes to convert between the two forms. Phytochromes are synthesized in the Pr form in dark-grown seedlings. It has been widely accepted that absorption of R light triggers a “Z” to “E” isomerization in the C15–C16 double bond between the C and D rings of the linear tetrapyrrole, resulting in the FR-absorbing Pfr form (Andel et al., 1996; Figure 2B). However, a recent NMR analysis showed that the A pyrrole ring around C4–C5 double bond rotates during photoconversion (Ulijasz et al., 2010). This discrepancy should be resolved in future studies. In addition, the Pr-to-Pfr transition is associated with rearrangement of the protein backbone (Figure 2B). The active Pfr form can be converted back to the inactive Pr form, either by a slow non-photoinduced reaction (dark reversion) or much faster upon absorption of FR light (Mancinelli, 1994; Quail, 1997a; Fankhauser, 2001; Figure 2B). This property allows phytochrome to function as a R/FR-dependent developmental switch.

Figure 1.

Absorption spectra of phytochromes.

Absorption spectra of the two forms (Pr and Pfr) of phytochromes. The Pr form absorbs maximally at 660 nm, while the Pfr form absorbs maximally at 730 nm. The visible light range of the human eye is approximately 380–700 nm. The light spectrum was adapted from Kami et al. (2010). Reprinted with permission from Elsevier.

Table 1.

Diagnostic Features of Different Phytochrome Action Modes

Figure 2.

Arabidopsis phytochrome chromophore.

(A) The biosynthesis pathway of Arabidopsis phytochrome chromophore. Image adapted from Kohchi et al. (2001).

(B) Red (R) light triggers a “Z” to “E” isomerization in the C15–C16 double bond between the C and D rings of the linear tetrapyrrole (upper panel), which is accompanied by rearrangement of the apoprotein backbone (lower panel; adapted from Bae and Choi, 2008). This results in the photoconversion of phytochromes from the Pr form to the Pfr form. Please note that the chromophore ring A rather than D is rotated during photoconversion according to a recent NMR analysis (Ulijasz et al., 2010). The discrepancy needs to be resolved in future studies. Far-red (FR) light converts the Pfr form back to the Pr form.

Upper panel image reprinted from Bae and Choi (2008) with permission, from the Annual Review of Plant Biology, Volume 59 © 2008 by Annual Reviews ( www.annualreviews.org).

Figure 3.

The phylogenetic tree of the five phytochrome species from Arabidopsis thaliana.

PHYB and PHYD share ∼80% amino acid sequence identity, and constitute a branch of the gene family. PHYE itself, PHYA and PHYC form two other branches of the evolutionary family tree.

Image adapted from Clack et al. (1994). Reprinted with permission from Springer.

It is generally assumed that all phytochromes have the same chromophore. Arabidopsis mutants defective in the PΦB-synthetic pathway have been isolated. These mutants (hy1 and hy2) have dramatically reduced levels of PΦB and consequently of functional phytochromes, and thus exhibit severely impaired photomorphogenesis (Parks and Quail, 1991). The Arabidopsis HY1 locus encodes a heme oxygenase (AtHO1) responsible for much of PΦB synthesis in Arabidopsis (Davis et al., 1999a; Muramoto et al., 1999). Three additional HO genes were found in the Arabidopsis genome, designated AtHO2 to AtHO4 (Davis et al., 2001). The Arabidopsis HY2 locus, likely a unique gene in the Arabidopsis genome, encodes the phytochromobilin synthase (Kohchi et al., 2001).

It should be pointed out that in addition to PΦB, phycocyanobilin (PCB), the chromophore of the light-harvesting pigment phycocyanin, can also bind phytochrome resulting in Pr and Pfr spectra that are slightly blue shifted compared with the PΦB adducts (Lagarias and Rapoport, 1980). This finding allowed the reconstitution of photoreversible phytochromes by expressing recombinant phytochrome proteins in yeast and assembling them in vitro.

The Phytochrome Gene Family

In Arabidopsis thaliana, there are five phytochromes, designated phytochrome A (phyA) to phyE. They are encoded by five distinct members of the phytochrome gene family and are classified into two groups according to their stability in light (Sharrock and Quail, 1989). phyA is a type I (light labile) phytochrome, and phyB to phyE are all type II (light stable) phytochromes. phyA is most abundant in dark-grown seedlings, whereas its level drops rapidly upon exposure to R or white (W) light. In light-grown plants, phyB is the most abundant phytochrome, whereas phyC-phyE are less abundant (Clack et al., 1994; Hirschfeld et al., 1998; Sharrock and Clack, 2002).

Sequence analysis suggests that these phytochromes can be clustered into three subfamilies: phyA/phyC, phyB/phyD, and phyE (Figure 3). Analysis of reconstituted recombinant phyA, phyB, phyC and phyE proteins revealed that they have similar but not identical spectral properties (Kunkel et al., 1996; Remberg et al., 1998; Eichenberg et al., 2000). Orthologs of Arabidopsis PHY genes are present in most, if not all, higher plants (Clack et al., 1994; Sharrock and Quail, 1989; Mathews and Sharrock, 1997).

Figure 4.

The domain structure of Arabidopsis phyA and phyB molecules.

H, hinge; NTE, N-terminal extension; PAS, Per (period circadian protein), Arnt (Ah receptor nuclear translocator protein), and Sim (single-minded protein); GAF, cGMP-stimulated phosphodiesterase, Anabaena adenylate cyclases and Escherichia coli FhlA; PHY, phytochrome; PRD, PAS-related domain; HKRD, histidine kinase—related domain. The chromophore is attached to a conserved cysteine residue in the GAF domain. The numbers indicate the positions of each domain.

Image adapted from Bae and Choi (2008). Reprinted with permission, from the Annual Review of Plant Biology, Volume 59 © 2008 by Annual Reviews ( www.annualreviews.org).

General Structures of Phytochromes

The phytochrome molecule consists of an N-terminal domain (∼ 70 kDa) and a C-terminal domain (∼ 55 kDa), connected by a flexible hinge region (Figure 4). The N-terminal domain can be further divided into four consecutive subdomains: N-terminal extension (NTE), Per-Arnt-Sim (PAS), GAF, and PHY, while the C-terminal domain can also be divided into two subdomains: the PAS-related domain (PRD) containing two PAS repeats, and the histidine kinase-related domain (HKRD) (Figure 4).

Among the N-terminal subdomains, the NTE domain is uniquely present in plant phytochromes, whereas the PAS, GAF and PHY domains are also found in phytochrome-like proteins of various organisms (see below). Among the C-terminal subdomains, the PRD domain is unique to plant phytochromes, whereas the HKRD domain is also found in phytochrome-like proteins (Rockwell et al., 2006; Bae and Choi, 2008; Nagatani, 2010). The chromophore is attached to a conserved cysteine residue in the GAF domain of plant phytochromes (Figure 4). The PAS domains can be used either as platforms for protein-protein interactions, or as response modules to small ligands or changes in light conditions, oxygen levels, and redox potentials (Quail, 1997a; Neff et al., 2000). The HKRD domain lacks a critical histidine residue, and thus may be an evolutionary remnant rather than an active histidine kinase (Boylan and Quail, 1996). The putative dimerization motifs of phytochromes are also localized in the C-terminal half of the phytochrome molecules (Quail, 1997a).

A number of point mutations in the C-terminal domains of both phyA and phyB do not affect photoreversibility but eliminate the biological activity (Quail et al., 1995; Quail, 1997a), suggesting that the C-terminal domain is essential for proper downstream signaling. Consistent with this idea, a domain-swapping and deletion analysis suggested that the N terminus of phytochrome is essential for its specific photosensory properties, while the C termini of phyA and phyB are interchangeable and function as the output domains (Wagner et al., 1996). However, this notion was later challenged by the finding that the N-terminal domain of phyB, when dimerized and localized in the nucleus, confers much higher photosensitivity than the full-length phyB (Matsushita et al., 2003). These results suggest that the N-terminal domain of phyB transduces the light signal to downstream targets, whereas the C-terminal domain attenuates the activity of phyB (Matsushita et al., 2003). Similarly, the N-terminal domain of phyA also showed a partial physiological activity when dimerized and localized in the nucleus (Mateos et al., 2006). A further study demonstrated that dimers of the N-terminal 450-aa fragment of phyB (lacking the PHY domain) can still transduce the light signal upon nuclear localization (Oka et al., 2004). Therefore, these reports suggest that the N-terminal 450-aa fragment (encompassing the NTE, PAS and GAF domains) constitutes the core signaling domain of phytochrome.

PHYTOCHROME FUNCTIONS

In most instances, the roles of individual phytochromes are studied in the context of specific responses and/or developmental stages. Loss-of-function studies of monogenic phytochrome mutant and higher order mutants of various combinations combined with gain-of-function analyses are revealing the roles of individual phytochromes in regulating different aspects of plant development. It is clear now that individual phytochromes play both unique and overlapping roles throughout the life cycle of plants, regulating a range of developmental processes from seed germination to the timing of reproductive development (Table 2). Phytochrome functions in Arabidopsis development were recently reviewed by Franklin and Quail (2010), and this chapter will mainly discuss the roles of phytochromes in seed germination, seedling de-etiolation, and shade avoidance.

Table 2.

Different Roles of Phytochrome Family Members in Seedling and Early Vegetative Development

Seed Germination

As mentioned above, the involvement of a R/FR-reversible photoreceptor in mediating seed germination was first demonstrated by Harry Borthwick and colleagues in 1952 by studying the reversible germination of Grand Rapids lettuce seeds by R and FR treatments (Borthwick et al., 1952). The involvement of individual phytochromes in mediating Arabidopsis seed germination has been documented in many mutant studies. At least three phytochromes, i.e. phyA, phyB and phyE are involved in the control of Arabidopsis seed germination. phyA is responsible for the irreversible VLFR responses triggered by a wide variety of irradiations (ultraviolet, visible and FR light), while phyB controls the R/FR photo reversible LFRs (Reed et al., 1994; Botto et al., 1996; Shinomura et al., 1996). Seed germination can be promoted by both VLFRs and LFRs (Table 2). In addition, phyA promotes germination in continuous FR light in the HIR mode (Johnson et al., 1994; Reed et al., 1994; Hennig et al., 2002). However, phyE was also found to play a role in controlling seed germination in continuous FR light. This could be either because phyE is directly involved in the photoperception of FR light for this response, or because phyA requires phyE to mediate seed germination (Hennig et al., 2002). It is interesting that ambient temperature modulates the light-regulation of Arabidopsis seed germination, and different phytochromes display altered functional hierarchies at different temperatures (Heschel et al., 2007).

Seedling De-etiolation

Dark-grown seedlings undergo skotomorphogenesis (etiolation) and are characterized by long hypocotyls, closed cotyledons and apical hooks, and development of the proplastids into etioplasts. Light-grown seedlings undergo photomorphogenesis (deetiolation) and are characterized by short hypocotyls, open and expanded cotyledons, and development of the proplastids into green mature chloroplasts (McNellis and Deng, 1995). Phytochromes perform a variety of overlapping functions in regulating seedling de-etiolation.

Mutants deficient in phyA display a wild-type photomorphogenic phenotype in W and R light. However, when grown in continuous FR light, phyA mutants display a skotomorphogenic phenotype (Figure 5), confirming that phyA is the primary photoreceptor responsible for perceiving and mediating various responses to FR light (Dehesh et al, 1993; Nagatani et al., 1993; Parks and Quail, 1993; Whitelam et al., 1993; Reed et al., 1994). Comparative transcriptional profiling of etiolated wild-type and phyA mutants subjected to FR light treatments revealed more than 800 phyA-regulated genes, providing the first insight into the phyA transcriptional network (Tepperman et al., 2001). It should be noted that phyA mutants also display elongated hypocotyls in continuous B light (Whitelam et al., 1993; Neff and Chory, 1998), suggesting that phyA also plays a pivotal role in perceiving and transducing B light.

phyB is the predominant phytochrome regulating de-etiolation in W and R light (Figure 5). However, transcriptional profiles of etiolated phyB mutants subjected to R light treatments did not differ dramatically from the wild-type controls (Tepperman et al., 2004). Subsequent studies showed that phyA plays a dominant role in regulating rapid gene expression responses to R light treatments (Tepperman et al., 2006). Moreover, the long hypocotyl and reduced cotyledon expansion phenotypes were enhanced in phyA phyB double mutants relative to phyB monogenic mutants in R light (Figure 5), revealing a role for phyA in responding to R light which is normally masked in the presence of phyB (Neff and Van Volkenburgh, 1994; Reed et al., 1994; Casal and Mazzella, 1998; Neff and Chory, 1998).

Mutants deficient in phyC exhibit a partial loss of sensitivity to R light, with longer hypocotyls and smaller cotyledons than the wild-type controls, indicating that phyC functions in regulating seedling de-etiolation in R light (Franklin et al., 2003a; Monte et al., 2003). However, no additive phenotype was observed in phyB phyC double mutants compared with their monogenic mutants, suggesting that phyC function is dependent on phyB (Monte et al., 2003). This might be explained by a recent finding that phyC does not homodimerize but rather forms heterodimers with phyB in vivo (Clack et al., 2009). Moreover, despite showing more sequence similarity to PHYA rather than to PHYB, PHYD and PHYE (Figure 3), phyC seems not to play a role in mediating seedling de-etiolation in FR light (Franklin et al., 2003a; Monte et al., 2003). However, the hypocotyls of phyA phyC double mutants are significantly longer than those of phyC monogenic mutants in R light (Franklin et al., 2003a; Monte et al., 2003), again confirming the contribution of phyA to seedling establishment under R light.

Figure 5.

Phenotypes of 4-d-old wild-type (WT), phyA, phyB and phyA phyB mutant plants grown in darkness (D) or under continuous white (W), far-red (FR), red (R) and blue (B) light conditions.

Although PHYD shows high sequence similarity to PHYB (Figure 3), the role of phyD in seedling de-etiolation in R seems minor, as it was reported that the Wassilewskija (Ws) ecotype of Arabidopsis contains a natural phyD deletion but its seedlings display only marginally longer hypocotyls in R than plants containing an introgressed PHYD gene (Aukerman et al., 1997). However, a synergistic relationship was observed between phyB and phyD in R, as the hypocotyls of phyB phyD double mutants are more than additively longer than those of each monogenic mutant (Aukerman et al., 1997). The contribution of phyE to seedling de-etiolation seems negligible, as it was shown that monogenic phyE mutants were indistinguishable from wild-type control plants in a variety of light conditions (Devlin et al., 1998).

As mentioned above, phytochromes also weakly absorb B light (Figure 1). In addition, phytochromes were shown to modulate cryptochrome-mediated seedling de-etiolation and phototropin-mediated phototropic curvature of Arabidopsis hypocotyls in B light (Parks et al., 1996; Ahmad and Cashmore, 1997; Hamazato et al., 1997; Janoudi et al., 1997; Casal and Mazzella, 1998; Neff and Chory, 1998). Direct physical interactions between phytochromes and cryptochromes were also reported (Ahmad et al., 1998; Mas et al., 2000). Therefore, phytochromes co-act with B/UV-A light photoreceptors to regulate seedling de-etiolation in B light.

Shade Avoidance

Plant development is regulated not only by the difference between light and darkness, but also by light quality, in particular the change of light quality due to shading by other plants. Light passed through or reflected from living vegetation is depleted in R and B wavebands, which are absorbed by chlorophyll and carotenoid pigments used for photosynthesis, leading to a reduction in the ratio of R to FR wavelengths (R:FR). This allows plants to initiate a suite of developmental responses called shade avoidance syndrome (SAS), which elevates leaves towards unfiltered daylight and enables plants to overtop competitors (reviewed in Smith and Whitelam, 1997). These responses include elongation of stems and petioles, accelerated flowering time, and increased apical dominance. The ability of plants to monitor their light environments and change their architecture provides them with a competitive strategy to survive and complete their life cycle in dense stands.

Reductions in R:FR ratio favor the conversion of phytochrome molecules to their inactive Prform. Therefore the shade avoidance syndrome must be suppressed under high R:FR ratio conditions. In this sense, shade avoidance is due to the relief of suppression rather than the induction of physiological responses. phyB is the predominant suppressor of shade avoidance responses in high R:FR, as phyB-deficient plants display a constitutive shade avoidance phenotype (elongated petiole and early flowering) (Nagatani et al., 1991; Somers et al., 1991; Figure 6). The shade avoidance responses enabled by low R:FR ratios can be effectively phenocopied by end-of-day far-red (EOD-FR) treatments. This led to the discovery of the roles of phyD and phyE in shade avoidance. Although the monogenic phyD mutant plants have no obviously abnormal phenotype to EOD-FR, plants impaired in both phyB and phyD display significantly longer hypocotyls under either R or W light, and flower earlier than the phyB monogenic mutants, suggesting that phyB and phyD function redundantly in suppressing shade avoidance (Aukerman et al., 1997; Devlin et al., 1999). As with phyD, monogenic phyE mutants show no phenotypic alterations unless in the phyB mutant background, and the phyB phyE double mutants flower much earlier than the phyB monogenic mutants (Devlin et al., 1998; Franklin et al., 2003b).

Figure 6.

Phenotypes of 3-week-old wild-type (WT), phyA, phyB, phyA phyB, phyB phyD phyE, phyA phyB phyD phyE plants grown under white light conditions (16-h light/8-h dark).

As phyA is the primary photoreceptor sensing FR wavelengths, enrichment of FR in transmitted/reflected light can lead to enhanced phyA signaling in the HIR mode. Therefore, the action of phyA can substitute for the loss of phyB, phyD and phyE activity due to their conversion to the inactive Pr forms. Indeed, Arabidopsis phyA mutant seedlings display enhanced shade avoidance responses relative to wild-type control plants when grown in low R:FR conditions (Johnson et al., 1994; Smith et al., 1997; Salter et al., 2003). Despite the relatively close phylogenetic relationship between phyA and phyC (Figure 3), no role for phyC in mediating shade avoidance responses has been reported. This conclusion is further confirmed by the observations that phyA phyB phyD phyE quadruple mutants display insensitivity to reductions in R:FR ratio and EOD-FR treatments (Franklin et al., 2003b).

LIGHT-REGULATED SUBCELLULAR LOCALIZATION OF PHYTOCHROMES

As mentioned above, phytochromes are synthesized in the cytosol in their inactive Pr forms. It was widely accepted before the mid-1990s that phytochromes were cytoplasmic photoreceptors based on early biochemical and immunocytochemical studies (Nagy and Schafer, 2002). However, extensive studies conducted in the last decade have established the notion that phytochromes must enter the nucleus to trigger most light responses (Nagatani, 2004; Kevei et al., 2007; Fankhauser and Chen, 2008). Thus, light-regulated translocation of the photoreceptors from the cytoplasm into the nucleus is a key event in the phytochrome signaling cascade. Recent publications are beginning to shed light on the molecular mechanisms underlying this central control step.

Regulation of phyB Nuclear Localization

phyB nuclear accumulation is efficiently initiated by continuous R light, and to a lesser extent by continuous B light, but completely ineffective by FR light. Single pulses of R, FR and B light cannot induce phyB nuclear accumulation (Gil et al., 2000). Moreover, phyB nuclear transport by R light is reversible by FR light, a typical characteristic of LFR (Kircher et al., 1999). A similar regulation of subcellular localization was also reported for phyC, phyD and phyE (Kircher et al., 2002). It should be noted that there is a weak, but detectable level of phyB-phyE present in the nucleus of dark-grown plants (Kircher et al., 2002).

A structure-function analysis first demonstrated that the C-terminal half of phyB (amino acids 594–1172) is sufficient to localize GUS to the nucleus, suggesting that the C-terminal domain of phyB harbors a putative nuclear localization signal (NLS) (Sakamoto and Nagatani, 1996). This result was confirmed by a later study, which showed that GFP fused to the N-terminal half of phyB (amino acids 1–651) localizes to the cytoplasm, whereas GFP fused to the C-terminal half of phyB (amino acids 625–1172) localizes to the nucleus (Matsushita et al., 2003). Subsequent further analyses of various truncations of the phyB C-terminal domain revealed that the PRD domain of phyB (amino acids 594–917) is both necessary and sufficient for nuclear localization, indicating that a putative NLS resides in this domain (Chen et al., 2005). Interestingly, this study also showed that the N-terminal photosensory GAF-PHY domains interact with the PRD domain in a light-dependent manner, thus providing a mechanistic link between light-dependent Pr/Pfr conformational alterations of phyB and the unmasking of the NLS that regulates phyB nuclear accumulation (Chen et al., 2005).

Regulation of phyA Nuclear Localization

The nuclear accumulation pattern of phyA is quite distinct from that of phyB. Firstly, all light illuminations (FR, R and B) are effective in inducing phyA nuclear translocation. A single, brief pulse of FR, R or B light induces phyA nuclear import as well (Hisada et al., 2000; Kim et al., 2000; Kircher et al., 2002). Therefore, phyA nuclear import is mediated by VLFR and HIR. Secondly, phyA nuclear translocation is very rapid (within minutes), whereas phyB nuclear import is relatively slow that takes hours (Kircher et al., 1999, Kim et al., 2000; Kircher et al., 2002). Finally, in contrast to phyB-phyE, phyA is exclusively localized in the cytosol in etiolated seedlings (Kircher et al., 2002).

FAR-RED ELONGATED HYPOCOTYL 1 (FHY1) and FHY1-LIKE (FHL). The rapid nuclear translocation of phyA, and the fact that phyA itself does not contain a typical NLS suggest the existence of an efficient transport machinery responsible for phyA nuclear import. Indeed, two small plant-specific proteins, FHY1 and FHL, have been shown to play an essential role in facilitating phyA nuclear translocation. The history of FHY1 essentially parallels the history of the molecular genetic analysis of phyA signaling pathway in Arabidopsis. The fhy1 mutant was firstly reported in 1993, together with two other mutants, i.e. fhy2 and fhy3, and all these fhy mutants develop elongated hypocotyls in FR light (Whitelam et al., 1993). This pioneering report showed that FHY2 locus corresponds to PHYA, while the FHY1 and FHY3 genes were cloned as separate loci in 2001 and 2002, respectively (Desnos et al., 2001; Wang and Deng, 2002). Subsequent studies identified a FHY1-like protein, named FHL, based on its sequence homology to FHY1 (Zhou et al., 2005).

FHY1 and FHL are two small proteins (202 and 201 amino acids, respectively) in Arabidopsis that were found to have homologs in both monocot and dicot plant species (Genoud et al., 2008; Li et al., 2010). Each protein contains an NLS and a nuclear exclusion signal (NES) at their N-termini and a septin-related domain (SRD) at their C termini (Desnos et al., 2001; Zhou et al., 2005). in vitro binding assays showed that both proteins are capable of homo- and hetero-dimerization through their C-terminal domains (Zhou et al., 2005). The NLS and SRD motifs are functionally important, because removal of either motif disrupts the function of FHY1 (Zeidler et al., 2004).

Earlier studies involving FHY1 suggested that FHY1 is responsible for mediating a branch of phyA signaling (Barnes et al., 1996). However, microarray analysis showed that all genes affected by phyA mutation are also affected by fhy1 mutation, although to a lesser degree (Wang et al., 2002). The roles of FHY1 in phyA signaling remained obscure until it was reported that FHY1/FHL physically interact with the Pfr form of phyA in vitro and in yeast cells through their SRD motifs, and that FHY1/FHL are required for nuclear accumulation of phyA since phyA is localized only in the cytosol of fhy1 fhl double mutants (Hiltbrunner et al., 2005, 2006). This conclusion was extended by a later report that the major function of FHY1/FHL is to act as adaptor proteins to chaperone photoactivated phyA into the nucleus (Genoud et al., 2008). Evidence supporting this proposed mode of action of FHY1 includes, first, sequence alignments show that the N-terminal NLS and the C-terminal phyA-interacting motifs are the only conserved motifs among all FHY1 homologs. Consistently, an artificial FHY1 consisting of a virus NLS motif and the C-terminal phyA-interacting motif of Arabidopsis FHY1 could rescue fhy1 mutant phenotypes and colocalizes with phyA in the nucleus. Second, FHY1 becomes functionally dispensable in transgenic seedlings expressing a constitutively nuclear phyA, i.e. if phyA could enter the nucleus by itself (Genoud et al., 2008).

FHY1/FHL specifically control the subcellular localization of phyA but not phyB, because fhy1/fhl mutants only show long-hypocotyl phenotype in FR but not R light, and phyB nuclear import is not affected in the fhy1 mutant (Hiltbrunner et al. 2005). As each of FHY1 and FHL has a functional monopartite NLS and NES that are indeed involved in the nuclear localization and exclusion of FHY1 (Zeidler et al., 2004), it is possible that phyA utilizes the NLS of FHY1/FHL for its nuclear transport. The fact that FHY1 homologs are widely distributed in angiosperms suggests that the mechanism uncovered in Arabidopsis may be conserved in higher plants (Genoud et al., 2008). However, interaction of phyA with FHY1/FHL alone appears to be insufficient for phyA nuclear translocation, because phyA-402, containing a missense mutation in the HKRD domain of phyA, is still capable of interacting with FHY1/FHL but does not translocate into the nucleus in R light (Muller et al., 2009).

Based on the evidence discussed above, one would assume that FHY1/FHL only function for phyA nuclear transport. However, this assumption was challenged by a recent report, which showed that FHY1/FHL might transmit phyA signals to downstream transcription factors (Yang et al., 2009). FHY1/FHL physically interact with two well-characterized transcription factors in phyA signaling network, LONG HYPOCOTYL IN FAR-RED 1 (HFR1) and LONG AFTER FAR-RED LIGHT 1 (LAF1), both in vitro and in vivo. Analysis of double and triple mutants showed that HFR1 and LAF1 independently transmit phyA signals downstream of FHY1 and FHL. Intriguingly, FHY1 was shown to mediate the assembly of a PHYA/FHY1/HFR1 signaling complex in vitro, suggesting that such kind of phyA signaling complexes may be assembled in vivo (Yang et al., 2009).

FHY3 and FAR-RED IMPAIRED RESPONSE1 (FAR1). The distinct role of FHY1/FHL in phyA nuclear accumulation suggests that any factor regulating FHY1/FHL transcriptionally or post-transcriptionally may indirectly affect phyA nuclear import. This notion is true, as demonstrated by the functional studies on FHY3 and FAR1. When the FHY3 gene was cloned in 2002, sequence alignments revealed that FHY3 shares high homology with FAR1, a previously identified phyA signaling component (Hudson et al., 1999; Wang and Deng, 2002). The function of FHY3 and FAR1 was not clear at that time, but some reports showed that FHY3 positively regulates the transcript levels of FHY1, and that FHY3/ FAR1 proteins share substantial similarity to Mutator-like element (MULE) transposases and may work as transcriptional regulators (Desnos et al., 2001; Hudson et al., 2003).

The breakthrough was made in 2007, when a study unequivocally demonstrated that FHY3 and FAR1 are transposase-derived transcription factors directly binding to the FHY1/FHL promoters via a specific cis-element called the FHY3/FAR1 binding site (FBS), and activating FHY1/FHL gene expression (Lin et al., 2007). The N-terminal C2H2 zinc finger domains of FHY3/FAR1 are essential for DNA binding, whereas the entire C-terminal regions are required for their transcriptional activation activity (Lin et al., 2007, 2008). Thus, FHY3 and FAR1 define a new type of transposase-derived transcription factors. Moreover, phyA nuclear accumulation is abolished in the fhy3 far1 double mutant, indicating that as the key transcriptional activators of FHY1/FHL expression, FHY3/FAR1 indirectly control phyA nuclear accumulation (Lin et al., 2007). This conclusion was further supported by another report that constitutively nuclear phyA could rescue the fhy3 mutant phenotypes (Genoud et al., 2008).

The discovery of FHY3/FAR1's role in phyA signaling invites the further question of how FHY1/FHL expression is down-regulated by phyA signaling. A negative feedback mechanism(s) controlling FHY1/FHL expression should exist, as previous studies showed that FHY1/FHL transcript levels were rapidly down-regulated when dark-grown plants were exposed to FR light (Desnos et al., 2001; Lin et al., 2007). A recent study showed that ELONGATED HYPOCOTYL 5 (HY5), a well-characterized bZIP transcription factor involved in promoting photomorphogenesis under various light conditions, plays a major role in this process (Li et al., 2010). HY5 achieves its goal by two distinct mechanisms. The first mechanism involves steric hindrance as HY5 directly binds ACGT-containing elements (ACEs) less than 10 bp away from the FHY3/FAR1 binding sites in the FHY1/FHL promoters. Thus, HY5's occupation of the ACEs consequently decreases the accessibility of the FHY1/FHL promoters to FHY3/FAR1. The second mechanism is called “sequestration” through the physical interactions between HY5 and FHY3/FAR1, a mechanism also used in the regulation of some plant bHLH transcription factors (de Lucas et al., 2008; Feng et al., 2008; Hornitschek et al., 2009). Therefore, HY5 acts as a repressor of FHY1/FHL expression by modulating the transcriptional activities of FHY3 and FAR1 (Li et al., 2010; Figure 7).

Nuclear Bodies (NBs)

Upon import into the nucleus, both phyA and phyB localize to discrete subnuclear foci, called nuclear bodies or speckles. Although most studies on NBs were conducted in cells overexpressing fluorescent-protein-tagged phytochromes, native phyA and phyB have each been shown to localize to NBs by immunocytochemical studies, suggesting that the formation of NBs is not an artifact because of the overexpression of phytochromes (Hisada et al., 2000; Kircher et al., 2002). The pattern of NBs is highly dynamic and directly regulated by light quality, quantity, and periodicity, and closely correlates to phytochrome-mediated responses (Kircher et al., 2002; Chen et al., 2003). However, the precise nature of NBs is still unknown. But several phytochrome signaling components colocalize to NBs, suggesting that NBs play important roles in phytochrome signaling (Chen, 2008).

Figure 7.

Control of FHY1/FHL expression and phyA nuclear accumulation.

FHY1 and FHL are required for phyA nuclear accumulation (Hiltbrunner et al., 2005, 2006; Genoud et al., 2008). FHY3 and FAR1 are two transposase-derived transcription factors that directly activate FHY1/FHL transcription, and thus indirectly regulate phyA nuclear accumulation and subsequent responses (Lin et al., 2007). phyA is localized exclusively in the cytosol in darkness in its inactive Pr form. Upon light exposure, the Pfr form of phyA is imported into the nucleus by FHY1/FHL, and thus triggers phyA signaling leading to multiple light responses, including the reduction of COP1 in the nucleus and accumulation of HY5 (Osterlund and Deng, 1998; Osterlund et al., 2000), and feedback regulation of FHY3 and FAR1 transcript levels (Lin et al., 2007). HY5 plays dual roles in phyA signaling: promoting photomorphogenesis, and down-regulating FHY1/FHL transcript levels by modulating the activities of the transcriptional activators FHY3 and FAR1 (Li et al., 2010). FHY3 and FHY1 (indicated by larger letters) are the more predominant players in the phyA signaling process compared to their respective homologs FAR1 and FHL. Pr: R-absorbing form of phyA (inactive); Pfr: FR-absorbing form of phyA (active). Arrow, positive regulation; bar, negative regulation. Image adapted from Li et al. (2010).

Based on the kinetics of phyB-GFP localization during the dark-to-light transition, two types of NBs have been defined for phyB. Within minutes of light exposure, small and transient phyBGFP NBs appear (Bauer et al., 2004). Interestingly, phyB colocalizes with PIF3 in the early transient NBs, and its localization to the early NBs is PIF3-dependent in vivo (Bauer et al., 2004). These early transient NBs disappear after 10–15 min in the light (Kevei et al., 2007), and interestingly, the disappearance of these early NBs correlates with the light-induced and phytochrome-dependent degradation of PIF3 (Al-Sady et al., 2006), thus implying that these early phyB NBs are the sites for phyB-PIF3 interaction, and their disappearance is due to PIF3 degradation (Chen, 2008).

After the disappearance of these early phyB NBs, longer R light treatment (2–3 h) leads to the appearance of larger and more stable NBs (Nagatani, 2004; Kevei et al., 2007; Chen, 2008). Interestingly, the size and number of these NBs depend on the fluence rate of R light (Chen et al., 2003). As increasing fluence rates of R not only induced a change in the pattern of phyB NBs but also enhanced inhibition of hypocotyl elongation, it was proposed that the formation of phyB NBs plays a role in the regulation of phyB-mediated signal transduction (Chen et al., 2003). The observations that PIF3 colocalizes with phyB only in the early but not late NBs indicate that the components of the early and late NBs may be different (Bauer et al., 2004).

phyA rapidly enters the nucleus in response to FR light and also forms early and late NBs (Bauer et al., 2004). PIF3 colocalizes with phyA in the early transient NBs but not late stable NBs (Bauer et al., 2004), consistent with the report that light-activated phyA also interacts with PIF3 and contributes to its degradation (Al-Sady et al., 2006). FHY1 and FHL, two phyA-interacting proteins required for phyA nuclear accumulation (see above), colocalize with phyA in the early transient NBs (Hiltbrunner et al., 2005, 2006). However, although a short R treatment also induces translocation of phyA into the nucleus, NB formation, and colocalization with PIF3, extended R light results in the complete loss of the phyA-GFP fluorescence (Bauer et al., 2004), possibly due to the photolabile nature and rapid degradation of phyA (see below).

Because both phyA and PIF3 are localized to NBs before their degradation, it has been proposed that NBs of phytochromes are sites for protein degradation (Bauer et al., 2004; Seo et al., 2004; Al-Sady et al., 2006). Recently, Chen et al. (2010) used a confocal microscopy-based screen to identify a gene, HEMERA (HMR), required for the localization of phyB-GFP to large NBs in high fluence rate of R light. Characterization of hmr mutants, localization of HMR protein within cells, and analysis of its biochemical function indicate that HMR is a specific and early phytochrome signaling component required for light-dependent proteolysis of phyA, PIF1, and PIF3 (Chen et al., 2010). Moreover, HMR is predicted to be structurally similar to the multiubiquitin-binding protein, RAD23, and can partially rescue yeast rad23 mutants, thus suggesting that phytochrome nuclear bodies may serve as sites of proteolysis (Chen et al., 2010).

PHYTOCHROMES AS LIGHT-REGULATED KINASES

How do phytochromes initiate their signal transduction upon photo-activation? A long-standing but much disputed hypothesis is that phytochromes act as light-regulated kinases (Wong et al., 1986; Kim et al., 1989). This hypothesis was initially supported by the observation that purified preparations of phyA catalyzed phosphorylation of serine residues on the photoreceptor itself, i.e. autophosphorylation activity (Wong et al., 1986; Yeh and Lagarias, 1998). The discovery of phytochrome-like photoreceptors in bacteria, collectively called bacteriophytochromes (BphPs), generated further supporting evidence for such a view (Fankhauser, 2000; Vierstra and Davis, 2000). Phytochrome-like sequences were identified in the cyanobacteria Fremeyella diplosiphon, Synechocystis sp. PCC6803, the purple photosynthetic bacterium Rhodospirillum and non-photosynthetic bacteria such as Deinococcus radiodurans, Pseudomononas putida and Pseudomonas aeruginosa (Kehoe and Grossman, 1996; Hughes et al., 1997; Davis et al., 1999b; Hughes and Lamparter, 1999; Jiang et al., 1999; Wu and Lagarias, 2000). Some of them, such as Cph1 of Synechocystis sp. PCC6803, can bind to the plant phytochrome chromophore (phytochromobilin PΦB or phycocyanobilin PCB) autocatalytically and display R/FR absorption spectra similar to plant phytochromes (Hughes et al., 1997; Lamparter et al., 1997; Yeh et al., 1997). Further, Cph1 was shown to be a light-regulated histidine kinase. Both autophosphorylation of Cph1 and transphosphorylation of Rcp1 (the response regulator for Cph1) are inhibited by R light and stimulated by FR light (Yeh et al., 1997), suggesting that in cyanobacteha phosphorylation is an important and very early step of phytochrome signal transduction.

Figure 8.

Structural comparison of Arabidopsis phytochromes and the bacterial phytochrome Cph1 (adapted from Yeh and Lagarias, 1998).

HKD, histidine kinase domain. The percent amino acid identities between the HKD domain of Cph1 and both PRD and HKRD domains of Arabidopsis phytochromes are indicated.

Image adapted from Yeh and Lagarias (1998). Reprinted with permission from the National Academy of Sciences.

However, higher plant phytochromes share limited sequence similarity with Cph1 at their C-termini (Figure 8). In addition, plant phytochromes have two additional domains compared with BphPs: a serine-rich NTE region, and a PRD domain located between PHY and HKRD domains (Figures 8). Moreover, mutating several critical residues required for bacterial His kinase activity does not affect the activity of plant phytochromes, suggesting that plant phytochromes are not active His kinases (Quail, 1997b). In fact, purified recombinant plant phytochromes exhibit a serine/threonine kinase activity, suggesting that eukaryotic phytochromes are histidine kinase paralogs with serine/threonine specificity (Yeh and Lagarias, 1998). Consistent with this discovery, several in vitro substrates of phytochrome kinase activity were subsequently discovered, such as histone H1, PHYTOCHROME KINASE SUBSTRATE 1 (PKS1), cryptochromes, AUX/IAA proteins, and FHY1 (Wong et al., 1989; Ahmad et al., 1998; Fankhauser et al., 1999; Colon-Carmona et al., 2000; Shen et al., 2009). However, the kinase domain of phytochrome has not been determined. It is notable that both PRD and HKRD domains show similarities to the HKD domain of Cph1 (Yeh and Lagarias, 1998; Figure 8). In addition, to date the kinase activity has only been proven for one higher plant phytochrome: oat phyA. Although a recent report indicated that phyB shows some kinase activity in vitro (Phee et al., 2008), more evidence is required before a firm conclusion can be reached. Whether all phytochromes have kinase activity, and whether different phytochromes behave differently as protein kinases need to be further characterized.

The claim that higher plant phytochromes function as protein kinases invites many questions, such as, is the Pfr form more active than the Pr form? What is the biological role of this kinase activity of phytochromes in plants? Answers to these questions have just begun to be unraveled. For example, autophosphorylation of recombinant oat phyA is both chromophore and light regulated, with Pfr being more active than Pr (Yeh and Lagarias, 1998). In vitro kinase assays also showed that the Pfr form of phytochrome more effectively phosphorylates some substrates, such as PKS1 and CRY1 (Ahmad et al., 1998; Fankhauser et al., 1999), but for some other substrates, such as AUX/IAA proteins and FHY1, the Pr and Pfr forms showed similar kinase activities (Colon-Carmona et al., 2000; Shen et al., 2009). However, in vivo assays are required to verify the conclusions of these experiments. For example, FHY1 phosphorylation in Arabidopsis seedlings is solely dependent on the active Pfr form of phyA, although both Pr and Pfr forms of phyA could phosphorylate FHY1 in vitro (Shen et al., 2009).

Recently it was shown that oat phyA autophosphorylates two serine sites in its NTE region in vitro (Han et al., 2010). Mutation of these two autophosphorylation sites in transgenic Arabidopsis plants caused hypersensitive light responses, indicating an increase in phyA activity (Han et al., 2010). Consistently, the degradation of the mutant phyA was significantly slower than the wild-type phyA under light conditions, suggesting that phyA autophosphorylation plays an important role in the regulation of phytochrome signaling through the control of phyA protein stability (Han et al., 2010). Another report showed that phyA is the only photoreceptor responsible for rapid R light-dependent FHY1 phosphorylation, and interestingly, this phosphorylation is R/FR light reversible, a typical LFR mode of phytochrome action (Shen et al., 2009). Notably, phosphorylated FHY1 was shown to be a preferred substrate for ubiquitin/26S proteasome-mediated degradation, suggesting that phyA-dependent FHY1 phosphorylation in R light may serve as a biochemical mechanism to desensitize FHY1-mediated phyA signaling (Shen et al., 2005b, 2009). These examples suggest that phytochrome autophosphorylation and kinase activity may play a negative role in light signal transduction.

However, the in vivo functional mechanism of phytochrome kinase activity is only beginning to be understood. For example, phytochrome kinase activity is stimulated in the presence of histone H1 in a Pr-specific manner, thus it is suggested that phytochrome kinase activity is activated in the nucleus that contains cationic molecules such as histones (Yeh and Lagarias, 1998; Kim et al., 2005; Han et al., 2010). Another example showing the complexity of in vivo phytochrome-related kinase activity comes from the study of the PHYTOCHROME-INTERACTING FACTORS (PIFs; see below). All PIFs except PIF7 are rapidly phosphorylated and then ubiquitinated in response to light in vivo prior to their degradation in a phytochrome-dependent manner (Al-Sady et al., 2008; Shen et al., 2007; Shen et al., 2008). A recent study reported that both phyA and phyB mediate PIF3 phosphorylation in vitro (Phee et al., 2008), suggesting that phytochromes may be the protein kinases responsible for phosphorylating the PIF proteins. However, compelling evidence supporting this assumption is still lacking. Recently, it was shown that CASEIN KINASE II (CK2), a highly conserved and ubiquitous Ser/Thr kinase, directly phosphorylates PIF1 (Bu et al., 2011). Therefore, whether phytochromes (both phyA and phyB) might directly phosphorylate PIFs in vivo, and how CK2 functions in phytochrome-induced rapid phosphorylation of PIFs await further investigation.

PHYTOCHROME SIGNALING INTERMEDIATES

The light signals perceived by the phytochrome photoreceptors are transduced to downstream signaling intermediates, which alter the expression of target genes and ultimately lead to the modulation of the biological responses (Quail, 2002; Jiao et al., 2007). Genetic research has identified a complex and interconnecting signaling network downstream of the phytochromes, together with a considerable number of positive or negative regulators, which act either in a specific pathway (such as LAF1 and HFR1) or in all branches of phytochrome signaling pathways (such as COP1 and HY5; Figure 9).

Negative Regulators of Phytochrome Signaling

Phylochrome-Interacting Factors (PIFs). Protein-protein interactions are necessary for many signal transduction cascades. Both general screenings for phytochrome-interacting proteins and targeted protein-protein interaction studies have identified a number of phytochtome-interacting partners. Those include PIF3 and other subsequently identified PIFs (Ni et al., 1998; Leivar and Quail, 2011), PKS1 (Fankhauser et al., 1999), NDPK2 (Choi et al., 1999), cryptochromes (both CRY1 and CRY2) (Ahmad et al., 1998; Mas et al., 2000), AUX/IAA proteins (Colon-Carmona et al., 2000), FyPP (Kim et al., 2002), COP1 (Seo et al., 2004; Jang et al., 2010), PAPP5 (Ryu et al., 2005), FHY1/FHL (Hiltbrunner et al., 2005, 2006), and were summarized recently by Bae and Choi (2008). Growing evidence demonstrates that the PIF proteins, a small subset of basic helix-loop-helix (bHLH) transcription factors, play central roles in phytochrome-mediated light signaling networks (Duek and Fankhauser, 2005; Castillon et al., 2007; Leivar and Quail, 2011).

The PIF proteins belong to the 15-member Subfamily 15 of the Arabidopsis bHLH transcription factor superfamily (Bailey et al., 2003; Heim et al., 2003; Toledo-Ortiz et al., 2003). PIF3 is the foundation member of the PIF subset initially identified in a yeast two-hybrid screen for phyB-interacting proteins (Ni et al., 1998). The second member of the PIF family, PIF4, was isolated by the convergence of both genetic and reverse-genetic approaches (Huq and Quail, 2002). Several other PIFs were then identified based on the sequence homology to PIF3, and were named PIF1, PIF5, PIF6 and PIF7 (Huq et al., 2004: Khanna et al., 2004; Oh et al., 2004; Leivar et al., 2008). As members of the bHLH superfamily transcription factors, all the PIF proteins contain a bHLH signature domain, consisting of a basic region (∼ 15 aa) involved in DNA binding and an HLH region (∼ 60 aa) involved in dimerization (Toledo-Ortiz et al., 2003; Castillon et al., 2007). The majority of the bHLH proteins bind to a cis-element called the E-box (CANNTG), whereas all the PIF proteins, where examined, bind specifically to a subtype of E-box, called the G-box (CACGTG) (Castillon et al., 2007; Leivar and Quail, 2011).

Figure 9.

A simplified model of the phytochrome signaling pathway.

phyA is the primary photoreceptor responsible for perceiving and mediating various responses to FR light, whereas phyB is the predominant phytochrome regulating responses to R light. Under light conditions, these photoreceptors act to suppress two main branches of light signaling: COP1-TFs and PIFs. COP1, whose activity is repressed by phytochromes in light conditions, is an E3 ubiquitin ligase targeting several photomorphogenesis-promoting transcription factors (such as HY5, HYH, LAF1 and HFR1) for degradation. PIFs are a subset of bHLH transcription factors required for skotomorphogenesis. Photo-activated phytochromes directly interact with PIFs, resulting in PIFs' phosphorylation and degradation, while COP1 positively regulates PIFs' protein levels. Phytochromes are targeted for degradation by COP1, and PIFs contribute to the degradation of phyB by promoting COP1/phyB interaction. Arrow, positive regulation; bar, negative regulation; solid line, direct regulation; dotted line, indirect regulation. Image adapted from Lau and Deng (2010). Reprinted with permission from Elsevier.

All PIF members contain a conserved motif in their N-termini, designated active phytochrome B-binding (APB) motif, which confers specific binding of PIFs to the biologically active Pfr form of phyB (Khanna et al., 2004; Duek and Fankhauser, 2005; Castillon et al., 2007; Leivarand Quail, 2011). However, only two PIF proteins, PIF1 and PIF3, also bind to the Pfr form of phyA, with PIF1 showing much stronger affinity for phyA than PIF3 (Ni et al., 1998; Huq et al., 2004). Accordingly, PIF1 and PIF3 each contain a motif called active phytochrome A-binding (APA) necessary for binding to phyA, but the actual sequences of these two APA motifs are not conserved (Al-Sady et al., 2006; Shen et al., 2008).

The recent finding that a quadruple mutant of PIFs, pif1 pif3 pif4 pif5 (pifq), develops a constitutively photomorphogenic (cop)like phenotype in the dark provides compelling evidence that the PIF proteins repress photomorphogenesis and promote skotomorphogenesis in etiolated seedlings (Leivar et al., 2008; Shin et al., 2009; Quail, 2011). Consistently, microarray analysis showed that the dark-grown pifq mutant has a gene expression pattern similar to that of R light-grown wild-type plants (Shin et al., 2009). By comparing rapidly light-responsive genes in wild-type seedlings with those responding in darkness in the pifq mutant, an overlapping subset of genes were identified as potential direct targets of these bHLH transcription factors (Leivar et al., 2009). Notably, transcription factor—encoding genes are highly enriched among these genes, suggesting that they may be potential primary targets of PIF transcriptional regulation.

At the same time, evidence obtained in the last decade demonstrates that one way phytochromes promote photomorphogenesis is by inducing rapid (within minutes) phosphorylation of most, if not all PIFs, upon light exposure. This subsequently leads to their ubiquitination and degradation via the ubiquitin/proteasome system (Bauer et al., 2004; Park et al., 2004; Shen et al., 2005a; Al-Sady et al., 2006; Oh et al., 2006; Nozue et al., 2007; Shen et al., 2007; Al-Sady et al., 2008; Lorrain et al., 2008; Shen et al., 2008). The interaction between phytochromes and PIFs is necessary for the light-dependent phosphorylation of PIFs, because mutant PIF1 and PIF3 proteins that abolish interactions with the Pfr forms of phyA and phyB do not undergo light-dependent phosphorylation (Al-Sady et al., 2006; Shen et al., 2008). Moreover, upon returning light-grown plants to darkness, PIF proteins rapidly re-accumulate to high levels. Subsequent re-exposure to light once again induces rapid degradation of PIFs (Leivar and Quail, 2011), indicating that this rapid regulation of PIFs is dynamic and controlled by phytochrome LFRs. Therefore, phytochrome-induced phosphorylation and proteolysis of PIFs may represent a major biochemical mechanism of signal transfer from the photoactivated phytochromes to their interacting signaling partners in the nucleus, which rapidly alters the gene expression profiles of the genome. In addition to controlling phytochrome-regulated gene expression, PIF proteins were recently shown to also modulate phyB abundance in R light, possibly by stimulating COP1-catalyzed ubiquitination and degradation of phyB (see below) (Khanna et al., 2007; Leivar et al., 2008; Al-Sady et al., 2008; Jang et al., 2010; Figure 9).

COP/DET/FUS Proteins. Genetic screens for Arabidopsis mutants involved in light-regulated seedling development followed by biochemical analyses have identified a group of pleiotropic Constitutive Photomorphogenic/De-etiolated/Fusca (COP/DET/FUS) proteins that are central negative regulators of photomorphogenesis (Sullivan et al., 2003; Yi and Deng, 2005). This group of COP/ DET/FUS proteins defines three biochemical entities: the COP1SPA complexes, the COP9 signalosome (CSN), and the CDD complex (COP10, DDB1, and DET1), all of which are involved in proteasomal degradation of photomorphogenesis-promoting factors (Saijo et al., 2003; Serino and Deng, 2003; Yanagawa et al., 2004; Yi and Deng, 2005; Zhu et al., 2008). Interestingly, the COP1 -SPA complexes and the CDD complex were recently shown to form two groups of CUL4-based E3 ligases in vivo (Chen et al., 2006, 2010). Therefore, these two groups of E3 ligases regulate the degradation of downstream factors to mediate light regulation of plant development (Chen et al., 2010).

COP1 is a conserved RING finger E3 ubiquitin ligase involved in multiple processes in many different organisms, including plant development and mammalian cell survival, growth, and metabolism (Yi and Deng, 2005). COP1 was first cloned and characterized in the model plant Arabidopsis as a repressor of light-regulated plant development (Deng et al., 1991, 1992; Figure 10). COP1 contains three domains: a RING finger domain in its N-terminal region, a WD40 repeat domain in its C-terminus, and a coiled-coil domain in the middle (Deng et al., 1992; Yi and Deng, 2005). COP1 has been shown to act as an E3 ligase targeting several photomorphogenesis-promoting proteins for degradation, including HY5 (Osterlund et al., 2000), HY5 HOMOLOG (HYH; Holm et al., 2002), LAF1 (Seo et al., 2003), HFR1 (Duek et al., 2004; Jang et al., 2005; Yang et al., 2005), and the phytochromes (Seo et al., 2004; Jang et al., 2010) (Figure 9). In addition, COP1 was recently shown to regulate flowering time by directly targeting transcriptional activator CONSTANS (CO) for degradation (Jang et al., 2008; Liu et al., 2008). Moreover, COP1 can interact with the substrate adaptor EARLY FLOWERING 3 (ELF3) to modulate light input signal to the circadian clock by destabilizing GIGANTEA (GI) protein (Yu et al., 2008).

SUPPRESSOR OF PHYA-105 (SPA1) was first identified as a repressor of phyA (Hoecker et al., 1998). Subsequent studies found three additional SPA1-like proteins in the Arabidopsis genome, named SPA2, SPA3, and SPA4 (Laubinger and Hoecker, 2003; Laubinger et al., 2004). Biochemical analysis demonstrated that SPA1 interacts with COP1 (Hoecker and Quail, 2001; Saijo et al., 2003; Seo et al., 2003), and interestingly, the SPA proteins can self-associate or interact with each other, forming a heterogeneous group of COP1/SPA complexes in Arabidopsis (Zhu et al., 2008). Genetic analysis showed that the four SPA genes are partially redundant in mediating light responses at both seedling and adult stages (Laubinger et al., 2004). Moreover, the quadruple spa mutant displays a phenotype similar to that of strong cop1 alleles (Laubinger et al., 2004), consistent with the notion that the SPA proteins work in concert with COP1 in controlling photomorphogenesis.

Figure 10.

Dark-grown cop1 mutant seedlings phenotypically mimic light-grown wild-type seedlings.

Empfindlicher Im Dunkelroten Licht 1 (EID1). EID1 is an F-box protein that functions as a negative regulator in phyA-specific light signaling (Buche et al., 2000; Dieterle et al., 2001). The fact that EID1 interacts with several Arabidopsis Skp1 -like (ASK) proteins and Cullin1 suggests that EID1 is a component of a SCF (SKP1/Cullin1/F-box protein) ubiquitin ligase complex targeting positively acting component(s) of phyA signaling pathway to ubiquitin-dependent proteolysis (Dieterle et al., 2001; Marrocco et al., 2006). A unique feature of the eid1 mutant is a shift in the peak of the action spectra of phyA-mediated hypocotyl elongation from FR to R part of the spectrum (Dieterle et al., 2001; Zhou et al., 2002). Although both EID1 and SPA1 function as negatively acting components in phyA-specific light signaling, mutant analysis indicated that EID1 and SPA1 have different but overlapping functions in phyA-dependent signal transduction chains (Zhou et al., 2002). Interestingly, a L946F mutation in the HKRD domain of phyA (named phyA-402 allele) was found to suppress the hypersensitive phenotype of the eid1–3 mutant (Muller et al., 2009). However, when phyA-402 is introgressed into the wild-type background, only moderate phenotype was observed, indicating that the mutation mainly alters phyA functions in an EID1-dependent signaling cascade (Muller et al., 2009).

Positive Regulators of Phytochrome Signaling

HY5 and HYH. HY5, a constitutively nuclear bZIP protein, is the first known and most extensively studied transcription factor involved in promoting photomorphogenesis under a wide spectrum of wavelengths, including FR, R, B, and UV-B (Koornneef et al., 1980; Oyama, et al., 1997; Osterlund et al., 2000; Ulm et al., 2004). It was shown that the abundance of HY5 protein is directly correlated with the extent of photomorphogenic development (Osterlund et al., 2000). Recent chromatin immunoprecipitation (ChIP)-chip studies revealed that HY5 binds directly to a large number of genomic sites, mainly at the promoter regions of annotated genes (Lee et al., 2007; Zhang et al., 2011). It seems that HY5 directly mediates both upregulation and downregulation of gene expression by light. The gene expression regulation attributable to HY5 is included largely within genes that are regulated by light and comprises ∼20% of all light-regulated genes (Ma et al., 2002). Therefore, HY5 is likely to be a high hierarchical regulator of the transcriptional cascades involved in seedling photomorphogenesis (Lee et al., 2007).

COP1 is capable of directly interacting with HY5 in the nucleus through its WD40 repeat domain and targets HY5 for proteasome-mediated degradation (Ang et al., 1998; Osterlund et al., 2000). HY5 has a homolog in the Arabidopsis genome, named HYH, and interestingly, HYH was also shown to be a target of COP1 (Holm et al., 2002). HY5 and HYH physically interact with COP1 through a COP1-interaction motif (Holm et al., 2001, 2002). Consistent with the finding that COP1 forms protein complexes with the SPA proteins, SPA1 contributes to the down-regulation of HY5 abundance (Saijo et al., 2003). Multiple photoreceptors, including phytochromes and cryptochromes, promote the accumulation of HY5 under specific light conditions, possibly by reducing the nuclear abundance of COP1 (Osterlund and Deng, 1998; Osterlund et al., 2000). However, little is known as to how the light-activated photoreceptors regulate the activities of COP1, as well as other COP/DET/FUS proteins.

HFR1 and LAF1. HFR1, an atypical bHLH protein, was originally identified as a positive regulator specific to phyA signaling (Fairchild et al., 2000; Fankhauser and Chory, 2000; Soh et al., 2000). However, subsequent studies revealed that HFR1 is also a component of cry1-mediated B light signaling (Duek and Fankhauser, 2003). Thus, HFR1 may represent a point of signal integration from phyA and cry1, either as a convergence of two independent signaling pathways or as a result of interaction of phyA and cry1 at the photoreceptor molecule level (Ahmad et al., 1998). It was demonstrated that HFR1 is capable of forming homodimers as well as heterodimers with PIF3. However, in contrast to PIF3, HFR1 does not bind directly to either phyA or phyB, although the HFR1/PIF3 complex can bind preferentially to the Pfr form of both phyA and phyB (Fairchild et al., 2000). In addition, unlike the PIF proteins, HFR1 contains an atypical basic domain which might not be functional for directly binding to DNA (Fairchild et al., 2000; Heim et al., 2003). Consistent with this proposal, HFR1 was recently shown to prevent an exaggerated shade avoidance response by forming non-DNA-binding heterodimers with PIF4 and PIF5, two bHLH transcription factors directly regulating the expression of shade-responsive marker genes (Sessa et al., 2005; Lorrain et al., 2008; Hornitschek et al., 2009; Galstyan et al., 2011). Moreover, the finding that a stabilized version of HFR1 leads to a constitutively photomorphogenic phenotype in darkness (similar to that of the pif1 pif3 pif4 pif5 quadruple mutants) suggest that HFR1 may function to sequester all these PIF proteins (Yang et al., 2003; Leivar et al., 2008; Shin et al., 2009).

LAF1 is an R2R3-MYB transcription factor with trans-activation activity, and functions as a positive component of phyA signaling (Ballesteros, et al., 2001). However, no direct target gene of LAF1 has been reported so far. The hfr1 laf1 double mutant has an additive phenotype compared to the two single mutants, indicating that HFR1 and LAF1 regulate largely independent pathways (Jang et al., 2007). Interestingly, both HFR1 and LAF1 were found to be the targets of COP1's E3 ubiquitin ligase activity in vitro, and in both cases, genetic and physical interactions between HFR1/LAF1 and COP1 were also observed (Seo et al., 2003; Jang et al., 2005; Yang et al., 2005). Moreover, it was recently shown that HFR1 also physically interacts with LAF1, and this interaction stabilizes each other through inhibition of ubiquitination by COP1, thereby enhancing phyA photoresponses (Jang et al., 2007).

PHYTOCHROME CONTROL OF NUCLEAR GENE EXPRESSION

Microarray analyses conducted in the last decade revealed genome-wide gene expression profiles regulated by light. About 10% or so of the genes in the Arabidopsis genome display phytochrome-regulated changes in expression during the seedling de-etiolation transition triggered by initial exposure of etiolated seedlings to light (Tepperman et al., 2004, 2006; Quail, 2011). These genes include numerous photosynthetic genes related to the biogenesis of active chloroplasts, various auxin-, gibberellin-, cytokinin- and ethylene hormone pathway-related genes potentially mediating growth responses, and metabolic genes reflecting the transition from heterotrophic to autotrophic growth (Tepperman et al., 2004, 2006; Quail, 2011). It is believed that the changes in expression of this large number of light-responsive genes ultimately lead to various morphogenic changes during seedling de-etiolation. Significantly, among functionally classifiable early light-responsive genes responding within 1 hour of FR or R light exposure, 44% (for FR light) and 25% (for R light) encode transcription factors (Tepperman et al., 2001, 2004, 2006), suggesting that they may representa master set of transcriptional regulators that orchestrate the expression of the downstream target genes in the phytochrome-directed transcriptional network.

Extensive studies have shed light on the mechanisms by which phytochromes regulate light-responsive gene expression. Firstly, phytochromes may promptly alter the expression of a large number of genes by inducing rapid phosphorylation and proteolysis of PIF transcription factors, as discussed above. Secondly, it is generally assumed that phytochromes rapidly inactivate the COP/ DET/FUS proteins in response to light, which leads to the accumulation of photomorphogenesis-promoting transcription factors, such as HY5, HYH, LAF1 and HFR1 although the mechanisms governing this process are largely unknown (Figure 9). In addition, it has been proposed that the direct protein-protein interactions between phytochromes and COP1/SPA proteins might be responsible for the rapid, initial inactivation of COP1 activity, whereas long-term inactivation of COP1 is achieved by subsequent depletion of the molecule from the nucleus (Wang and Deng, 2003).

Moreover, it has been suggested that phytochromes might directly target light signals to the light-responsive gene promoters. phyB was shown to bind reversibly to G-box-bound PIF3 specifically upon light-triggered conversion of the photoreceptor to its biologically active Pfr form, thus suggesting a provocative model in which phytochromes may function as integral light-switchable components of transcription regulator complexes, permitting direct targeting of light signals to target gene promoters (Martinez-Garcia et al., 2000; Quail, 2002). The direct interactions of PIF1 and PIF3 with phyA suggest that phyA could be targeted to gene promoters as well. However, conclusive evidence in favor of this model is still lacking. There is no evidence that phytochromes are associated with DNA in vivo, and that phytochromes indeed modulate transcription on the target gene promoters.

PHYTOCHROME SIGNALING AND THE CIRCADIAN CLOCK

The circadian clock controls many metabolic, developmental and physiological processes in a time-of-day-specific manner in both plants and animals (McClung, 2008; de Montaigu et al., 2010). Although circadian rhythms are endogenously generated, they can be modulated by external cues such as light and temperature, thus allowing plants to anticipate and adapt to daily and seasonal changes in their environment. Light signals perceived and transduced by phytochromes and cryptochromes ensure the clock is in tune with the external light/dark cycles. This process is known as entrainment. phyA, phyB, phyD, and phyE act as photoreceptors in R light input to the clock, while phyA and the cryptochromes cry1 and cry2 act as photoreceptors in B light input (Devlin and Kay, 2000). Interestingly, it was shown that phyA acts in low-intensity R light for circadian control, while phyB functions in high-intensity R light (Somers et al., 1998). In FR light, phyA is expected to be the only active photoreceptor transducing the light input to the circadian clock (Wenden et al., 2011). A recent report showed that Arabidopsis mutants deficient in all five phytochromes still displayed clock-controlled robust rhythmic oscillations of leaf position, indicating that phytochromes are not part of the core mechanism of the circadian clock and that other photoreceptors are sufficient for entrainment (Strasser et al., 2010). It should be noted that the LOV/F-box/Kelch-repeat family photoreceptors ZTL, LKP2 and FKF are also involved in the regulation of the circadian clock in Arabidopsis (Nelson et al., 2000; Somers et al., 2000; Schultz et al., 2001; Imaizumi et al., 2003; Mas et al., 2003; Somers et al., 2004; Kim et al., 2007; Demarsy and Fankhauser, 2009; Baudry et al., 2010).

The rhythm of leaf movement in the wild-type Arabidopsis plants can be reset by FR light, but this resetting was absent in the phyA, fhy1 and fhy3 mutants, suggesting that phyA signaling pathway is required for the entrainment of the circadian clock (Yanovsky et al., 2000, 2001). Interestingly, the fhy3 mutant also shows an enhanced response to R light during seedling de-etiolation, and shows disrupted rhythmicity of central-clock and clockoutput gene expression in continuous R light (Allen et al., 2008). Further, FHY3 is required for the clock resetting in response to R light pulses, suggesting that FHY3 functions in gating phytochrome signaling to the circadian clock (Allen et al., 2006). Recently, FHY3 and its homolog FAR1 were shown to bind directly to the promoter of EARLY FLOWERING 4 (ELF4), a component of the central oscillator of Arabidopsis circadian clock (Li et al., 2011). Interestingly, HY5, a well-characterized bZIP transcription factor involved in promoting photomorphogenesis, and CIRCADIAN CLOCK ASSOCIATED 1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY), two MYB-related transcription factors that are key components of the central oscillator, were also shown to bind directly to the ELF4 gene promoter via their respective cis-elements. FHY3, FAR1 and HY5 activate ELF4 expression during the day, whereas CCA1 and LHY suppress ELF4 expression periodically at dawn. Thus, this set of light- and circadian-regulated transcription factors act directly and coordinately at the ELF4 promoter to regulate its cyclic expression (Li et al., 2011). In addition, phyA itself is essential for the expression of ELF4 (Khanna et al., 2003), and ELF4 is a likely target mediating phyA-transduced FR light input to the circadian clock (Wenden et al., 2011). Moreover, phyA was shown to directly associate with FHY3 in vivo (Saijo et al., 2008). These data suggest that FHY3 may represent a potential molecular link directly gating the environmental light signals perceived by phytochrome to the circadian clock. It is also interesting to note that FHY3 directly binds to the promoter regions of other circadian genes such as CCA1 and CCA1 HIKING EXPEDITION (CHE) (Li et al., 2011; Ouyang et al., 2011), while its co-regulator HY5 binds to the promoters of CCA1, LHY, TOC1, ELF3 and GI (Lee et al., 2007). Thus, whether FHY3 could regulate other clock genes (such as CCA1) together with HY5 will be an intriguing question for future studies.

Another mechanism of gating the light input to the circadian clock, called the external coincidence model, has been proposed in the recent years. In Arabidopsis, CONSTANS (CO) plays a central role in the induction of flowering by long days (LDs) (Putterill et al., 1995; Robson et al., 2001). CO encodes a nuclear protein containing zinc fingers, and activates the transcription of floral regulators such as FLOWERING LOCUS T(FT) in the light. The mRNA level of CO is tightly controlled by the circadian clock and shows a striking temporal pattern of expression. Under LDs CO mRNA peaks before dusk and stays high until the following dawn, whereas under short days (SDs) CO mRNA peaks during the night (Suarez-Lopez et al., 2001). Under these conditions, when CO mRNA expression coincides with the exposure of plants to light, photoreceptors enhance the level and activity of the CO protein (Valverde et al., 2004). CO then triggers flowering by activating FT transcription. Genetic experiments demonstrate that photoreceptors cry1, cry2 and phyA promote flowering and stabilize the CO protein, whereas phyB delays flowering and promotes the degradation of CO (Johnson et al., 1994; Guo et al., 1998; El-Assal et al., 2001; Yanovsky and Kay, 2002; Cerdan and Chory, 2003; Searle and Coupland, 2004; Valverde et al., 2004). Therefore, the coincidence between CO mRNA and exposure to light (thus CO stability could be regulated post-transcoptionally by photoreceptors) is required to promote flowering.

On the other hand, several clock-related genes are involved in phytochrome signaling. For example, GIGANTEA (GI) encodes a novel protein found only in plants, and regulates flowering, circadian rhythms, and seedling photomorphogenesis under continuous R and B light (Araki and Komeda, 1993; Fowler et al., 1999; Park et al., 1999; Huq et al., 2000; Tseng et al., 2004; Mizoguchi et al., 2005; Martin-Tryon et al., 2007). Interestingly, although GI does not affect the phyA HIRs, it does affect phyA-mediated VLFRs via mechanisms that do not obviously involve its circadian functions (Oliverio et al., 2007). ELF3 is a highly conserved plant-specific nuclear protein which has been suggested to be part of the central clock oscillator and to act as a link between light and the circadian clock (Hicks et al., 1996; Zagotta et al., 1996; McWatters et al., 2000; Covington et al., 2001; Hicks et al., 2001; Liu et al., 2001; Thines and Harmon, 2010). Like phyB mutants, both gi and elf3 mutants display elongated hypocotyls in R light (Zagotta et al., 1996; Huq et al., 2000; Reed et al., 2000). However, the gi mutants are late flowering, which is in contrast with the early flowering phenotype of the phyB and elf3 mutants, suggesting that ELF3 and Gl play different roles or use different mechanisms in controlling hypocotyl elongation and flowering responses. Interestingly, ELF3 was shown to interact with both phyB and GI (Liu et al., 2001; Yu et al., 2008). In addition, ELF3 may act as an adaptor protein allowing COP1 to interact with GI which leads to GI degradation (Yu et al., 2008).

PHOSPHORYLATION/DEPHOSPHORYLATION AND DEGRADATION OF PHYTOCHROMES

Phosphorylation and Dephosphorylation of Phytochromes

It is well established that phytochromes are phosphoproteins, because they could be readily labeled with 32P isotope in vivo (Wong et al., 1986; Biermann et al., 1994). The phosphorylation sites of oat phyA have been investigated since the 1980s. There are two phosphorylation sites (Ser8 and Ser18) in the NTE region, and one site (Ser599) in the hinge region of oat phyA, among which, Ser8 and Ser599 were confirmed as in vivo phosphorylation sites (Wong et al., 1986; McMichael and Lagarias, 1990; Lapko et al., 1997, 1999). Ser8 is constitutively phosphorylated in both the Pr and Pfr forms, whereas Ser599 phosphorylation is light dependent and is preferentially phosphorylated in the Pfr form in vivo (Lapko et al., 1997, 1999).

The observation that phytochromes are phosphoproteins suggests the existence of protein kinase(s) and phosphatase(s) responsible for phosphorylating and dephosphorylating phytochromes. However, despite the extensive investigations of phytochrome-interacting proteins, there is no report thus far of a protein kinase that can specifically phosphorylate phytochromes. Instead, as mentioned above, two phosphorylation sites in the NTE region of oat phyA, i.e. Ser8 and Ser18, are autophosphorylated by phyA itself in vitro (Han et al., 2010). Mutations of these two serines to alanines effectively abolished autophosphorylation of oat phyA in vitro (Han et al., 2010). However, another phosphorylation site of oat phyA, Ser599, is not autophosphorylated by phyA (Kim et al., 2004; Han et al., 2010). Therefore, the protein kinase(s) responsible for phosphorylating this site of oat phyA will be a major interest for future studies.

In contrast to the failure of identifying a phytochrome-associated kinase, there are several studies reporting the discovery of protein phosphatases that could dephosphorylate phytochromes. A phytochrome-associated protein phosphatase 2A, designated FyPP, was shown to interact with and dephosphorylate phyA (Kim et al., 2002). The transgenic Arabidopsis plants with over-expressed or suppressed FyPP levels exhibited delayed or accelerated flowering, respectively, indicating that FyPP modulates phytochrome-mediated light signaling in the timing of flowering (Kim et al., 2002). PHYTOCHROME-ASSOCIATED PROTEIN PHOSPHATASE 5 (PAPP5) was isolated from yeast two-hybrid screen using the full-length Arabidopsis phyA as bait (Ryu et al., 2005). PAPP5 interacts with both phyA and phyB, and dephosphorylates all three phosphor-serine residues of oat phyA in a Pfr-specific manner. It was shown that PAPP5 positively influences the protein stability of phytochrome and the interaction of phytochrome with a downstream signal transducer NDPK2 (Ryu et al., 2005). Another phosphatase, PHYTOCHROME-ASSOCIATED PROTEIN PHOSPHATASE 2C (PAPP2C), was recently shown to interact with both phyA and phyB and effectively dephosphorylate phytochromes in vitro (Phee et al., 2008).

What are the in vivo functional roles of phytochrome phosphorylation and dephosphorylation? Recent studies suggest two possible roles: controlling phytochrome stability, and regulating interactions of phytochromes with downstream signal transducers. In the last two decades, great efforts have been made to investigate the role of a stretch of serine residues (including Ser8 and Ser18 of oat phyA) in the first 20 amino acids of phyA proteins, and it has been well-documented that substitution of these serines to alanines, or deletion of this 20-aa region, results in an increased biological activity of phyA, suggesting that these serine residues are involved in desensitization of phyA signaling (Stockhaus et al., 1992; Jordan et al., 1996, 1997; Casal et al., 2002; Han et al., 2010). Recently it was shown that the increased activity of mutated phyA proteins is related to the increased phyA stability (Han et al., 2010). Therefore, it is likely that autophosphorylation of serine sites in the NTE of phyA controls phyA stability.

However, phosphorylation of another serine residue, Ser599 in the hinge region of oat phyA, does not affect phytochrome stability (Kim et al., 2004). Instead, it was shown that phosphorylation of Ser599 prevents the interaction of phyA with its signal transducers such as NDPK2 and PIF3, suggesting that phosphorylation of Ser599 serves as a signal modulating switch affecting protein-protein interactions between phytochrome and its signal transducers (Kim et al., 2004). Interestingly, an earlier study showed that if Ser599 was mutated (Ser599Lys), phyA autophosphorylation and phosphorylation of its substrate PKS1 were no longer regulated by light (Fankhauser et al., 1999), suggesting that Ser599 plays an important role in the regulation of phytochrome kinase activity as well.

A recent study firstly detected an in vivo phosphorylated form of Arabidopsis phyA protein (Saijo et al., 2008). It was shown that phosphorylated phyA preferentially associates with COP1/ SPA1 complex for degradation, whereas unphosphorylated phyA predominantly associates with phyA signaling intermediates FHY1 and FHY3 for signal transduction. Interestingly, COP1 associates with unphosphorylated phyA in the absence of FHY1 or FHY3, suggesting that phyA associations with FHY1 and FHY3 protect unphosphorylated phyA from being recognized by COP1/SPA1 complex (Saijo et al., 2008). Therefore, these data suggest that light-induced phyA phosphorylation in vivo acts as a switch controlling differential interactions of the photoreceptor with signal propagation or attenuation machineries. However, how many phosphorylation sites contribute to produce this phosphorylated form of Arabidopsis phyA protein needs to be further characterized.

Degradation of Phytochromes

As mentioned above, phyA is a light labile phytochrome. phyA is most abundant in etiolated seedlings, but its level drops up to 100-fold after exposure to light (Clough and Vierstra, 1997; Sharrock and Clack, 2002). The proteolysis of phyA following photoconversion from Pr to Pfr is rapid, as the Pr form has a half-life of approximately 1 week, whereas the Pfr form has a half-life of only 1–2 h (Clough and Vierstra, 1997). It was observed more than two decades ago that phyA is ubiquitinated in vivo, and is degraded by the ubiquitin/26S proteasome pathway (Shanklin et al., 1987; Jabben et al., 1989a, b; Clough and Vierstra, 1997).

However, the E3 ligase responsible for phyA ubiquitination had not been identified until 2004, when COP1 was shown to ubiquitinate phyA in vitro (Seo et al., 2004). COP1 and phyA colocalize in the nuclear bodies, and the two proteins physically interact with each other mediated by the PRD domain of phyA and the WD40 domain of COP1. Consistently, the degradation rate of phyA is decreased in cop1 mutants (Seo et al., 2004). This finding was extended by a later report that the light signaling repressors, SPA proteins, contribute to COP1-mediated degradation of phyA, and that a COP1/SPA1 protein complex is tightly associated with phyA ubiquitination activity (Saijo et al., 2008). Consistent with the notion that phyA phosphorylation status controls its protein stability, the phosphorylated phyA form was suggested to be a preferred target for COP1-mediated degradation (Saijo et al., 2008).

Moreover, it is notable that light-induced phyA degradation still occurs in the null cop1–5 allele, suggesting the presence of a COP1-independent phyA-degradation pathway (Saijo et al., 2008). Consistent with this assumption, it was observed long ago that a R light pulse promotes the rapid formation of phyA-containing cytosolic spots, referred to as sequestered areas of phytochormes (SAPs), which appear prior to the nuclear transport of phyA and is thought to be the place of phyA ubiquitination and degradation (Speth et al., 1986; Nagatani, 2004; Kevei et al., 2007). The cytosolic degradation of phyA is also supported by the recent reports that phyA is still degraded in the fhy1 fhl double mutants, in which phyA is maintained in the cytosol, although at a slower rate than in the wild-type plants (Rosler et al., 2007; Debrieux and Fankhauser, 2010).

Although the type II phytochromes (phyB-phyE in Arabidopsis) are described as “light stable”, they are slowly degraded upon irradiation with R light (Sharrock and Clack, 2002). Recently, a report showed that phyB is stable in darkness, but in R, a fraction of phyB translocates into the nucleus and becomes degraded by the ubiquitin/26S proteasome pathway (Jang et al., 2010). Nuclear phyB degradation is also mediated by COP1 E3 ligase, which preferentially interacts with the N-terminal region of phyB. Interestingly, phyB polyubiquitination by COP1 in vitro can be enhanced by different PIF proteins that promote COP1/phyB interaction. Consistent with these results, nuclear phyB accumulates to higher levels in pif and cop1 mutants. Moreover, COP1 was shown to interact with and ubiquitinate phyC-phyE in vitro (Jang et al., 2010). Therefore, COP1 is an E3 ligase for all five phytochromes in Arabidopsis.

CYTOPLASMIC EVENTS OF PHYTOCHROME SIGNALING

Besides nuclear signaling events, phytochrome signaling likely entails cytoplasmic events based on the following reasons. Firstly, as mentioned above, the nuclear import of some phytochrome species (such as phyB) takes hours, and their nuclear actions obviously could not explain some rapid phytochrome responses which occur within minutes of light irradiation, such as the change in hypocotyl growth rate (Parks and Spalding, 1999). Secondly, phytochromes interact with a number of cytoplasmic proteins, such as PKS1 and NDPK2 (Choi et al., 1999; Fankhauser et al., 1999). Thirdly, genetic studies have identified several cytoplasmic proteins as signaling intermediates, such as PAT1 and FIN219 for phyA signaling (Bolle et al., 2000; Hsieh et al., 2000).

The fastest phytochrome response described so far is the cytoplasmic streaming of Vallisneria gigantea, which can be locally stimulated with pulsed R light and becomes measurable within 2.5 s, and is likely mediated by phyB-type phytochromes (Takagi et al., 2003). As FHY1/FHL are indispensible for phyA nuclear accumulation (see above), the cytoplasmic responses regulated by phyA were identified recently using the fhy1 fhl double mutant (Rosler et al., 2007). The phyA-specific cytoplasmic responses include the R-enhanced phototropism, abrogation of gravitropism in B, and the inhibition of hypocotyl elongation in B (Rosler et al., 2007). It is noteworthy that the latter two cytoplasmic responses of phyA occur in B light, which might be explained in part by a recent observation that phyA coordinates the localization and distribution of PHOT1 (Han et al., 2008).

Previous microinjection and pharmacological studies suggested the involvement of G proteins, cGMP and Ca2+/calmodulin in phytochrome signaling (Bowler et al., 1994; Neuhaus et al., 1997). In particular, it was reported that transgenic Arabidopsis plants conditionally overexpressing the α subunit (encoded by GPA1 gene) of the heterotrimeric G protein under the control of a glucocorticoid-inducible promoter exhibited a light-dependent hypersensitive response as a result of reduced hypocotyl cell elongation (Okamoto et al., 2001). However, a separate study reported that loss-of-function gpa1 mutants display partial de-etiolation in the dark, with short hypocotyls and open apical hooks typical of light-irradiated seedlings (Ullah et al., 2001). The short hypocotyl of gpa1 seedlings was reportedly due to a defect in cell division, but not cell elongation (Ullah et al., 2001). To resolve these discrepancies regarding GPA1 function in photomorphogenesis, Jones et al. (2003) re-evaluated the roles of the heterotrimeric G protein employing multiple alleles of gpa1 and agb1 (impaired in the β subunit) and their double mutants. They concluded that these mutants have wild-type sensitivity to R and FR light. In addition, there is no apparent alteration in R or FR sensitivity in transgenic plants overexpressing GPA1 or AGB1 The observed shorter hypocotyls and partially open cotyledons of gpa1 and agb1 mutants grown in the dark are mainly caused by a defect in cell division, rather than cell elongation (Jones et al., 2003).

Genetic studies have also provided supporting evidence for the involvement of Ca2+ in phytochrome signaling. The sub1 mutant exhibits hypersensitive responses to both FR and B light (Guo et al., 2001). The SUB1 gene was found to encode a Ca2+-binding protein. Genetic interaction studies suggest that SUB1 is a component of cryptochrome signaling pathway and is a modulator of phyA signaling pathway. Further, SUB1 negatively regulates HY5, a photomorphogenesis-promoting transcription factor (Guo et al., 2001).

FUTURE ISSUES

The last two decades have seen significant progress in unraveling the signaling mechanisms of plant phytochromes. However, several important questions about phytochromes still remain to be answered:

  1. We still do not understand the earliest signaling events following photo-activation. Which activities of phytochromes are sufficient to trigger light responses?

  2. Further to the first question, are phytochromes light-regulated kinases? If so, which domain is the kinase domain? What is the biological role of this kinase activity in inducing light responses?

  3. How do phytochromes induce the phosphorylation of PIFs? How are PIFs degraded?

  4. How are nuclear bodies formed? What are their precise functions in phytochrome signaling?

  5. Are phytochromes associated with the target gene promoters to directly regulate their gene expression?

  6. How do phytochromes inactivate the COP1/SPA protein degradation machinery upon photo-activation?

  7. How are phytochromes phosphorylated? In addition to autophosphorylation, which kinase(s) are responsible for phosphorylating phytochromes?

  8. How are phytochromes degraded in the nucleus and in the cytosol?

Further investigation and ultimate elucidation of these questions and others will undoubtedly shed more light on phytochrome signaling mechanisms and photomorphogenesis in general.

ACKNOWLEDGMENTS

We apologize to the authors whose work might have been overlooked here. We thank William Terzaghi for reading and commenting on the manuscript. This work was supported by a National Institute of Health grant (GM47850) to X.W.D. and a National Science Foundation award (IOS-1026630) to H.W. Xing Wang Deng's laboratory at Peking University was supported by grants from the Ministry of Science and Technology of China (2009DFB30030) and the Ministry of Agriculture of China (2009ZX08012-021B).

REFERENCES

  1. M. Ahmad , and A.R. Cashmore (1997). The blue-light receptor cryptochrome 1 shows functional dependence on phytochrome A or phytochrome B in Arabidopsis thaliana. Plant J. 11: 421–427. Google Scholar
  2. M. Ahmad , J.A. Jarillo , O. Smirnova , and A.R. Cashmore (1998). The CRY1 blue light photoreceptor of Arabidopsis interacts with phytochrome A in vitro. Mol. Cell 1: 939–948. Google Scholar
  3. T. Allen , A. Koustenis , G. Theodorou , D.E. Somers , S.A. Kay , G.C. Whitelam , and P.F. Devlin (2006). Arabidopsis FHY3 specifically gates phytochrome signaling to the circadian clock. Plant Cell 18: 2506–2516. Google Scholar
  4. B. Al-Sady , E.A. Kikis , E. Monte , and P.H. Quail (2008). Mechanistic duality of transcription factor function in phytochrome signaling. Proc. Natl. Acad. Sci. USA 105: 2232–2237. Google Scholar
  5. B. Al-Sady , W. Ni , S. Kircher , E. Schafer , and P.H. Quail (2006). Photoactivated phytochrome induces rapid PIF3 phosphorylation prior to proteasome-mediated degradation. Mol. Cell 23: 439–446. Google Scholar
  6. F. Andel 3rd, J.C. Lagarias , and R.A. Mathies (1996). Resonance raman analysis of chromophore structure in the lumi-R photoproduct of phytochrome. Biochemistry 35: 15997–16008. Google Scholar
  7. L.H. Ang , S. Chattopadhyay , N. Wei , T. Oyama , K. Okada , A. Batschauer , and X.W. Deng (1998). Molecular interaction between COP1 and HY5 defines a regulatory switch for light control of Arabidopsis development. Mol. Cell 1: 213–222. Google Scholar
  8. T. Araki , and Y. Komeda (1993). Flowering in darkness in Arabidopsis thaliana. Plant J. 4: 801–811. Google Scholar
  9. M.J. Aukerman , M. Hirschfeld , L. Wester , M. Weaver , T. Clack , R.M. Amasino , and R.A. Sharrock (1997). A deletion in the PHYD gene of the Arabidopsis Wassilewskija ecotype defines a role for phytochrome D in red/far-red light sensing. Plant Cell 9: 1317–1326. Google Scholar
  10. G. Bae , and G. Choi (2008). Decoding of light signals by plant phytochromes and their interacting proteins. Annu. Rev. Plant Biol. 59: 281–311. Google Scholar
  11. P.C. Bailey , C. Martin , G. Toledo-Ortiz , P.H. Quail , E. Huq , M.A. Heim , M. Jakoby , M. Werber , and B. Weisshaar (2003). Update on the basic helix-loop-helix transcription factor gene family in Arabidopsis thaliana. Plant Cell 15: 2497–2502. Google Scholar
  12. M.L. Ballesteros , C. Bolle , L.M. Lois , J.M. Moore , J.P. Vielle-Caizada , U. Grossniklaus , and N.H. Chua (2001). LAF1, a MYB transcription activator for phytochrome A signaling. Genes Dev. 15: 2613– 2625. Google Scholar
  13. S.A. Barnes , R.B. Quaggio , G.C. Whitelam , and N.H. Chua (1996). fhy1 defines a branch point in phytochrome A signal transduction pathways for gene expression. Plant J. 10: 1155–1161. Google Scholar
  14. A. Baudry , S. Ito , Y.H. Song , Strait A.A ., T. Kiba , S. Lu , R. Henriques , J.L. Pruneda-Paz , N.H. Chua , E.M. Tobin , S.A. Kay , and T. Imaizumi (2010). F-box proteins FKF1 and LKP2 act in concert with ZEITLUPE to control Arabidopsis clock progression. Plant Cell 22: 606–622. Google Scholar
  15. D. Bauer , A. Viczian , S. Kircher , T. Nobis , R. Nitschke , T. Kunkel , K.C. Panigrahi , E. Adam , E. Fejes , E. Schafer , and F. Nagy (2004). Constitutive Photomorphogenesis 1 and multiple photoreceptors control degradation of Phytochrome Interacting Factor 3, a transcription factor required for light signaling in Arabidopsis. Plant Cell 16: 1433–1445. Google Scholar
  16. B.J. Biermann , L.I. Pao , and L.J. Feldman (1994). Pr-specific phytochrome phosphorylation in vitro by a protein kinase present in antiphytochrome maize immunoprecipitates. Plant Physiol. 105: 243–251. Google Scholar
  17. C. Bolle , C. Koncz , and N.H. Chua (2000). PAT1, a new member of the GRAS family, is involved in phytochrome A signal transduction. Genes Dev. 14: 1269–1278. Google Scholar
  18. H.A. Borthwick , S.B. Hendricks , M.W. Parker , E.H. Toole , and V.K. Toole (1952). A reversible photoreaction controlling seed germination. Proc. Natl. Acad. Sci. USA 38: 662–666. Google Scholar
  19. J.F. Botto , R.A. Sanchez , G.C. Whitelam , and J.J. Casal (1996). Phytochrome A mediates the promotion of seed germination by very low fluences of light and canopy shade light in Arabidopsis. Plant Physiol. 110: 439–444. Google Scholar
  20. C. Bowler , G. Neuhaus , H. Yamagata , and N.H. Chua (1994). Cyclic GMP and calcium mediate phytochrome phototransduction. Cell 77: 73–81. Google Scholar
  21. M.T. Boylan , and P.H. Quail (1996). Are the phytochromes protein kinases? Protoplasma 195: 12–17 Google Scholar
  22. Q. Bu , L. Zhu , M.D. Dennis , L. Yu , S.X. Lu , M.D. Person , E.M. Tobin , K.S. Browning , and E. Huq (2011). Phosphorylation by CK2 enhances the rapid light-induced degradation of Phytochrome Interacting Factor 1 in Arabidopsis. J. Biol. Chem. 286: 12066–12074. Google Scholar
  23. C. Buche , C. Poppe , E. Schafer , and T. Kretsch (2000). eid1. a new Arabidopsis mutant hypersensitive in phytochrome A-dependent highirradiance responses. Plant Cell 12: 547–558. Google Scholar
  24. J.J. Casal , S.J. Davis , D. Kirchenbauer , A. Viczian , M.J. Yanovsky , R.C. Clough , S. Kircher , E.T. Jordan-Beebe , E. Schafer , F. Nagy , and R.D. Vierstra (2002). The serine-rich N-terminal domain of oat phytochrome A helps regulate light responses and subnuclear localization of the photoreceptor. Plant Physiol. 129: 1127–1137. Google Scholar
  25. J.J. Casal , and M.A. Mazzella (1998). Conditional synergism between cryptochrome 1 and phytochrome B is shown by the analysis of phyA, phyB, and hy4 simple, double, and triple mutants in Arabidopsis. Plant Physiol. 118: 19–25. Google Scholar
  26. J.J. Casal , R.A. Sanchez , and J.F. Botto (1998). Modes of action of phytochromes. J. Exp. Bot. 49: 127–138. Google Scholar
  27. A. Castillon , H. Shen , and E. Huq (2007). Phytochrome Interacting Factors: central players in phytochrome-mediated light signaling networks. Trends Plant Sci. 12: 514–521. Google Scholar
  28. P.D. Cerdan , and J. Chory (2003). Regulation of flowering time by light quality. Nature 423: 881–885. Google Scholar
  29. H. Chen , X. Huang , G. Gusmaroli , W. Terzaghi , O.S. Lau , Y. Yanagawa , Y. Zhang , J. Li , J.H. Lee , D. Zhu , and X.W. Deng (2010). Arabidopsis CULLIN4-damaged DNA binding protein 1 interacts with CONSTITUTIVELY PHOTOMORPHOGENIC1-SUPPRESSOR OF PHYA complexes to regulate photomorphogenesis and flowering time. Plant Cell 22: 108–123. Google Scholar
  30. H. Chen , Y. Shen , X. Tang , L. Yu , J. Wang , L. Guo , Y. Zhang , H. Zhang , S. Feng , E. Strickland , N. Zheng , and X.W. Deng (2006). Arabidopsis CULLIN4 forms an E3 ubiquitin ligase with RBX1 and the CDD complex in mediating light control of development. Plant Cell 18: 1991–2004. Google Scholar
  31. M. Chen (2008). Phytochrome nuclear body: an emerging model to study interphase nuclear dynamics and signaling. Curr. Opin. Plant Biol. 11: 503–508. Google Scholar
  32. M. Chen , R.M. Galvao , M. Li , B. Burger , J. Bugea , J. Bolado , and J. Chory (2010). Arabidopsis HEMERA/pTAC12 initiates photomorphogenesis by phytochromes. Cell 141: 1230–1240. Google Scholar
  33. M. Chen , R. Schwab , and J. Chory (2003). Characterization of the requirements for localization of phytochrome B to nuclear bodies. Proc Natl. Acad. Sci. USA 100: 14493–14498. Google Scholar
  34. M. Chen , Y. Tao , J. Lim , A. Shaw , and J. Chory (2005). Regulation of phytochrome B nuclear localization through light-dependent unmasking of nuclear-localization signals. Curr. Biol. 15: 637–642. Google Scholar
  35. G. Choi , H. Yi , J. Lee , Y.K. Kwon , M.S. Soh , B. Shin , Z. Luka , T.R. Hahn , and P.S. Song (1999). Phytochrome signalling is mediated through nucleoside diphosphate kinase 2. Nature 401: 610–613. Google Scholar
  36. T. Clack , S. Mathews , and R.A. Sharrock (1994). The phytochrome apoprotein family in Arabidopsis is encoded by five genes: the sequences and expression of PHYD and PHYE. Plant Mol. Biol. 25: 413–427. Google Scholar
  37. T. Clack , A. Shokry , M. Moffet , P. Liu , M. Faul , and R.A. Sharrock (2009). Obligate heterodimerization of Arabidopsis phytochromes C and E and interaction with the PIF3 basic helix-loop-helix transcription factor. Plant Cell 21: 786–799. Google Scholar
  38. R.C. Clough , and R.D. Vierstra (1997). Phytochrome degradation. Plant Cell Environ. 20: 713–721. Google Scholar
  39. A. Colon-Carmona , D.L. Chen , K.C. Yeh , and S. Abel (2000). Aux/ IAA proteins are phosphorylated by phytochrome in vitro. Plant Physiol. 124: 1728–1738. Google Scholar
  40. M.F. Covington , S. Panda , X.L. Liu , C.A. Strayer , D.R. Wagner , and S.A. Kay (2001). ELF3 modulates resetting of the circadian clock in Arabidopsis. Plant Cell 13: 1305–1315. Google Scholar
  41. S.J. Davis , S.H. Bhoo , A.M. Durski , J.M. Walker , and R.D. Vierstra (2001). The heme-oxygenase family required for phytochrome chromophore biosynthesis is necessary for proper photomorphogenesis in higher plants. Plant Physiol. 126: 656–669. Google Scholar
  42. S.J. Davis , J. Kurepa , and R.D. Vierstra (1999a). The Arabidopsis thaliana HY1 locus, required for phytochrome-chromophore biosynthesis, encodes a protein related to heme oxygenases. Proc. Natl. Acad. Sci. USA 96: 6541–6546. Google Scholar
  43. S.J. Davis , A.V. Vener , and R.D. Vierstra (1999b). Bacteriophytochromes: phytochrome-like photoreceptors from nonphotosynthetic eubacteria. Science 286: 2517–2520. Google Scholar
  44. M. de Lucas , J.M. Daviere , M. Rodriguez-Falcon , M. Pontin , J.M. Iglesias-Pedraz , S. Lorrain , C. Fankhauser , M.A. Blazquez , E. Titarenko , and S. Prat (2008). A molecular framework for light and gibberellin control of cell elongation. Nature 451: 480–484. Google Scholar
  45. A. de Montaigu , R. Toth , and G. Coupland (2010). Plant development goes like clockwork. Trends Genet. 26: 296–306. Google Scholar
  46. D. Debrieux , and C. Fankhauser (2010). Light-induced degradation of phyA is promoted by transfer of the photoreceptor into the nucleus. Plant Mol. Biol. 73: 687–695. Google Scholar
  47. K. Dehesh , C. Franci , B.M. Parks , K.A. Seeley , T.W. Short , J.M. Tepperman , and P.H. Quail (1993). Arabidopsis HY8 locus encodes phytochrome A. Plant Cell 5: 1081–1088. Google Scholar
  48. E. Demarsy , and C. Fankhauser (2009). Higher plants use LOV to perceive blue light. Curr. Opin. Plant Biol. 12: 69–74. Google Scholar
  49. X.W. Deng , T. Caspar , and P.H. Quail (1991). cop1: a regulatory locus involved in light-controlled development and gene expression in Arabidopsis. Genes Dev. 5: 1172–1182. Google Scholar
  50. X.W. Deng , M. Matsui , N. Wei , D. Wagner , A.M. Chu , K.A. Feldmann , and P.H. Quail (1992). COP1, an Arabidopsis regulatory gene, encodes a protein with both a zinc-binding motif and a G beta homologous domain. Cell 71: 791–801. Google Scholar
  51. X.W. Deng , and P.H. Quail (1999). Signalling in light-controlled development. Semin. Cell Dev. Biol. 10: 121–129. Google Scholar
  52. T. Desnos , P. Puente , G.C. Whitelam , and N.P. Harberd (2001). FHY1: a phytochrome A-specific signal transducer. Genes Dev. 15: 2980–2990. Google Scholar
  53. P.F. Devlin , and S.A. Kay (2000). Cryptochromes are required for phytochrome signaling to the circadian clock but not for rhythmicity. Plant Cell 12: 2499–2510. Google Scholar
  54. P.F. Devlin , S.R. Patel , and G.C. Whitelam (1998). Phytochrome E influences internode elongation and flowering time in Arabidopsis. Plant Cell 10: 1479–1487. Google Scholar
  55. P.F. Devlin , P.R. Robson , S.R. Patel , L. Goosey , R.A. Sharrock , and G.C. Whitelam (1999). Phytochrome D acts in the shade-avoidance syndrome in Arabidopsis by controlling elongation growth and flowering time. Plant Physiol. 119: 909–915. Google Scholar
  56. M. Dieterle , Y.C. Zhou , E. Schafer , M. Funk , and T. Kretsch (2001). EID1. an F-box protein involved in phytochrome A-specific light signaling. Genes Dev. 15: 939–944. Google Scholar
  57. P.D. Duek , M.V. Elmer , V.R. van Oosten , and C. Fankhauser (2004). The degradation of HFR1, a putative bHLH class transcription factor involved in light signaling, is regulated by phosphorylation and requires COP1. Curr. Biol. 14: 2296–2301. Google Scholar
  58. P.D. Duek , and C. Fankhauser (2003). HFR1, a putative bHLH transcription factor, mediates both phytochrome A and cryptochrome signalling. Plant J. 34: 827–836. Google Scholar
  59. P.D. Duek , and C. Fankhauser (2005). bHLH class transcription factors take centre stage in phytochrome signalling. Trends Plant Sci. 10: 51–54. Google Scholar
  60. K. Eichenberg , I. Baurle , N. Paulo , R.A. Sharrock , W. Rudiger , and E. Schafer (2000). Arabidopsis phytochromes C and E have different spectral characteristics from those of phytochromes A and B. FEBS Lett. 470: 107–112. Google Scholar
  61. S.E.D. El-Assal , C. Alonso-Blanco , A.J. Peeters , V. Raz , and M. Koornneef (2001). A QTL for flowering time in Arabidopsis reveals a novel allele of CRY2. Nat. Genet. 29: 435–440. Google Scholar
  62. C.D. Fairchild , M.A. Schumaker , and P.H. Quail (2000). HFR1 encodes an atypical bHLH protein that acts in phytochrome A signal transduction. Genes Dev. 14: 2377–2391. Google Scholar
  63. C. Fankhauser (2000). Phytochromes as light-modulated protein kinases. Semin. Cell. Dev. Biol. 11: 467–473. Google Scholar
  64. C. Fankhauser (2001). The phytochromes, a family of red/far-red absorbing photoreceptors. J. Biol. Chem. 276: 11453–11456. Google Scholar
  65. C. Fankhauser , and M. Chen (2008). Transposing phytochrome into the nucleus. Trends Plant Sci. 13: 596–601. Google Scholar
  66. C. Fankhauser , and J. Chory (2000). RSF1, an Arabidopsis locus implicated in phytochrome A signaling. Plant Physiol. 124: 39–45. Google Scholar
  67. C. Fankhauser , K.C. Yeh , J.C. Lagarias , H. Zhang , T.D. Elich , and J. Chory (1999). PKS1, a substrate phosphorylated by phytochrome that modulates light signaling in Arabidopsis. Science 284: 1539–1541. Google Scholar
  68. S. Feng , C. Martinez , G. Gusmaroli , Y. Wang , J. Zhou , F. Wang , L. Chen , L. Yu , J.M. Iglesias-Pedraz , S. Kircher , E. Schafer , X. Fu , L.M. Fan , and X.W. Deng (2008). Coordinated regulation of Arabidopsis thaliana development by light and gibberellins. Nature 451: 475–479. Google Scholar
  69. S. Fowler , K. Lee , H. Onouchi , A. Samach , K. Richardson , B. Morris , G. Coupland , and J. Putterill (1999). GIGANTEA: a circadian clock-controlled gene that regulates photoperiodic flowering in Arabidopsis and encodes a protein with several possible membrane-spanning domains. EMBO J. 18: 4679–4688. Google Scholar
  70. K.A. Franklin , S.J. Davis , W.M. Stoddart , R.D. Vierstra , and G.C. Whitelam (2003a). Mutant analyses define multiple roles for phytochrome C in Arabidopsis photomorphogenesis. Plant Cell 15: 1981– 1989. Google Scholar
  71. K.A. Franklin , U. Praekelt , W.M. Stoddart , O.E. Billingham , K.J. Halliday , and G.C. Whitelam (2003b). Phytochromes B, D, and E act redundantly to control multiple physiological responses in Arabidopsis. Plant Physiol. 131: 1340–1346. Google Scholar
  72. K.A. Franklin , and P.H. Quail (2010). Phytochrome functions in Arabidopsis development. J. Exp. Bot. 61: 11–24. Google Scholar
  73. M. Furuya , and P.S. Song (1994). Assembly and properties of holophytochrome. In Photomorphogenesis in plants, Kendrick R.E. , and Kronenberg G.H.M. , eds, (Dordrecht, The Netherlands: Kluwer Academic Publishers), pp. 105–140. Google Scholar
  74. A. Galstyan , N. Cifuentes-Esquivel , J. Bou-Torrent , and J.F. MartinezGarcia (2011). The shade avoidance syndrome in Arabidopsis: a fundamental role for atypical basic helix-loop-helix proteins as transcriptional cofactors. Plant J. 66: 258–267. Google Scholar
  75. W.W. Garner , and H.A. Allard (1920). Effect of the relative length of day and night and other factors of the environment on growth and reproduction in plants. J. Agric. Res. 18: 553–606. Google Scholar
  76. T. Genoud , F. Schweizer , A. Tscheuschler , D. Debrieux , J.J. Casai , E. Schafer , A. Hiltbrunner , and C. Fankhauser (2008). FHY1 mediates nuclear import of the light-activated phytochrome A photoreceptor. PLoS Genet. 4: e1000143. Google Scholar
  77. P. Gil , S. Kircher , E. Adam , E. Bury , L. Kozma-Bognar , E. Schafer , and F. Nagy (2000). Photocontrol of subcellular partitioning of phytochrome-B:GFP fusion protein in tobacco seedlings. Plant J. 22: 135–145. Google Scholar
  78. H. Guo , T. Mockler , H. Duong , and C. Lin (2001). SUB1. an Arabidopsis Ca2+-binding protein involved in cryptochrome and phytochrome coaction. Science 291: 487–490. Google Scholar
  79. H. Guo , H. Yang , T.C. Mockler , and C. Lin (1998). Regulation of flowering time by Arabidopsis photoreceptors. Science 279: 1360–1363. Google Scholar
  80. F. Hamazato , T. Shinomura , H. Hanzawa , J. Chory , and M. Furuya (1997). Fluence and wavelength requirements for Arabidopsis CAB gene induction by different phytochromes. Plant Physiol. 115: 1533– 1540. Google Scholar
  81. I.S. Han , T.S. Tseng , W. Eisinger , and W.R. Briggs (2008). Phytochrome A regulates the intracellular distribution of phototropin 1-green fluorescent protein in Arabidopsis thaliana. Plant Cell 20: 2835–2847. Google Scholar
  82. Y.J. Han , H.S. Kim , Y.M. Kim , A.Y. Shin , S.S. Lee , S.H. Bhoo , P.S. Song , and J.I. Kim (2010). Functional characterization of phytochrome autophosphorylation in plant light signaling. Plant Cell Physiol. 51: 596– 609. Google Scholar
  83. W. Haupt , and D.P. Hader (1994). Photomovement. In Photomorphogenesis in plants, R.E. Kendrick , and G.H.M. Kronenberg , eds, (Dordrecht, The Netherlands: Kluwer Academic Publishers), pp. 707–732. Google Scholar
  84. M.A. Heim , M. Jakoby , M. Werber , C. Martin , B. Weisshaar , and P.C. Bailey (2003). The basic helix-loop-helix transcription factor family in plants: a genome-wide study of protein structure and functional diversity. Mol. Biol. Evol. 20: 735–747. Google Scholar
  85. L. Hennig , W.M. Stoddart , M. Dieterle , G.C. Whitelam , and E. Schafer (2002). Phytochrome E controls light-induced germination of Arabidopsis. Plant Physiol. 128: 194–200. Google Scholar
  86. M.S. Heschel , J. Selby , C. Butler , G.C. Whitelam , R.A. Sharrock , and K. Donohue (2007). A new role for phytochromes in temperature-dependent germination. New Phytol. 174: 735–741. Google Scholar
  87. K.A. Hicks , T.M. Albertson , and D.R. Wagner (2001). EARLY FLOWERING3 encodes a novel protein that regulates circadian clock function and flowering in Arabidopsis. Plant Cell 13: 1281–1292. Google Scholar
  88. K.A. Hicks , A.J. Millar , I.A. Carre , D.E. Somers , M. Straume , D.R. Meeks-Wagner , and S.A. Kay (1996). Conditional circadian dysfunction of the Arabidopsis early-flowering 3 mutant. Science 274: 790–792. Google Scholar
  89. A. Hiltbrunner , A. Tscheuschler , A. Viczian , T. Kunkel , S. Kircher , and E. Schafer (2006). FHY1 and FHL act together to mediate nuclear accumulation of the phytochrome A photoreceptor. Plant Cell Physiol. 47: 1023–1034. Google Scholar
  90. A. Hiltbrunner , A. Viczian , E. Bury , A. Tscheuschler , S. Kircher , R. Toth , A. Honsberger , F. Nagy , C. Fankhauser , and E. Schafer (2005). Nuclear accumulation of the phytochrome A photoreceptor requires FHY1. Curr. Biol. 15: 2125–2130. Google Scholar
  91. M. Hirschfeld , J.M. Tepperman , T. Clack , P.H. Quail , and R.A. Sharrock (1998). Coordination of phytochrome levels in phyB mutants of Arabidopsis as revealed by apoprotein-specific monoclonal antibodies. Genetics 149: 523–535. Google Scholar
  92. A. Hisada , H. Hanzawa , J.L. Weller , A. Nagatani , J.B. Reid , and M. Furuya (2000). Light-induced nuclear translocation of endogenous pea phytochrome A visualized by immunocytochemical procedures. Plant Cell 12: 1063–1078. Google Scholar
  93. U. Hoecker , Y. Xu , and P.H. Quail (1998). SPA1: a new genetic locus involved in phytochrome A-specific signal transduction. Plant Cell 10: 19–33. Google Scholar
  94. U. Hoecker , and P.H. Quail (2001). The phytochrome A-specific signaling intermediate SPA1 interacts directly with COP1, a constitutive repressor of light signaling in Arabidopsis. J. Biol. Chem. 276: 38173– 38178. Google Scholar
  95. M. Holm , C.S. Hardtke , R. Gaudet , and X.W. Deng (2001). Identification of a structural motif that confers specific interaction with the WD40 repeat domain of Arabidopsis COP1. EMBO J. 20: 118–127. Google Scholar
  96. M. Holm , L.G. Ma , L.J. Qu , and X.W. Deng (2002). Two interacting bZIP proteins are direct targets of COP1-mediated control of light-dependent gene expression in Arabidopsis. Genes Dev. 16: 1247–1259. Google Scholar
  97. P. Hornitschek , S. Lorrain , V. Zoete , Q. Michielin , and C. Fankhauser (2009). Inhibition of the shade avoidance response by formation of non-DNA binding bHLH heterodimers. EMBO J. 28: 3893–3902. Google Scholar
  98. H.L. Hsieh , H. Okamoto , M. Wang , L.H. Ang , M. Matsui , H. Goodman , and X.W. Deng (2000). FIN219. an auxin-regulated gene, defines a link between phytochrome A and the downstream regulator COP1 in light control of Arabidopsis development. Genes Dev. 14: 1958–1970. Google Scholar
  99. M. Hudson , C. Ringli , M.T. Boylan , and P.H. Quail (1999). The FAR1 locus encodes a novel nuclear protein specific to phytochrome A signaling. Genes Dev. 13: 2017–2027. Google Scholar
  100. M.E. Hudson , D.R. Lisch , and P.H. Quail (2003). The FHY3 and FAR1 genes encode transposase-related proteins involved in regulation of gene expression by the phytochrome A-signaling pathway. Plant J. 34: 453–471. Google Scholar
  101. J. Hughes , and T. Lamparter (1999). Prokaryotes and phytochrome. The connection to chromophores and signaling. Plant Physiol. 121: 1059–1068. Google Scholar
  102. J. Hughes , T. Lamparter , F. Mittmann , E. Hartmann , W. Gartner , A. Wilde , and T. Borner (1997). A prokaryotic phytochrome. Nature 386: 663. Google Scholar
  103. E. Huq , B. Al-Sady , M. Hudson , C. Kim , K. Apel , and P.H. Quail (2004). Phytochrome-Interacting Factor 1 is a critical bHLH regulator of chlorophyll biosynthesis. Science 305: 1937–1941. Google Scholar
  104. E. Huq , and P.H. Quail (2002). PIF4. a phytochrome-interacting bHLH factor, functions as a negative regulator of phytochrome B signaling in Arabidopsis. EMBO J. 21: 2441–2450. Google Scholar
  105. E. Huq , J.M. Tepperman , and P.H. Quail (2000). GIGANTEA is a nuclear protein involved in phytochrome signaling in Arabidopsis. Proc. Natl. Acad. Sci. USA 97: 9789–9794. Google Scholar
  106. T. Imaizumi , H.G. Tran , T.E. Swartz , W.R. Briggs , and S.A. Kay (2003). FKF1 is essential for photoperiodic-specific light signalling in Arabidopsis. Nature 426: 302–306. Google Scholar
  107. M. Jabben , J. Shanklin , and R.D. Vierstra (1989a). Ubiquitin-phytochrome conjugates. Pool dynamics during in vivo phytochrome degradation. J. Biol. Chem. 264: 4998–5005. Google Scholar
  108. M. Jabben , J. Shanklin , and R.D. Vierstra (1989b). Red light-induced accumulation of ubiquitin-phytochrome conjugates in both monocots and dicots. Plant Physiol. 90: 380–384. Google Scholar
  109. I.C. Jang , R. Henriques , H.S. Seo , A. Nagatani , and N.H. Chua (2010). Arabidopsis PHYTOCHROME INTERACTING FACTOR proteins promote phytochrome B polyubiquitination by COP1 E3 ligase in the nucleus. Plant Cell 22: 2370–2383. Google Scholar
  110. I.C. Jang , J.Y. Yang , H.S. Seo , and N.H. Chua (2005). HFR1 is targeted by COP1 E3 ligase for post-translational proteolysis during phytochrome A signaling. Genes Dev. 19: 593–602. Google Scholar
  111. I.C. Jang , S.W. Yang , J.Y. Yang , and N.H. Chua (2007). Independent and interdependent functions of LAF1 and HFR1 in phytochrome A signaling. Genes Dev. 21: 2100–2111. Google Scholar
  112. S. Jang , V. Marchal , K.C. Panigrahi , S. Wenkel , W. Soppe , X.W. Deng , F. Valverde , and G. Coupland (2008). Arabidopsis COP1 shapes the temporal pattern of CO accumulation conferring a photoperiodic flowering response. EMBO J. 27: 1277–1288. Google Scholar
  113. A.K. Janoudi , W.R. Gordon , D. Wagner , P. Quail , and K.L. Poff (1997). Multiple phytochromes are involved in red-light-induced enhancement of first-positive phototropism in Arabidopsis thaliana. Plant Physiol. 113: 975–979. Google Scholar
  114. Z. Jiang , L.R. Swem , B.G. Rushing , S. Devanathan , G. Tollin , and C.E. Bauer (1999). Bacterial photoreceptor with similarity to photoactive yellow protein and plant phytochromes. Science 285: 406–409. Google Scholar
  115. Y. Jiao , O.S. Lau , and X.W. Deng (2007). Light-regulated transcriptional networks in higher plants. Nat. Rev. Genet. 8: 217–230. Google Scholar
  116. E. Johnson , M. Bradley , N.P. Harberd , and G.C. Whitelam (1994). Photoresponses of light-grown phyA mutants of Arabidopsis (phytochrome A is required for the perception of daylength extensions). Plant Physiol. 105: 141–149. Google Scholar
  117. A.M. Jones , J.R. Ecker , and J.G. Chen (2003). A reevaluation of the role of the heterotrimeric G protein in coupling light responses in Arabidopsis. Plant Physiol. 131: 1623–1627. Google Scholar
  118. E.T. Jordan , J.R. Cherry , J.M. Walker , and R.D. Vierstra (1996). The amino-terminus of phytochrome A contains two distinct functional domains. Plant J. 9: 243–257. Google Scholar
  119. E.T. Jordan , J.M. Marita , R.C. Clough , and R.D. Vierstra (1997). Characterization of regions within the N-terminal 6-kilodalton domain of phytochrome A that modulate its biological activity. Plant Physiol. 115: 693–704. Google Scholar
  120. C. Kami , S. Lorrain , P. Hornitschek , and C. Fankhauser (2010). Light-regulated plant growth and development. Curr. Top. Dev. Biol. 91: 29–66. Google Scholar
  121. D.M. Kehoe , and A.R. Grossman (1996). Similarity of a chromatic adaptation sensor to phytochrome and ethylene receptors. Science 273: 1409–1412. Google Scholar
  122. E. Kevei , E. Schafer , and F. Nagy (2007). Light-regulated nucleo-cytoplasmic partitioning of phytochromes. J. Exp. Bot. 58: 3113–3124. Google Scholar
  123. R. Khanna , E. Huq , E.A. Kikis , B. Al-Sady , C. Lanzatella , and P.H. Quail (2004). A novel molecular recognition motif necessary for targeting photoactivated phytochrome signaling to specific basic helix-loop-helix transcription factors. Plant Cell 16: 3033–3044. Google Scholar
  124. R. Khanna , E.A. Kikis , and P.H. Quail (2003). EARLY FLOWERING 4 functions in phytochrome B-regulated seedling de-etiolation. Plant Physiol. 133: 1530–1538. Google Scholar
  125. R. Khanna , Y. Shen , C.M. Marion , A. Tsuchisaka , A. Theologis , E. Schafer , and P.H. Quail (2007). The basic helix-loop-helix transcription factor PIF5 acts on ethylene biosynthesis and phytochrome signaling by distinct mechanisms. Plant Cell 19: 3915–3929. Google Scholar
  126. D.H. Kim , J.G. Kang , S.S. Yang , K.S. Chung , P.S. Song , and CEVI. Park (2002). A phytochrome-associated protein phosphatase 2A modulates light signals in flowering time control in Arabidopsis. Plant Cell 14: 3043–3056. Google Scholar
  127. I.S. Kim , U. Bai , and P.S. Song (1989). A purified 124-kDa oat phytochrome does not possess a protein kinase activity. Photochem. Photobiol. 49: 319–323. Google Scholar
  128. J.I. Kim , J.E. Park , X. Zarate , and P.S. Song (2005). Phytochrome phosphorylation in plant light signaling. Photochem. Photobiol. Sci. 4: 681–687. Google Scholar
  129. J.I. Kim , Y. Shen , Y.J. Han , J.E. Park , D. Kirchenbauer , M.S. Soh , F. Nagy , E. Schafer , and P.S. Song (2004). Phytochrome phosphorylation modulates light signaling by influencing the protein-protein interaction. Plant Cell 16: 2629–2640. Google Scholar
  130. L. Kim , S. Kircher , R. Toth , E. Adam , E. Schafer , and F. Nagy (2000). Light-induced nuclear import of phytochrome-A:GFP fusion proteins is differentially regulated in transgenic tobacco and Arabidopsis. Plant J. 22: 125–133. Google Scholar
  131. W.Y. Kim , S. Fujiwara , S.S. Suh , J. Kim , Y. Kim , L. Han , K. David , J. Putterill , H.G. Nam , and D.E. Somers (2007). ZEITLUPE is a circadian photoreceptor stabilized by GIGANTEA in blue light. Nature 449: 356–360. Google Scholar
  132. S. Kircher , P. Gil , L. Kozma-Bognar , E. Fejes , V. Speth , T. Husselstein-Muller , D. Bauer , E. Adam , E. Schafer , and F. Nagy (2002). Nucleocytoplasmic partitioning of the plant photoreceptors phytochrome A, B, C, D, and E is regulated differentially by light and exhibits a diurnal rhythm. Plant Cell 14: 1541–1555. Google Scholar
  133. S. Kircher , L. Kozma-Bognar , L. Kim , E. Adam , K. Harter , E. Schafer , and F. Nagy (1999). Light quality-dependent nuclear import of the plant photoreceptors phytochrome A and B. Plant Cell 11: 1445–1456. Google Scholar
  134. T. Kohchi , K. Mukougawa , N. Frankenberg , M. Masuda , A. Yokota , and J.C. Lagarias (2001). The Arabidopsis HY2 gene encodes phytochromobilin synthase, a ferredoxin-dependent biliverdin reductase. Plant Cell 13: 425–436. Google Scholar
  135. M. Koornneef , E. Rolff , and C.J.P. Spruit (1980). Genetic control of light-inhibited hypocotyl elongation in Arabidopsis thaliana (L.) Heynh. Z. Pflanzenphysiol. 100: 147–160. Google Scholar
  136. T. Kunkel , G. Neuhaus , A. Batschauer , N.H. Chua , and E. Schafer (1996). Functional analysis of yeast-derived phytochrome A and B phycocyanobilin adducts. Plant J. 10: 625–636. Google Scholar
  137. J.C. Lagarias , and H. Rapoport (1980). Chromopeptides from phytochrome. The structure and linkage of the Pr form of the phytochrome chromophore. J. Am. Chem. Soc. 102: 4821–4828. Google Scholar
  138. T. Lamparter , F. Mittmann , W. Gartner , T. Borner , E. Hartmann , and J. Hughes (1997). Characterization of recombinant phytochrome from the cyanobacterium Synechocystis. Proc. Natl. Acad. Sci. USA 94: 11792–11797. Google Scholar
  139. V.N. Lapko , X.Y. Jiang , D.L. Smith , and P.S. Song (1997). Posttranslational modification of oat phytochrome A: phosphorylation of a specific serine in a multiple serine cluster. Biochemistry 36: 10595–10599. Google Scholar
  140. V.N. Lapko , X.Y. Jiang , D.L. Smith , and P.S. Song (1999). Mass spectrometric characterization of oat phytochrome A: isoforms and posttranslational modifications. Protein Sci. 8: 1032–1044. Google Scholar
  141. O.S. Lau , and X.W. Deng (2010). Plant hormone signaling lightens up: integrators of light and hormones. Curr. Opin. Plant Biol. 13: 571–577. Google Scholar
  142. S. Laubinger , and U. Hoecker (2003). The SPA1-like proteins SPA3 and SPA4 repress photomorphogenesis in the light. Plant J. 35: 373–385. Google Scholar
  143. S. Laubinger , K. Fittinghoff , and U. Hoecker (2004). The SPA quartet: a family of WD-repeat proteins with a central role in suppression of photomorphogenesis in Arabidopsis. Plant Cell 16: 2293–2306. Google Scholar
  144. J. Lee , K. He , V. Stolc , H. Lee , P. Figueroa , Y. Gao , W. Tongprasit , H. Zhao , I. Lee , and X.W. Deng (2007). Analysis of transcription factor HY5 genomic binding sites revealed its hierarchical role in light regulation of development. Plant Cell 19: 731–749. Google Scholar
  145. P. Leivar , E. Monte , B. Al-Sady , C. Carle , A. Storer , J.M. Alonso , J.R. Ecker , and P.H. Quail (2008). The Arabidopsis phytochrome-interacting factor PIF7, together with PIF3 and PIF4, regulates responses to prolonged red light by modulating phyB levels. Plant Cell 20: 337– 352. Google Scholar
  146. P. Leivar , E. Monte , Y. Oka , T. Liu , C. Carle , A. Castillon , E. Huq , and P.H. Quail (2008). Multiple phytochrome-interacting bHLH transcription factors repress premature seedling photomorphogenesis in darkness. Curr. Biol. 18: 1815–1823. Google Scholar
  147. P. Leivar , J.M. Tepperman , E. Monte , R.H. Calderon , T.L. Liu , and P.H. Quail (2009). Definition of early transcriptional circuitry involved in light-induced reversal of PIF-imposed repression of photomorphogenesis in young Arabidopsis seedlings. Plant Cell 21: 3535–3553. Google Scholar
  148. P. Leivar , and P.H. Quail (2011). PIFs: pivotal components in a cellular signaling hub. Trends Plant Sci. 16: 19–28. Google Scholar
  149. G. Li , H. Siddiqui , Y. Teng , R. Lin , X.Y. Wan , J. Li , O.S. Lau , X. Ouyang , M. Dai , J. Wan , P.F. Devlin , X.W. Deng , and H. Wang (2011). Coordinated transcriptional regulation underlying the circadian clock in Arabidopsis. Nat. Cell. Biol. 13: 616–622. Google Scholar
  150. J. Li , G. Li , S. Gao , C. Martinez , G. He , Z. Zhou , X. Huang , J.H. Lee , H. Zhang , Y. Shen , H. Wang , and X.W. Deng (2010). Arabidopsis transcription factor ELONGATED HYPOCOTYL 5 plays a role in the feedback regulation of phytochrome A signaling. Plant Cell 22: 3634–3649. Google Scholar
  151. R. Lin , L. Ding , C. Casola D.R. Ripoll , C. Feschotte , and H. Wang (2007). Transposase-derived transcription factors regulate light signaling in Arabidopsis. Science 318: 1302–1305. Google Scholar
  152. R. Lin , Y. Teng , H.J. Park , L. Ding , C. Black , P. Fang , and H. Wang (2008). Discrete and essential roles of the multiple domains of Arabidopsis FHY3 in mediating phytochrome A signal transduction. Plant Physiol. 148: 981–992. Google Scholar
  153. L.J. Liu , Y.C. Zhang , Q.H. Li , Y. Sang , J. Mao , H.L. Lian , L. Wang , and H.Q. Yang (2008). COP1-mediated ubiquitination of CONSTANS is implicated in cryptochrome regulation of flowering in Arabidopsis. Plant Cell 20: 292–306. Google Scholar
  154. X.L. Liu , M.F. Covington , C. Fankhauser , J. Chory , and D.R. Wagner (2001). ELF3 encodes a circadian clock-regulated nuclear protein that functions in an Arabidopsis PHYB signal transduction pathway. Plant Cell 13: 1293–1304. Google Scholar
  155. S. Lorrain , T. Allen , P.D. Duek , G.C. Whitelam , and C. Fankhauser (2008). Phytochrome-mediated inhibition of shade avoidance involves degradation of growth-promoting bHLH transcription factors. Plant J. 53: 312–323. Google Scholar
  156. L. Ma , Y. Gao , L. Qu , Z. Chen , J. Li , H. Zhao , and X.W. Deng (2002). Genomic evidence for COP1 as a repressor of light-regulated gene expression and development in Arabidopsis. Plant Cell 14: 2383–2398. Google Scholar
  157. A.L. Mancinelli (1994). The physiology of phytochrome action. In Photomorphogenesis in plants, R.E. Kendrick , and G.H.M. Kronenberg , eds, (Dordrecht, The Netherlands: Kluwer Academic Publishers), pp. 211–269. Google Scholar
  158. K. Marrocco , Y. Zhou , E. Bury , M. Dieterle , M. Funk , P. Genschik , M. Krenz , T. Stolpe , and T. Kreisch (2006). Functional analysis of EID1, an F-box protein involved in phytochrome A-dependent light signal transduction. Plant J. 45: 423–438. Google Scholar
  159. J.F. Martinez-Garcia , E. Huq , and P.H. Quail (2000). Direct targeting of light signals to a promoter element-bound transcription factor. Science 288: 859–863. Google Scholar
  160. E.L. Martin-Tryon , J.A. Kreps , and S.L. Harmer (2007). GIGANTEA acts in blue light signaling and has biochemically separable roles in circadian clock and flowering time regulation. Plant Physiol. 143: 473–486. Google Scholar
  161. P. Mas , P.F. Devlin , S. Panda , and S.A. Kay (2000). Functional interaction of phytochrome B and cryptochrome 2. Nature 408: 207–211. Google Scholar
  162. P. Mas , W.Y. Kim , D.E. Somers , and S.A. Kay (2003). Targeted degradation of TOC1 by ZTL modulates circadian function in Arabidopsis thaliana. Nature 426: 567–570. Google Scholar
  163. J.L. Mateos , J.P. Luppi , O.B. Ogorodnikova , V.A. Sineshchekov , M.J. Yanovsky , S.E. Braslavsky , W. Gartner , and J.J. Casal (2006). Functional and biochemical analysis of the N-terminal domain of phytochrome A. J. Biol. Chem. 281: 34421–34429. Google Scholar
  164. S. Mathews , and R.A. Sharrock (1997). Phytochrome gene diversity. Plant Cell Environ. 20: 666–671. Google Scholar
  165. T. Matsushita , N. Mochizuki , and A. Nagatani (2003). Dimers of the N-terminal domain of phytochrome B are functional in the nucleus. Nature 424: 571–574. Google Scholar
  166. C.R. McClung (2008). Comes a time. Curr. Opin. Plant Biol. 11: 514–520. Google Scholar
  167. C.R. McClung , P.A. Salome , and T.P. Michael (2002). The Arabidopsis circadian system. The Arabidopsis Book 1: e0044. doi: 0010.1199/tab.0044Google Scholar
  168. R.W. McMichael Jr. , and J.C. Lagarias (1990). Phosphopeptide mapping of Avena phytochrome phosphorylated by protein kinases in vitro. Biochemistry 29: 3872–3878. Google Scholar
  169. T.W. McNellis , and X.W. Deng (1995). Light control of seedling morphogenetic pattern. Plant Cell 7: 1749–1761. Google Scholar
  170. H.G. McWatters , R.M. Bastow , A. Hall , and A.J. Millar (2000). The ELF3 Zeitnehmer regulates light signalling to the circadian clock. Nature 408: 716–720. Google Scholar
  171. T. Mizoguchi , L. Wright , S. Fujiwara , F. Cremer , K. Lee , H. Onouchi , A. Mouradov , S. Fowler , H. Kamada , J. Putteriil , and G. Coupland (2005). Distinct roles of GIGANTEA in promoting flowering and regulating circadian rhythms in Arabidopsis. Plant Cell 17: 2255–2270. Google Scholar
  172. E. Monte , J.M. Alonso , J.R. Ecker , Y. Zhang , X. Li , J. Young , S. Austin-Phillips , and P.H. Quail (2003). Isolation and characterization of phyC mutants in Arabidopsis reveals complex crosstalk between phytochrome signaling pathways. Plant Cell 15: 1962–1980. Google Scholar
  173. R. Muller , A.P. Fernandez , A. Hiltbrunner , E. Schafer , and T. Kretsch (2009). The histidine kinase-related domain of Arabidopsis phytochrome A controls the spectral sensitivity and the subcellular distribution of the photoreceptor. Plant Physiol. 150: 1297–1309. Google Scholar
  174. T. Muramoto , T. Kohchi , A. Yokota , I. Hwang , and H.M. Goodman (1999). The Arabidopsis photomorphogenic mutant hy1 is deficient in phytochrome chromophore biosynthesis as a result of a mutation in a plastid heme oxygenase. Plant Cell 11: 335–348. Google Scholar
  175. A.C. Mustilli , and C. Bowler (1997). Tuning in to the signals controlling photoregulated gene expression in plants. EMBO J. 16: 5801–5808. Google Scholar
  176. A. Nagatani (2004). Light-regulated nuclear localization of phytochromes. Curr. Opin. Plant Biol. 7: 708–711. Google Scholar
  177. A. Nagatani (2010). Phytochrome: structural basis for its functions. Curr. Opin. Plant Biol. 13: 565–570. Google Scholar
  178. A. Nagatani , J. Chory , and M. Furuya (1991). Phytochrome B is not detectable in the hy3 mutant of Arabidopsis, which is deficient in responding to end-of-day far-red light treatments. Plant Cell Physiol. 32: 1119–1122. Google Scholar
  179. A. Nagatani , J.W. Reed , and J. Chory (1993). Isolation and initial characterization of Arabidopsis mutants that are deficient in phytochrome A. Plant Physiol. 102: 269–277. Google Scholar
  180. F. Nagy , and E. Schafer (2002). Phytochromes control photomorphogenesis by differentially regulated, interacting signaling pathways in higher plants. Annu. Rev. Plant Biol. 53: 329–355. Google Scholar
  181. M.M. Neff , and J. Chory (1998). Genetic interactions between phytochrome A, phytochrome B, and cryptochrome 1 during Arabidopsis development. Plant Physiol. 118: 27–35. Google Scholar
  182. M.M. Neff , C. Fankhauser , and J. Chory (2000). Light: an indicator of time and place. Genes Dev. 14: 257–271. Google Scholar
  183. M.M. Neff , and E. Van Volkenburgh (1994). Light-stimulated cotyledon expansion in Arabidopsis seedlings (the role of phytochrome B). Plant Physiol. 104: 1027–1032. Google Scholar
  184. D.C. Nelson , J. Lasswell , L.E. Rogg , M.A. Cohen , and B. Bartel (2000). FKF1, a clock-controlled gene that regulates the transition to flowering in Arabidopsis. Cell 101: 331–340. Google Scholar
  185. J. Nemhauser , and J. Chory (2002). Photomorphogenesis. The Arabidopsis Book 1: e0054 doi:  0010.1199/tab.0054Google Scholar
  186. G. Neuhaus , C. Bowler , K. Hiratsuka , H. Yamagata , and N.H. Chua (1997). Phytochrome-regulated repression of gene expression requires calcium and cGMP. EMBO J. 16: 2554–2564. Google Scholar
  187. M. Ni , J.M. Tepperman , and P.H. Quail (1998). PIF3. a phytochromeinteracting factor necessary for normal photoinduced signal transduction, is a novel basic helix-loop-helix protein. Cell 95: 657–667. Google Scholar
  188. K. Nozue , M.F. Covington , P.D. Duek , S. Lorrain , C. Fankhauser , S.L. Harmer , and J.N. Maloof (2007). Rhythmic growth explained by coincidence between internal and external cues. Nature 448: 358–361. Google Scholar
  189. E. Oh , J. Kim , E. Park , J.I. Kim , C. Kang , and G. Choi (2004). PIL5, a phytochrome-interacting basic helix-loop-helix protein, is a key negative regulator of seed germination in Arabidopsis thaliana. Plant Cell 16: 3045–3058. Google Scholar
  190. E. Oh , S. Yamaguchi , Y. Kamiya , G. Bae , W.I. Chung , and G. Choi (2006). Light activates the degradation of PIL5 protein to promote seed germination through gibberellin in Arabidopsis. Plant J. 47: 124–139. Google Scholar
  191. Y. Oka , T. Matsushita , N. Mochizuki , T. Suzuki , S. Tokutomi , and A. Nagatani (2004). Functional analysis of a 450-amino acid N-terminal fragment of phytochrome B in Arabidopsis. Plant Cell 16: 2104–2116. Google Scholar
  192. H. Okamoto , M. Matsui , and X.W. Deng (2001). Overexpression of the heterotrimeric G-protein alpha-subunit enhances phytochrome-mediated inhibition of hypocotyl elongation in Arabidopsis. Plant Cell 13: 1639–1652. Google Scholar
  193. K.A. Oliverio , M. Crepy , E.L. Martin-Tryon , R. Milich , S.L. Harmer , J. Putterill , M.J. Yanovsky , and J.J. Casai (2007). GIGANTEA regulates phytochrome A-mediated photomorphogenesis independently of its role in the circadian clock. Plant Physiol. 144: 495–502. Google Scholar
  194. M.T. Osterlund , and X.W. Deng (1998). Multiple photoreceptors mediate the light-induced reduction of GUS-COP1 from Arabidopsis hypocotyl nuclei. Plant J. 16: 201–208. Google Scholar
  195. M.T. Osterlund , C.S. Hardtke , N. Wei , and X.W. Deng (2000). Targeted destabilization of HY5 during light-regulated development of Arabidopsis. Nature 405: 462–466. Google Scholar
  196. X. Ouyang , J. Li , G. Li , B. Li , B. Chen , H. Shen , X. Huang , X. Mo , X. Wan , R. Lin , S. Li , H. Wang , and X.W. Deng (2011). Genomewide binding site analysis of FAR-RED ELONGATED HYPOCOTYL 3 reveals its novel function in Arabidopsis development. Plant Cell 23: 2514–2535. Google Scholar
  197. T. Oyama , Y. Shimura , and K. Okada (1997). The Arabidopsis HY5 gene encodes a bZIP protein that regulates stimulus-induced development of root and hypocotyl. Genes Dev. 11: 2983–2995. Google Scholar
  198. D.H. Park , D.E. Somers , Y.S. Kim , Y.H. Choy , H.K. Lim , M.S. Soh , H.J. Kim , S.A. Kay , and H.G. Nam (1999). Control of circadian rhythms and photoperiodic flowering by the Arabidopsis GIGANTEA gene. Science 285: 1579–1582. Google Scholar
  199. E. Park , J. Kim , Y. Lee , J. Shin , E. Oh , W.I. Chung , J.R. Liu , and G. Choi (2004). Degradation of phytochrome interacting factor 3 in phytochrome-mediated light signaling. Plant Cell Physiol. 45: 968–975. Google Scholar
  200. B.M. Parks , and P.H. Quail (1991). Phytochrome-deficient hy1 and hy2 long hypocotyl mutants of Arabidopsis are defective in phytochrome chromophore biosynthesis. Plant Cell 3: 1177–1186. Google Scholar
  201. B.M. Parks , and P.H. Quail (1993). hy8, a new class of Arabidopsis long hypocotyl mutants deficient in functional phytochrome A. Plant Cell 5: 39–48. Google Scholar
  202. B.M. Parks , P.H. Quail , and R.P. Hangarter (1996). Phytochrome A regulates red-light induction of phototropic enhancement in Arabidopsis. Plant Physiol. 110: 155–162. Google Scholar
  203. B.M. Parks , and E.P. Spalding (1999). Sequential and coordinated action of phytochromes A and B during Arabidopsis stem growth revealed by kinetic analysis. Proc. Natl. Acad. Sci. USA 96: 14142–14146. Google Scholar
  204. U.V. Pedmale , R.B. Celaya , and E. Liscum (2010). Phototropism: Mechanism and outcomes. The Arabidopsis Book 8: e0125 doi: 0110.1199/tab.0125Google Scholar
  205. B.K. Phee , J.I. Kim , D.H. Shin , J. Yoo , K.J. Park , Y.J. Han , Y.K. Kwon , M.H. Cho , J.S. Jeon , S.H. Bhoo , and T.R. Hahn (2008). A novel protein phosphatase indirectly regulates phytochrome-interacting factor 3 via phytochrome. Biochem J. 415: 247–255. Google Scholar
  206. J. Putterill , F. Robson , K. Lee , R. Simon , and G. Coupland (1995). The CONSTANS gene of Arabidopsis promotes flowering and encodes a protein showing similarities to zinc finger transcription factors. Cell 80: 847–857. Google Scholar
  207. P.H. Quail (1997a). An emerging molecular map of the phytochromes. Plant Cell Environ. 20: 657–665. Google Scholar
  208. P.H. Quail (1997b). The phytochromes: a biochemical mechanism of signaling in sight? BioEssays 19: 571–579. Google Scholar
  209. P.H. Quail (2002). Phytochrome photosensory signalling networks. Nat. Rev. Mol. Cell Biol. 3: 85–93. Google Scholar
  210. P.H. Quail (2011). Phytochromes. Curr. Biol. 20: R504–507. Google Scholar
  211. P.H. Quail , M.T. Boylan , B.M. Parks , T.W. Short , Y. Xu , and D. Wagner (1995). Phytochromes: photosensory perception and signal transduction. Science 268: 675–680. Google Scholar
  212. J.W. Reed , A. Nagatani , T.D. Elich , M. Fagan , and J. Chory (1994). Phytochrome A and phytochrome B have overlapping but distinct functions in Arabidopsis development. Plant Physiol. 104: 1139–1149. Google Scholar
  213. J.W. Reed , P. Nagpal , R.M. Bastow , K.S. Solomon , M.J. Dowson-Day , R.P. Elumalai , and A.J. Millar (2000). Independent action of ELF3 and phyB to control hypocotyl elongation and flowering time. Plant Physiol. 122: 1149–1160. Google Scholar
  214. A. Remberg , A. Ruddat , S.E. Braslavsky , W. Gartner , and K. Schaffner (1998). Chromophore incorporation, Pr to Pfr kinetics, and Pfr thermal reversion of recombinant N-terminal fragments of phytochrome A and B chromoproteins. Biochemistry 37: 9983–9990. Google Scholar
  215. L. Rizzini , J.J. Favory , C. Cloix , D. Faggionato , A. O'Hara , E. Kaiserli , R. Baumeister , E. Schafer , F. Nagy , G.I. Jenkins , and R. Ulm (2011). Perception of UV-B by the Arabidopsis UVR8 protein. Science 332: 103–108. Google Scholar
  216. F. Robson , M.M. Costa , S.R. Hepworth , I. Vizir , M. Pineiro , P.H. Reeves , J. Putterill , and G. Coupland (2001). Functional importance of conserved domains in the flowering-time gene CONSTANS demonstrated by analysis of mutant alleles and transgenic plants. Plant J. 28: 619–631. Google Scholar
  217. N.C. Rockwell , Y.S. Su , and J.C. Lagarias (2006). Phytochrome structure and signaling mechanisms. Annu. Rev. Plant Biol. 57: 837–858. Google Scholar
  218. J. Rosler , I. Klein , and M. Zeidler (2007). Arabidopsis fhl/fhy1 double mutant reveals a distinct cytoplasmic action of phytochrome A. Proc. Natl. Acad. Sci. USA 104: 10737–10742. Google Scholar
  219. S.J. Roux (1994). Signal transduction in phytochrome responses. In Photomorphogenesis in plants, R.E. Kendrick , and G.H.M. Kronenberg , eds, (Dordrecht, The Netherlands: Kluwer Academic Publishers), pp. 187–209. Google Scholar
  220. J.S. Ryu , J.I. Kim , T. Kunkel , B.C. Kim , D.S. Cho , S.H. Hong , S.H. Kim , A.P. Fernandez , Y. Kim , J.M. Alonso , J.R. Ecker , F. Nagy , P.O. Lim , P.S. Song , E. Schafer , and H.G. Nam (2005). Phytochrome-specific type 5 phosphatase controls light signal flux by enhancing phytochrome stability and affinity for a signal transducer. Cell 120: 395–406. Google Scholar
  221. Y. Saijo , J.A. Sullivan , H. Wang , J. Yang , Y. Shen , V. Rubio , L. Ma , U. Hoecker , and X.W. Deng (2003). The COP1-SPA1 interaction defines a critical step in phytochrome A-mediated regulation of HY5 activity. Genes Dev. 17: 2642–2647. Google Scholar
  222. Y. Saijo , D. Zhu , J. Li , V. Rubio , Z. Zhou , Y. Shen , U. Hoecker , H. Wang , and X.W. Deng (2008). Arabidopsis COP1/SPA1 complex and FHY1/FHY3 associate with distinct phosphorylated forms of phytochrome A in balancing light signaling. Mol. Cell 31: 607–613. Google Scholar
  223. K. Sakamoto , and A. Nagatani (1996). Nuclear localization activity of phytochrome B. Plant J. 10: 859–868. Google Scholar
  224. M.G. Salter , K.A. Franklin , and G.C. Whitelam (2003). Gating of the rapid shade-avoidance response by the circadian clock in plants. Nature 426: 680–683. Google Scholar
  225. T.F. Schultz , T. Kiyosue , M. Yanovsky , M. Wada , and S.A. Kay (2001). A role for LKP2 in the circadian clock of Arabidopsis. Plant Cell 13: 2659–2670. Google Scholar
  226. I. Searle , and G. Coupland (2004). Induction of flowering by seasonal changes in photoperiod. EMBO J. 23: 1217–1222. Google Scholar
  227. H.S. Seo , E. Watanabe , S. Tokutomi , A. Nagatani , and N.H. Chua (2004). Photoreceptor ubiquitination by COP1 E3 ligase desensitizes phytochrome A signaling. Genes Dev. 18: 617–622. Google Scholar
  228. H.S. Seo , J.Y. Yang , M. Ishikawa , C. Bolle , M.L. Ballesteros , and N.H. Chua (2003). LAF1 ubiquitination by COP1 controls photomorphogenesis and is stimulated by SPA1. Nature 423: 995–999. Google Scholar
  229. G. Serino , and X.W. Deng (2003). The COP9 signalosome: regulating plant development through the control of proteolysis. Annu. Rev. Plant Biol. 54: 165–182. Google Scholar
  230. G. Sessa , M. Carabelli , M. Sassi , A. Cioifi , M. Possenti , F. Mittempergher , J. Becker , G. Morelli , and I. Ruberti (2005). A dynamic balance between gene activation and repression regulates the shade avoidance response in Arabidopsis. Genes Dev. 19: 2811–2815. Google Scholar
  231. J. Shanklin , M. Jabben , and R.D. Vierstra (1987). Red light-induced formation of ubiquitin-phytochrorne conjugates: identification of possible intermediates of phytochrome degradation. Proc. Natl. Acad. Sci. USA 84: 359–363. Google Scholar
  232. R.A. Sharrock , and T. Clack (2002). Patterns of expression and normalized levels of the five Arabidopsis phytochromes. Plant Physiol. 130: 442–456. Google Scholar
  233. R.A. Sharrock , and P.H. Quail (1989). Novel phytochrome sequences in Arabidopsis thaliana: structure, evolution, and differential expression of a plant regulatory photoreceptor family. Genes Dev. 3: 1745–1757. Google Scholar
  234. H. Shen , J. Moon , and E. Huq (2005a). PIF1 is regulated by light-mediated degradation through the ubiquitin-26S proteasome pathway to optimize photomorphogenesis of seedlings in Arabidopsis. Plant J. 44: 1023–1035. Google Scholar
  235. H. Shen , L. Zhu , A. Castillon , M. Majee , B. Downie , and E. Huq (2008). Light-induced phosphorylation and degradation of the negative regulator PHYTOCHROME-INTERACTING FACTOR 1 from Arabidopsis depend upon its direct physical interactions with photoactivated phytochromes. Plant Cell 20: 1586–1602. Google Scholar
  236. Y. Shen , S. Feng , L. Ma , R. Lin , L.J. Qu , Z. Chen , H. Wang , and X.W. Deng (2005b). Arabidopsis FHY1 protein stability is regulated by light via phytochrome A and 26S proteasome. Plant Physiol. 139: 1234–1243. Google Scholar
  237. Y. Shen , R. Khanna , C.M. Carle , and P.H. Quail (2007). Phytochrome induces rapid PIF5 phosphorylation and degradation in response to red-light activation. Plant Physiol. 145: 1043–1051. Google Scholar
  238. Y. Shen , Z. Zhou , S. Feng , J. Li , A. Tan-Wilson , L.J. Qu , H. Wang , and X.W. Deng (2009). Phytochrome A mediates rapid red light-induced phosphorylation of Arabidopsis FAR-RED ELONGATED HYPOCOTYL1 in a low fluence response. Plant Cell 21: 494–506. Google Scholar
  239. J. Shin , K. Kim , H. Kang , I.S. Zulfugarov , G. Bae , C.H. Lee , D. Lee , and G. Choi (2009). Phytochromes promote seedling light responses by inhibiting four negatively-acting phytochrome-interacting factors. Proc. Natl. Acad. Sci. USA 106: 7660–7665. Google Scholar
  240. T. Shinomura , A. Nagatani , H. Hanzawa , M. Kubota , M. Watanabe , and M. Furuya (1996). Action spectra for phytochrome A- and B-specific photoinduction of seed germination in Arabidopsis thaliana. Proc. Nail. Acad. Sci. USA 93: 8129–8133. Google Scholar
  241. H. Smith , and G.C. Whitelam (1997). The shade avoidance syndrome: multiple responses mediated by multiple phytochromes. Plant Cell Environ. 20: 840–844. Google Scholar
  242. H. Smith , Y. Xu , and P.H. Quail (1997). Antagonistic but complementary actions of phytochromes A and B allow seedling de-etiolation. Plant Physiol. 114: 637–641. Google Scholar
  243. M.S. Soh , Y.M. Kim , S.J. Han , and P.S. Song (2000). REP1, a basic helix-loop-helix protein, is required for a branch pathway of phytochrome A signaling in Arabidopsis. Plant Cell 12: 2061–2074. Google Scholar
  244. D.E. Somers , P.F. Devlin , and S.A. Kay (1998). Phytochromes and cryptochromes in the entrainment of the Arabidopsis circadian clock. Science 282: 1488–1490. Google Scholar
  245. D.E. Somers , W.Y. Kim , and R. Geng (2004). The F-box protein ZEITLUPE confers dosage-dependent control on the circadian clock, photomorphogenesis, and flowering time. Plant Cell 16: 769–782. Google Scholar
  246. D.E. Somers , T.F. Schultz , M. Milnamow , and S.A. Kay (2000). ZEIT-LUPE encodes a novel clock-associated PAS protein from Arabidopsis. Cell 101: 319–329. Google Scholar
  247. D.E. Somers , R.A. Sharrock , J.M. Tepperman , and P.H. Quail (1991). The hy3 long hypocotyl mutant of Arabidopsis is deficient in phytochrome E. Plant Cell 3: 1263–1274. Google Scholar
  248. V. Speth , V. Otto , and E. Schafer (1986). Intracellular localization of phytochrome in oat coleoptiles by electron microscopy. Planta 168: 299–304. Google Scholar
  249. J. Stockhaus , A. Nagatani , U. Halfter , S. Kay , M. Furuya , and N.H. Chua (1992). Serine-to-alanine substitutions at the amino-terminal region of phytochrome A result in an increase in biological activity. Genes Dev. 6: 2364–2372. Google Scholar
  250. B. Strasser , M. Sanchez-Lamas , M.J. Yanovsky , J.J. Casai , and P.D. Cerdan (2010). Arabidopsis thaliana life without phytochromes. Proc. Natl. Acad. Sci. USA 107: 4776–4781. Google Scholar
  251. P. Suarez-Lopez , K. Wheatley , F. Robson , H. Onouchi , F. Valverde , and G. Coupland (2001). CONSTANS mediates between the circadian clock and the control of flowering in Arabidopsis. Nature 410: 1116–1120. Google Scholar
  252. J.A. Sullivan , K. Shirasu , and X.W. Deng (2003). The diverse roles of ubiquitin and the 26S proteasome in the life of plants. Nat. Rev. Genet. 4: 948–958. Google Scholar
  253. S. Takagi , S.G. Kong , Y. Mineyuki , and M. Furuya (2003). Regulation of actin-dependent cytoplasmic motility by type II phytochrome occurs within seconds in Vallisneria gigantea epidermal cells. Plant Cell 15: 331–345. Google Scholar
  254. J.M. Tepperman , M.E. Hudson , R. Khanna , T. Zhu , S.H. Chang , X. Wang , and P.H. Quail (2004). Expression profiling of phyB mutant demonstrates substantial contribution of other phytochromes to redlight-regulated gene expression during seedling de-etiolation. Plant J. 38: 725–739. Google Scholar
  255. J.M. Tepperman , Y.S. Hwang , and P.H. Quail (2006). phyA dominates in transduction of red-light signals to rapidly responding genes at the initiation of Arabidopsis seedling de-etiolation. Plant J. 48: 728–742. Google Scholar
  256. J.M. Tepperman , T. Zhu , H.S. Chang , X. Wang , and P.H. Quail (2001). Multiple transcription-factor genes are early targets of phytochrome A signaling. Proc. Natl. Acad. Sci. USA 98: 9437–9442. Google Scholar
  257. M.J. Terry (1997). Phytochrome chromophore-deficient mutants. Plant Cell Environ. 20: 740–745. Google Scholar
  258. B. Thines , and F.G. Harmon (2010). Ambient temperature response establishes ELF3 as a required component of the core Arabidopsis circadian clock. Proc. Natl. Acad. Sci. USA 107: 3257–3262. Google Scholar
  259. G. Toledo-Ortiz , E. Huq , and P.H. Quail (2003). The Arabidopsis basic/ helix-loop-helix transcription factor family. Plant Cell 15: 1749–1770. Google Scholar
  260. T.S. Tseng , P.A. Salome , C.R. McClung , and N.E. Olszewski (2004). SPINDLY and GIGANTEA interact and act in Arabidopsis thaliana pathways involved in light responses, flowering, and rhythms in cotyledon movements. Plant Cell 16: 1550–1563. Google Scholar
  261. A.T. Ulijasz , G. Cornilescu , C.C. Cornilescu , J. Zhang , M. Rivera , J.L. Markley , and R.D. Vierstra (2010). Structural basis for the photoconversion of a phytochrome to the activated Pfr form. Nature 463: 250–254. Google Scholar
  262. H. Ullah , J.G. Chen , J.C. Young , K.H. Im , M.R. Sussman , and A.M. Jones (2001). Modulation of cell proliferation by heterotrimeric G protein in Arabidopsis. Science 292: 2066–2069. Google Scholar
  263. R. Ulm , A. Baumann , A. Oravecz , Z. Mate , E. Adam , E.J. Oakeley , E. Schafer , and F. Nagy (2004). Genome-wide analysis of gene expression reveals function of the bZIP transcription factor HY5 in the UV-B response of Arabidopsis. Proc. Natl. Acad. Sci. USA 101: 1397–1402. Google Scholar
  264. F. Valverde , A. Mouradov , W. Soppe , D. Ravenscroft , A. Samach , and G. Coupland (2004). Photoreceptor regulation of CONSTANS protein in photoperiodic flowering. Science 303: 1003–1006. Google Scholar
  265. R.D. Vierstra , and S.J. Davis (2000). Bacieriophytochromes: new tools for understanding phytochrome signal transduction. Semin. Cell Dev. Biol. 11: 511–521. Google Scholar
  266. D. Vince-Prue (1994). The duration of light and photoperiodic responses. In Photomorphogenesis in plants, R.E. Kendrick , and G.H.M. Kronenberg , eds, (Dordrecht, The Netherlands: Kluwer Academic Publishers), pp. 447–490. Google Scholar
  267. D. Wagner , C.D. Fairchild , R.M. Kuhn , and P.H. Quail (1996). Chromophore-bearing NH2-terminal domains of phytochromes A and B determine their photosensory specificity and differential light lability. Proc. Natl. Acad. Sci. USA 93: 4011–4015. Google Scholar
  268. H. Wang , and X.W. Deng (2002). Arabidopsis FHY3 defines a key phytochrome A signaling component directly interacting with its homologous partner FAR1. EMBO J. 21: 1339–1349. Google Scholar
  269. H. Wang , and X.W. Deng (2003). Dissecting the phytochrome A-dependent signaling network in higher plants. Trends Plant Sci. 8: 172–178. Google Scholar
  270. H. Wang , L. Ma , J. Habashi , J. Li , H. Zhao , and X.W. Deng (2002). Analysis of far-red light-regulated genome expression profiles of phytochrome A pathway mutants in Arabidopsis. Plant J. 32: 723–733. Google Scholar
  271. B. Wenden , L. Kozma-Bognar , K.D. Edwards , A.J. Hall , J.C. Locke , and A.J. Millar (2011). Light inputs shape the Arabidopsis circadian system. Plant J. 66: 480–491. Google Scholar
  272. G.C. Whitelam , E. Johnson , J. Peng , P. Carol , M.L. Anderson , J.S. Cowl , and N.P. Harberd (1993). Phytochrome A null mutants of Arabidopsis display a wild-type phenotype in white light. Plant Cell 5: 757–768. Google Scholar
  273. Y.S. Wong , H.C. Cheng , D.A. Walsh , and J.C. Lagarias (1986). Phosphorylation of Avena phytochrome in vitro as a probe of light-induced conformational changes. J. Biol. Chem. 261: 12089–12097. Google Scholar
  274. Y.S. Wong , R.W. McMichael , and J.C. Lagarias (1989). Properties of a polycation-stimulated protein kinase associated with purified Avena phytochrome. Plant Physiol. 91: 709–718. Google Scholar
  275. S.H. Wu , and J.C. Lagarias (2000). Defining the bilin lyase domain: lessons from the extended phytochrome superfamily. Biochemistry 39: 13487–13495. Google Scholar
  276. Y. Yanagawa , J.A. Sullivan , S. Komatsu , G. Gusmaroli , G. Suzuki , J. Yin , T. Ishibashi , Y. Saijo , V. Rubio , S. Kimura , J. Wang , and X.W. Deng (2004). Arabidopsis COP10 forms a complex with DDB1 and DET1 in vivo and enhances the activity of ubiquitin conjugating enzymes. Genes Dev. 18: 2172–2181. Google Scholar
  277. J. Yang , R. Lin , J. Sullivan , U. Hoecker , B. Liu , L. Xu , X.W. Deng , and H. Wang (2005). Light regulates COP1-mediated degradation of HFR1, a transcription factor essential for light signaling in Arabidopsis. Plant Cell 17: 804–821. Google Scholar
  278. K.Y. Yang , Y.M. Kim , S. Lee , P.S. Song , and M.S. Soh (2003). Overexpression of a mutant basic helix-loop-helix protein HFR1, HFR1-deltaN105, activates a branch pathway of light signaling in Arabidopsis. Plant Physiol. 133: 1630–1642. Google Scholar
  279. S.W. Yang , I.C. Jang , R. Henriques , and N.H. Chua (2009). FAR-RED ELONGATED HYPOCOTYL1 and FHY1-LIKE associate with the Arabidopsis transcription factors LAF1 and HFR1 to transmit phytochrome A signals for inhibition of hypocotyl elongation. Plant Cell 21: 1341–1359. Google Scholar
  280. M.J. Yanovsky , M. Izaguirre , J.A. Wagmaister , C. Gatz , S.D. Jackson , B. Thomas , and J.J. Casal (2000). Phytochrome A resets the circadian clock and delays tuber formation under long days in potato. Plant J. 23: 223–232. Google Scholar
  281. M.J. Yanovsky , and S.A. Kay (2002). Molecular basis of seasonal time measurement in Arabidopsis. Nature 419: 308–312. Google Scholar
  282. M.J. Yanovsky , M.A. Mazzella , G.C. Whitelam , and J.J. Casal (2001). Resetting of the circadian clock by phytochromes and cryptochromes in Arabidopsis. J. Biol. Rhythms 16: 523–530. Google Scholar
  283. K.C. Yeh , and J.C. Lagarias (1998). Eukaryotic phytochromes: light-regulated serine/threonine protein kinases with histidine kinase ancestry. Proc. Natl. Acad. Sci. USA 95: 13976–13981. Google Scholar
  284. K.C. Yeh , S.H. Wu , J.T. Murphy , and J.C. Lagarias (1997). A cyanobacterial phytochrome two-component light sensory system. Science 277: 1505–1508. Google Scholar
  285. C. Yi , and X.W. Deng (2005). COP1 -from plant photomorphogenesis to mammalian tumorigenesis. Trends Cell Biol. 15: 618–625. Google Scholar
  286. J.W. Yu , V. Rubio , N.Y. Lee , S. Bai , S.Y. Lee , S.S. Kim , L. Liu , Y. Zhang , M.L. Irigoyen , J.A. Sullivan , I. Lee , Q. Xie , N.C. Paek , and X.W. Deng (2008). COP1 and ELF3 control circadian function and photoperiodic flowering by regulating Gl stability. Mol. Cell 32: 617–630. Google Scholar
  287. X. Yu , H. Liu , J. Klejnot , and C. Lin (2010). The cryptochrome blue light receptors. The Arabidopsis Book 8: e0135. doi: 0110.1199/tab.0135Google Scholar
  288. M.T. Zagotta , K.A. Hicks , C.I. Jacobs , J.C. Young , R.P. Hangarter , and D.R. Meeks-Wagner (1996). The Arabidopsis ELF3 gene regulates vegetative photomorphogenesis and the photoperiodic induction of flowering. Plant J. 10: 691–702. Google Scholar
  289. M. Zeidler , Q. Zhou , X. Sarda , C.P. Yau , and N.H. Chua (2004). The nuclear localization signal and the C-terminal region of FHY1 are required for transmission of phytochrome A signals. Plant J. 40: 355–365. Google Scholar
  290. H. Zhang , H. He , X. Wang , X. Yang , L. Li , and X.W. Deng (2011). Genome-wide mapping of the HY5-mediated gene networks in Arabidopsis that involve both transcriptional and post-transcriptional regulation. Plant J. 65: 346–358. Google Scholar
  291. Q. Zhou , P.D. Hare , S.W. Yang , M. Zeidler , L.F. Huang , and N.H. Chua (2005). FHL is required for full phytochrome A signaling and shares overlapping functions with FHY1. Plant J. 43: 356–370. Google Scholar
  292. Y.C. Zhou , M. Dieterle , C. Buche , and T. Kreisch (2002). The negatively acting factors EID1 and SPA1 have distinct functions in phytochrome A-specific light signaling. Plant Physiol. 128: 1098–1108. Google Scholar
  293. D. Zhu , A. Maier , J.H. Lee , S. Laubinger , Y. Saijo , H. Wang , L.J. Qu , U. Hoecker , and X.W. Deng (2008). Biochemical characterization of Arabidopsis complexes containing CONSTITUTIVELY PHOTOMORPHOGENIC1 and SUPPRESSOR OF PHYA proteins in light control of plant development. Plant Cell 20: 2307–2323. Google Scholar
© 2011 American Society of Plant Biologists
Jigang Li, Gang Li, Haiyang Wang and Xing Wang Deng "Phytochrome Signaling Mechanisms," The Arabidopsis Book 2011(9), (1 August 2011). https://doi.org/10.1199/tab.0148
JOURNAL ARTICLE
PAGES


SHARE
ARTICLE IMPACT
Back to Top