Other components that function within or close to the circadian oscillator have recently been identified: FIONA 1, TIME FOR COFFEE, LIGHT REGULATED WD-1 (LWD1) and LWD2 (Ding *et al.*, 2007; Kim *et al.*, 2008; Wu *et al.*, 2008). However, it is not known whether these clock proteins are part of pre-existing loops or constitute unidentified regulatory loops. It has been recently reported that LWD1/2 regulate the expression of multiple oscillator genes and attenuate light signals to adjust period length. Further, it was also proposed that LWD1 and PRR9 form a positive feedback loop within the central oscillator which is also involved in regulating the light input pathway (Wang *et al.*, 2011) (Figure 2). These results underscore the difficulties in dissecting which signalling events are part of the circadian oscillator and which ones are input pathways.

#### **4.2 Light signalling input to the circadian clock**

Several different photoreceptors mediate light input to the clock, including the phytochromes and the cryptochromes (Somers *et al.*, 1998; Devlin & Kay, 2000; Yanovsky *et al.*, 2001). However, the molecular mechanisms are only partially known. The ztl family of photoreceptors interacts with clock components and regulates their turnover; hence they are potentially part of input mechanisms. ztl interacts with TOC1 and PRR5, leading to their degradation via the proteasome pathway in the dark (Kiba *et al.*, 2007; Mas *et al.*, 2003). The TOC1-ztl interaction does not depend on light, but an interaction between ztl and GI is bluelight dependent, stabilizes both ztl and GI, and contributes to the robust rhythms of TOC1

the Myb-related transcription factors CIRCADIAN CLOCK ASSOCIATED 1 (CCA1) and LATE ELONGATED HYPOCOTYL (LHY) and the pseudo-response regulator TIMING OF CAB EXPRESSION 1 (TOC1/PRR1) (Loop 1, figure 2). CCA1 and LHY proteins have partially redundant functions, bind directly to the TOC1 promoter and inhibit its expression during the day (Alabadi *et al.*, 2001). In turn, TOC1 promotes the expression of *CCA1* and *LHY* indirectly via a hypothetical component X in the morning. The mechanism by which TOC1 induces *CCA1* and *LHY1* is not completely understood, but it includes CCA1 HIKING EXPEDITION (CHE), a TCP type transcription factor, which associates with TOC1 to regulate *CCA1* (Pruneda-Paz *et al.*, 2009). Eventually, CHE also forms an additional loop

Mathematical modelling suggests that an evening-phased negative loop is coupled to the first loop, with an unknown component Y that positively regulates *TOC1* whereas *Y* is negatively regulated by TOC1, CCA1 and LHY (Locke *et al.*, 2005) (Loop 2, figure 2). It was suggested that a portion of Y activity is provided by the protein GIGANTEA (GI) (Locke *et*

The Arabidopsis genome contains four genes encoding proteins with similarity to TOC1: PSEUDORESPONSE REGULATOR (PRR), PRR3, 5, 7 and 9. All these *PRR* genes play a role in the circadian system, although the effect of single mutations is subtle. Multiple mutants generally have stronger phenotypes, for example the triple *prr5 prr7 prr9* mutants are essentially arrhythmic (Nakamichi *et al.*, 2005a; Nakamichi *et al.*, 2005b). Experimental and modelling studies suggest that morning expression of *CCA1* and *LHY* activates the transcription of *PRR7* and *PRR9* (Farre *et al.*, 2005; Nakamichi *et al.*, 2005b; Zeilinger *et al.*, 2006). This loop is called morning loop (Loop 3, figure 2) and is closed when PRR7 and PRR9 feedback to inhibit *CCA1* and *LHY* expression. Together the three interlinked feedback loops form an important part of the clock regulatory mechanism and enhance the robustness

Other components that function within or close to the circadian oscillator have recently been identified: FIONA 1, TIME FOR COFFEE, LIGHT REGULATED WD-1 (LWD1) and LWD2 (Ding *et al.*, 2007; Kim *et al.*, 2008; Wu *et al.*, 2008). However, it is not known whether these clock proteins are part of pre-existing loops or constitute unidentified regulatory loops. It has been recently reported that LWD1/2 regulate the expression of multiple oscillator genes and attenuate light signals to adjust period length. Further, it was also proposed that LWD1 and PRR9 form a positive feedback loop within the central oscillator which is also involved in regulating the light input pathway (Wang *et al.*, 2011) (Figure 2). These results underscore the difficulties in dissecting which signalling events are part of the circadian oscillator and

Several different photoreceptors mediate light input to the clock, including the phytochromes and the cryptochromes (Somers *et al.*, 1998; Devlin & Kay, 2000; Yanovsky *et al.*, 2001). However, the molecular mechanisms are only partially known. The ztl family of photoreceptors interacts with clock components and regulates their turnover; hence they are potentially part of input mechanisms. ztl interacts with TOC1 and PRR5, leading to their degradation via the proteasome pathway in the dark (Kiba *et al.*, 2007; Mas *et al.*, 2003). The TOC1-ztl interaction does not depend on light, but an interaction between ztl and GI is bluelight dependent, stabilizes both ztl and GI, and contributes to the robust rhythms of TOC1

*al.*, 2005), but this is still unclear (Ito *et al.*, 2009; Martin-Tryon *et al.*, 2007).

of the network against environmental perturbations (Harmer, 2009).

with CCA1 (Imaizumi, 2010).

which ones are input pathways.

**4.2 Light signalling input to the circadian clock**

(Kim *et al.*, 2007), contributing to a faster degradation of ztl, GI, TOC1 and PRR5 in darkness than in light (Loop 4, figure 2) (Kiba *et al.*, 2007; Kim *et al.*, 2007; Mas *et al.*, 2003). Within this loop 4, TOC1 binds to PRR3, interfering with TOC1 binding to ztl (Para *et al.*, 2007). Thus, PRR3 seems to stabilize TOC1 avoiding its recruitment to the SCF complex and its degradation by the proteasome (Loop 4, figure 2).

Fig. 2. A model for the *Arabidopsis* clock. The circadian clock is composed of several interlocking positive and negative loops.

The other members of the ztl family, fkf1 and lkp2, were also studied. Mutant combinations showed that fkf1 and lkp2 play similar roles to ztl in the circadian clock when ztl is absent, and that both of them interact with TOC1 and PRR5. These results indicate that ztl, fkf1 and lkp2 regulate TOC1 and PRR5 degradation and are important to determine the period of circadian oscillation (Baudry *et al.*, 2010).

Cryptochromes also signal to the circadian clock. However, the mechanisms are still unclear. One possibility is through the regulation of COP1 activity; COP1 directly interacts with ELF3 and with GI to promote GI degradation by the proteasome. This could be a mechanism by which cryptochromes regulate the activity of GI, a protein closely associated with circadian clock function (Yu *et al.*, 2008).

As mentioned above, one interesting aspect of phytochrome and circadian clock is that mutants affected in the clock are also affected in phytochrome responses (Ito *et al.*, 2007). However, how phytochromes contribute to the entrainment of the clock is still unclear. It was previously suggested that PIF3 could directly induce *CCA1* and *LHY* mRNA expression (Martinez-Garcia *et al.*, 2000). Later, it was shown that TOC1 interacts with PIF3 and PIL1

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 203

Genome-wide CHIP-chip analysis was used to identify HY5 binding regions and to compare this information to HY5-global expression data. This approach allowed the identification of more that 1100 direct targets where HY5 can either activate or repress transcription. However, not all the targets were light responsive genes, suggesting that HY5 must act in concert with other factors to confer light responsiveness (Zhang *et al.*,

**5.3 The dissection of single light responsive promoters reveals another layer of**

paradigm of light-regulated gene expression (Akhilesh & Gaur, 2003).

different light inputs, as will be explained below.

Most of the photoreceptors, signalling components and transcription factors mentioned above were identified using genetic approaches, after Arabidopsis was established as the model plant. Another strategy to understand light signalling and photosynthetic gene expression has been underway since late mid 80s, after the first transgenic plants became available. This strategy was simple, the generation of transgenic plants bearing promoter:reporter gene fusions. With this approach, light responsive promoters were the subject of extensive research with the aim of finding the light responsive elements (LREs) and their cognate binding factors. The genes encoding the small subunits of the Rubisco (RbcS) and the light-harvesting chlorophyll a/b-binding proteins (Lhc; previously known as Cab), were considered a

Several LREs were described, as GT-1-Boxes (core sequence GGTTAA), I-Boxes (GATAA), G-Boxes (CACGTG), H-Boxes (CCTACC), AT-rich sequences (consensus AATATTTTTATT) (Akhilesh & Gaur, 2003). Using complementary approaches as Gel Shift analysis and DNA footprinting, some of the cognate binding factors were identified. However, three difficulties hampered this approaches. First, the LREs identified were not always enough to sustain light regulation. Hence, it was proposed that combinations of different motifs but not multimerisation of single motifs could function as LREs, confirming the complex nature of these regulatory elements (Chattopadhyay *et al.*, 1998; Puente *et al.*, 1996). Second, when the cognate transcription factors were studied in Arabidopsis with available mutants, a direct role in light signal was not evident. This can be illustrated by the GT-element binding factors, a small family of plant trihelix DNAbinding proteins comprising Arabidopsis GT2 (AT1G76890), DF1L (AT1G76880), PTL (At5g03680), GT-2-LIKE1 (GTL1, AT1G33240), GT2L (At5g28300), EDA31 (AT3G10000) and GTL1L (AT5G47660). Some of these transcription factors have roles in the fusion of the polar nuclei, in the development of the embryo sac or even perianth development (Brewer *et al.*, 2004; Pagnussat *et al.*, 2005), but were not involved in responses to light. The third difficulty was the apparent "redundancy" of LREs in single promoters. This redundancy could be just the consequence of a single promoter responding to several

In a few examples, thorough analysis of promoter sequences, combined with genetic approaches significantly advanced our understanding of light-regulated transcription, but also revealed the complex nature underneath this process. The Arabidopsis *Lhcb1\*1* (*Cab 2*) promoter fused to luciferase reporters has been extensively used as a marker for light and circadian expression. Genetic screenings using this construct led to the isolation of *toc1* mutants (Strayer *et al.*, 2000). Promoter analysis of *Lhcb1\*1* allowed the identification of a 78 bp fragment that was sufficient to confer phytochrome and circadian regulation to a minimal promoter (Anderson *et al.*, 1994). Further analysis of this promoter allowed the

2011).

**complexity**

(Yamashino *et al.*, 2003). However, thorough analysis of PIF3 function has led to the conclusion that it does not play a significant role in controlling light input to the circadian clock (Viczian *et al.*, 2005).

Indeed, there is circumstantial evidence of phytochromes regulating CCA1 and LHY. Both genes are rapidly induced in a TOC1 dependent manner upon transfer of dark grown seedlings to red light. This induction requires *EARLY FLOWERING 4* (*ELF4*), which forms with CCA1 and LHY a negative feedback loop in an analogous manner to TOC1 (Kikis *et al.*, 2005) and ELF4 is itself a direct target of FHY3, FAR1 and HY5 (Li *et al.*, 2011). *ELF3*, is also necessary for light-induced expression of *CCA1* and *LHY* and this event seems to occur indirectly, through a direct repression of PRR9 by physically interacting with its promoter (Dixon *et al.*, 2011).

#### **5. Downstream targets of light and clock signalling**

#### **5.1 The impact of the circadian clock in the expression of photosynthesis related genes**

As presented above, the interconnections between the clock and light signalling are extremely complex. The regulation of outputs is not an exception. One unbiased measure of the impact of the circadian clock on plant development is the finding that at least one third of the Arabidopsis genome is circadian regulated (Covington *et al.*, 2008). The genes involved in photosynthesis are an important target group of the circadian clock, and tend to be expressed at the middle of the subjective day, together with genes involved in the phenylpropanoid pathway (Edwards *et al.*, 2006). In another global analysis it was shown that PRR5, PRR7 and PRR9 are negative regulators of the chlorophyll and carotenoid biosynthetic pathways (Fukushima *et al.*, 2009).

Despite what we know of the clock impact on photosynthetic gene expression, the mechanisms are still poorly understood. One such mechanism may involve CCA1. CCA1 was originally identified by its binding to an AA(CA)AATCT motif in the *lhcb1\*3* promoter, and also shown to be required for phytochrome responsivity (Wang *et al.*, 1997). Hence, CCA1 can represent one of the mechanisms by which the clock regulates photosynthetic gene expression. Nevertheless, the reality is more complex. CCA1 binding site is similar to the Evening Element (AAAATATCT) found in promoters of clock regulated genes that peak toward the end of the subjective day (Harmer *et al.*, 2000), including TOC1, which is repressed by CCA1 (Alabadi *et al.*, 2001). However, *lhcb1\*3* expression peaks earlier and is promoted by CCA1 (Wang *et al.*, 1997). These apparent contradictions can be reconciled by the finding that CCA1 effects depend on the context, showing also another level of complexity (Harmer & Kay, 2005).

#### **5.2 Global expression analysis identifies the targets of photomorphogenesis master regulators**

HY5, the bZIP targeted by COP1 for degradation, is necessary for responses to a broad spectrum of wavelengths of light and, as explained above, acts as a positive regulator in photomorphogenesis. Arabidopsis plants defective in HY5 show aberrant light mediated phenotypes, including an elongated hypocotyl, reduced chlorophyll/anthocyanin accumulation and reduced chloroplast development in greening hypocotyls (Lee *et al.*, 2007). HY5 regulates the transcription of multiple genes in response to light signals through binding to G-box elements in their promoters such as RBCS1A or CHS1 genes.

(Yamashino *et al.*, 2003). However, thorough analysis of PIF3 function has led to the conclusion that it does not play a significant role in controlling light input to the circadian

Indeed, there is circumstantial evidence of phytochromes regulating CCA1 and LHY. Both genes are rapidly induced in a TOC1 dependent manner upon transfer of dark grown seedlings to red light. This induction requires *EARLY FLOWERING 4* (*ELF4*), which forms with CCA1 and LHY a negative feedback loop in an analogous manner to TOC1 (Kikis *et al.*, 2005) and ELF4 is itself a direct target of FHY3, FAR1 and HY5 (Li *et al.*, 2011). *ELF3*, is also necessary for light-induced expression of *CCA1* and *LHY* and this event seems to occur indirectly, through a direct repression of PRR9 by physically interacting with its promoter

**5.1 The impact of the circadian clock in the expression of photosynthesis related**

As presented above, the interconnections between the clock and light signalling are extremely complex. The regulation of outputs is not an exception. One unbiased measure of the impact of the circadian clock on plant development is the finding that at least one third of the Arabidopsis genome is circadian regulated (Covington *et al.*, 2008). The genes involved in photosynthesis are an important target group of the circadian clock, and tend to be expressed at the middle of the subjective day, together with genes involved in the phenylpropanoid pathway (Edwards *et al.*, 2006). In another global analysis it was shown that PRR5, PRR7 and PRR9 are negative regulators of the chlorophyll and carotenoid

Despite what we know of the clock impact on photosynthetic gene expression, the mechanisms are still poorly understood. One such mechanism may involve CCA1. CCA1 was originally identified by its binding to an AA(CA)AATCT motif in the *lhcb1\*3* promoter, and also shown to be required for phytochrome responsivity (Wang *et al.*, 1997). Hence, CCA1 can represent one of the mechanisms by which the clock regulates photosynthetic gene expression. Nevertheless, the reality is more complex. CCA1 binding site is similar to the Evening Element (AAAATATCT) found in promoters of clock regulated genes that peak toward the end of the subjective day (Harmer *et al.*, 2000), including TOC1, which is repressed by CCA1 (Alabadi *et al.*, 2001). However, *lhcb1\*3* expression peaks earlier and is promoted by CCA1 (Wang *et al.*, 1997). These apparent contradictions can be reconciled by the finding that CCA1 effects depend on the context, showing also another level of

**5.2 Global expression analysis identifies the targets of photomorphogenesis master**

HY5, the bZIP targeted by COP1 for degradation, is necessary for responses to a broad spectrum of wavelengths of light and, as explained above, acts as a positive regulator in photomorphogenesis. Arabidopsis plants defective in HY5 show aberrant light mediated phenotypes, including an elongated hypocotyl, reduced chlorophyll/anthocyanin accumulation and reduced chloroplast development in greening hypocotyls (Lee *et al.*, 2007). HY5 regulates the transcription of multiple genes in response to light signals through binding to G-box elements in their promoters such as RBCS1A or CHS1 genes.

**5. Downstream targets of light and clock signalling**

biosynthetic pathways (Fukushima *et al.*, 2009).

complexity (Harmer & Kay, 2005).

**regulators**

clock (Viczian *et al.*, 2005).

(Dixon *et al.*, 2011).

**genes**

Genome-wide CHIP-chip analysis was used to identify HY5 binding regions and to compare this information to HY5-global expression data. This approach allowed the identification of more that 1100 direct targets where HY5 can either activate or repress transcription. However, not all the targets were light responsive genes, suggesting that HY5 must act in concert with other factors to confer light responsiveness (Zhang *et al.*, 2011).

#### **5.3 The dissection of single light responsive promoters reveals another layer of complexity**

Most of the photoreceptors, signalling components and transcription factors mentioned above were identified using genetic approaches, after Arabidopsis was established as the model plant. Another strategy to understand light signalling and photosynthetic gene expression has been underway since late mid 80s, after the first transgenic plants became available. This strategy was simple, the generation of transgenic plants bearing promoter:reporter gene fusions. With this approach, light responsive promoters were the subject of extensive research with the aim of finding the light responsive elements (LREs) and their cognate binding factors. The genes encoding the small subunits of the Rubisco (RbcS) and the light-harvesting chlorophyll a/b-binding proteins (Lhc; previously known as Cab), were considered a paradigm of light-regulated gene expression (Akhilesh & Gaur, 2003).

Several LREs were described, as GT-1-Boxes (core sequence GGTTAA), I-Boxes (GATAA), G-Boxes (CACGTG), H-Boxes (CCTACC), AT-rich sequences (consensus AATATTTTTATT) (Akhilesh & Gaur, 2003). Using complementary approaches as Gel Shift analysis and DNA footprinting, some of the cognate binding factors were identified. However, three difficulties hampered this approaches. First, the LREs identified were not always enough to sustain light regulation. Hence, it was proposed that combinations of different motifs but not multimerisation of single motifs could function as LREs, confirming the complex nature of these regulatory elements (Chattopadhyay *et al.*, 1998; Puente *et al.*, 1996). Second, when the cognate transcription factors were studied in Arabidopsis with available mutants, a direct role in light signal was not evident. This can be illustrated by the GT-element binding factors, a small family of plant trihelix DNAbinding proteins comprising Arabidopsis GT2 (AT1G76890), DF1L (AT1G76880), PTL (At5g03680), GT-2-LIKE1 (GTL1, AT1G33240), GT2L (At5g28300), EDA31 (AT3G10000) and GTL1L (AT5G47660). Some of these transcription factors have roles in the fusion of the polar nuclei, in the development of the embryo sac or even perianth development (Brewer *et al.*, 2004; Pagnussat *et al.*, 2005), but were not involved in responses to light. The third difficulty was the apparent "redundancy" of LREs in single promoters. This redundancy could be just the consequence of a single promoter responding to several different light inputs, as will be explained below.

In a few examples, thorough analysis of promoter sequences, combined with genetic approaches significantly advanced our understanding of light-regulated transcription, but also revealed the complex nature underneath this process. The Arabidopsis *Lhcb1\*1* (*Cab 2*) promoter fused to luciferase reporters has been extensively used as a marker for light and circadian expression. Genetic screenings using this construct led to the isolation of *toc1* mutants (Strayer *et al.*, 2000). Promoter analysis of *Lhcb1\*1* allowed the identification of a 78 bp fragment that was sufficient to confer phytochrome and circadian regulation to a minimal promoter (Anderson *et al.*, 1994). Further analysis of this promoter allowed the

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 205

(Tanaka & Tanaka, 2007). Phytochrome and cryptochrome mutants contain lower levels of chlorophyll (Strasser *et al.*, 2010) stressing out the importance of the photomorphogenic signal for proper assembly of the photosynthetic machinery. In the next paragraphs we

review how light signalling pathways regulate chlorophyll biosynthesis (Figure 4).

Fig. 3. Light interactions in plastid development. Phytochrome and PIFs roles during the

2004; McCormac *et al.*, 2001; McCormac & Terry, 2002a; McCormac & Terry, 2002b).

biogenesis with circadian and light signalling (Stephenson *et al.*, 2009).

Glutamyl-tRNA reductase activity is regulated by negative feedback loops; the accumulation of Haeme, Mg-Protoporphyrin IX or Divinyl protochlorofilide a antagonise Glutamyl-tRNA reductase activity (Srivastava *et al.*, 2005). At the transcriptional level, *HEMA1* expression is induced by red and far-red light, implicating at least phyA and phyB, and blue light perceived by cry1 (McCormac *et al.*, 2001; McCormac & Terry, 2002a). *pif1* and *pif3* mutants contain higher levels of *HEMA1* mRNA, higher levels of protochlorophyllide and partially developed chloroplasts in the dark, a phenotype observed in *cop* mutants. The effects of *pif1* and *pif3* mutations are essentially additive, suggesting a model where phytochromes promote chloroplast biogenesis by antagonizing the activity of at least PIF1 and PIF3. As PIF1 and PIF3 are regulated by the circadian clock, but do not seem to affect central clock components (TOC1, CCA1, LHY), these PIFs seem to integrate chloroplast

The expression of photosynthetic nuclear genes is repressed by plastid signals if chloroplast biogenesis is blocked (retrograde signalling). This finding led to the isolation of mutants that disrupt chloroplast to nucleus communication, the genomes uncoupled mutants (*gun*) (Nott *et al.*, 2006). These mutants show high levels of *lhcb1* mRNA in the presence of norfluorzazon and were named *gun1* to *gun5*. *gun2* and *gun3* are allelic to *hy1* and *hy2* and disrupt phytochromobilin synthesis, leading to haeme accumulation and feedback

Chlorophyll synthesis occurs in plastids; in the first step glutamate is activated to GlutamyltRNA by the Glutamyl-tRNA synthetase, a step shared with plastid protein synthesis. The following step, the reduction of the Glutamyl-tRNA to produce glutamate-1-semialdehyde is subjected to tight regulation (Figure 4). In Arabidopsis, the Glutamyl-tRNA reductases are encoded by a little family of nuclear genes called *HEMA*. Of this family, the expression of *HEMA1* correlates with the expression of *Lhcb1* genes, which encode light-harvesting proteins of the photosystem II; in some way the expression of *HEMA1* reflects the demand of chlorophyll synthesis. On the other hand, *HEMA2* is not light regulated (Matsumoto *et al.*,

transition from proplastid or amyloplast to chloroplast.

identification of HY5, CCA1 and a DET1 responsive elements (Maxwell *et al.*, 2003). Similarly, it has been shown that HY5 binds to the *Lhcb1\*3* promoter and physically interacts with CCA1 to synergistically regulate expression (Andronis *et al.*, 2008).

Another promoter analysed in more detail was the tobacco *Lhcb1\*2*. First, a 146 bp promoter fragment sufficed to confer VLFR (mediated by phyA), LFR (mediated by phyB) and HIR (mediated by phyA) to a minimal promoter (Cerdan *et al.*, 1997). Then, the motifs for VLFR and LFR were dissected from the HIR responsive motifs (Cerdan *et al.*, 2000) and finally, the TGGA motif was shown to bind Bell-like homeodomain 1 (BLH1) as part of the phyA mediated HIR (Staneloni *et al.*, 2009). This promoter is an example of how several different photoreceptors can regulate a single gene and integrate their signalling pathways at the promoter level; at least four different photoreceptors were shown to regulate this single promoter (Casal *et al.*, 1998; Cerdan *et al.*, 1999; Mazzella *et al.*, 2001) .

#### **6. Light promotes chloroplast development**

Proplastids are found in the embryo; they are undifferentiated plastids that are converted to other kind of plastids like chromoplasts, amyloplasts, chloroplasts and etioplasts. During skotomorphogenic development, proplastids turn into etioplasts, the chloroplast precursors. Etioplasts contain the prolamellar body, a structure rich in protochlorophyllide, the chlorophyll precursor, and the enzyme protochlorophyllide oxidoreductase (POR). During the development of etioplasts into chloroplasts, the POR is directly activated by light to convert protochlorophyllide into divinyl-chlorophyllide a, which is chlorophyll a and b precursor (Tanaka & Tanaka, 2007). This light-dependent step can be promoted by red-light in Arabidopsis, even in the absence of phytochromes (Strasser *et al.*, 2010). However, other events that occur during chloroplast biogenesis require the signals transduced by photoreceptors. These signals ensure proper coordination of synthesis and import of LHCB proteins, which are essential for the assembly of the photosynthetic complexes. These events are also coordinated with the synthesis of carotenoids, which are necessary for photoprotection (Cazzonelli & Pogson, 2010).

Phytochromes, through the action of PIFs, regulate the transition from amiloplasts to etioplasts and to chloroplasts. For example, the PIFs inhibit the conversion of endodermal amyloplasts to etioplasts, whereas the phytochromes antagonise this inhibition, promoting the formation of chloroplasts (Figure 3) (Kim *et al.*, 2011).

#### **6.1 Chlorophyll biosynthesis is regulated by light**

Chlorophyll biosynthesis and the synthesis of other components of the photosystems are tightly regulated by light and the circadian clock. This coordination is necessary because when the chlorophyll synthesis exceeds the accumulation of chlorophyll-binding apoproteins, reactive oxygen species are generated, ultimately leading to cell death. However, when the chlorophyll synthesis is not enough, the amount of fully functional chlorophyll-binding proteins is not sufficient to gain optimal photosynthetic activity. Another example highlighting the importance of proper coordination is that PIF deficient plants accumulate protochlorophyllide in the dark during skotomorphogenic development, but this accumulation leads to bleaching upon exposure to light (Stephenson *et al.*, 2009).

Plants have four classes of tetrapyrroles: chlorophyll, phytochromobilin, haeme and siroheme, all derived from the same biosynthetic pathway. The flow of the tetrapyrrole pathway is strictly regulated, keeping at low levels the potentially toxic intermediates

identification of HY5, CCA1 and a DET1 responsive elements (Maxwell *et al.*, 2003). Similarly, it has been shown that HY5 binds to the *Lhcb1\*3* promoter and physically interacts

Another promoter analysed in more detail was the tobacco *Lhcb1\*2*. First, a 146 bp promoter fragment sufficed to confer VLFR (mediated by phyA), LFR (mediated by phyB) and HIR (mediated by phyA) to a minimal promoter (Cerdan *et al.*, 1997). Then, the motifs for VLFR and LFR were dissected from the HIR responsive motifs (Cerdan *et al.*, 2000) and finally, the TGGA motif was shown to bind Bell-like homeodomain 1 (BLH1) as part of the phyA mediated HIR (Staneloni *et al.*, 2009). This promoter is an example of how several different photoreceptors can regulate a single gene and integrate their signalling pathways at the promoter level; at least four different photoreceptors were shown to regulate this single

Proplastids are found in the embryo; they are undifferentiated plastids that are converted to other kind of plastids like chromoplasts, amyloplasts, chloroplasts and etioplasts. During skotomorphogenic development, proplastids turn into etioplasts, the chloroplast precursors. Etioplasts contain the prolamellar body, a structure rich in protochlorophyllide, the chlorophyll precursor, and the enzyme protochlorophyllide oxidoreductase (POR). During the development of etioplasts into chloroplasts, the POR is directly activated by light to convert protochlorophyllide into divinyl-chlorophyllide a, which is chlorophyll a and b precursor (Tanaka & Tanaka, 2007). This light-dependent step can be promoted by red-light in Arabidopsis, even in the absence of phytochromes (Strasser *et al.*, 2010). However, other events that occur during chloroplast biogenesis require the signals transduced by photoreceptors. These signals ensure proper coordination of synthesis and import of LHCB proteins, which are essential for the assembly of the photosynthetic complexes. These events are also coordinated with the synthesis of carotenoids, which are necessary for

Phytochromes, through the action of PIFs, regulate the transition from amiloplasts to etioplasts and to chloroplasts. For example, the PIFs inhibit the conversion of endodermal amyloplasts to etioplasts, whereas the phytochromes antagonise this inhibition, promoting

Chlorophyll biosynthesis and the synthesis of other components of the photosystems are tightly regulated by light and the circadian clock. This coordination is necessary because when the chlorophyll synthesis exceeds the accumulation of chlorophyll-binding apoproteins, reactive oxygen species are generated, ultimately leading to cell death. However, when the chlorophyll synthesis is not enough, the amount of fully functional chlorophyll-binding proteins is not sufficient to gain optimal photosynthetic activity. Another example highlighting the importance of proper coordination is that PIF deficient plants accumulate protochlorophyllide in the dark during skotomorphogenic development, but this accumulation leads to bleaching upon exposure to light (Stephenson *et al.*, 2009). Plants have four classes of tetrapyrroles: chlorophyll, phytochromobilin, haeme and siroheme, all derived from the same biosynthetic pathway. The flow of the tetrapyrrole pathway is strictly regulated, keeping at low levels the potentially toxic intermediates

with CCA1 to synergistically regulate expression (Andronis *et al.*, 2008).

promoter (Casal *et al.*, 1998; Cerdan *et al.*, 1999; Mazzella *et al.*, 2001) .

**6. Light promotes chloroplast development**

photoprotection (Cazzonelli & Pogson, 2010).

the formation of chloroplasts (Figure 3) (Kim *et al.*, 2011).

**6.1 Chlorophyll biosynthesis is regulated by light**

(Tanaka & Tanaka, 2007). Phytochrome and cryptochrome mutants contain lower levels of chlorophyll (Strasser *et al.*, 2010) stressing out the importance of the photomorphogenic signal for proper assembly of the photosynthetic machinery. In the next paragraphs we review how light signalling pathways regulate chlorophyll biosynthesis (Figure 4).

Fig. 3. Light interactions in plastid development. Phytochrome and PIFs roles during the transition from proplastid or amyloplast to chloroplast.

Chlorophyll synthesis occurs in plastids; in the first step glutamate is activated to GlutamyltRNA by the Glutamyl-tRNA synthetase, a step shared with plastid protein synthesis. The following step, the reduction of the Glutamyl-tRNA to produce glutamate-1-semialdehyde is subjected to tight regulation (Figure 4). In Arabidopsis, the Glutamyl-tRNA reductases are encoded by a little family of nuclear genes called *HEMA*. Of this family, the expression of *HEMA1* correlates with the expression of *Lhcb1* genes, which encode light-harvesting proteins of the photosystem II; in some way the expression of *HEMA1* reflects the demand of chlorophyll synthesis. On the other hand, *HEMA2* is not light regulated (Matsumoto *et al.*, 2004; McCormac *et al.*, 2001; McCormac & Terry, 2002a; McCormac & Terry, 2002b).

Glutamyl-tRNA reductase activity is regulated by negative feedback loops; the accumulation of Haeme, Mg-Protoporphyrin IX or Divinyl protochlorofilide a antagonise Glutamyl-tRNA reductase activity (Srivastava *et al.*, 2005). At the transcriptional level, *HEMA1* expression is induced by red and far-red light, implicating at least phyA and phyB, and blue light perceived by cry1 (McCormac *et al.*, 2001; McCormac & Terry, 2002a). *pif1* and *pif3* mutants contain higher levels of *HEMA1* mRNA, higher levels of protochlorophyllide and partially developed chloroplasts in the dark, a phenotype observed in *cop* mutants. The effects of *pif1* and *pif3* mutations are essentially additive, suggesting a model where phytochromes promote chloroplast biogenesis by antagonizing the activity of at least PIF1 and PIF3. As PIF1 and PIF3 are regulated by the circadian clock, but do not seem to affect central clock components (TOC1, CCA1, LHY), these PIFs seem to integrate chloroplast biogenesis with circadian and light signalling (Stephenson *et al.*, 2009).

The expression of photosynthetic nuclear genes is repressed by plastid signals if chloroplast biogenesis is blocked (retrograde signalling). This finding led to the isolation of mutants that disrupt chloroplast to nucleus communication, the genomes uncoupled mutants (*gun*) (Nott *et al.*, 2006). These mutants show high levels of *lhcb1* mRNA in the presence of norfluorzazon and were named *gun1* to *gun5*. *gun2* and *gun3* are allelic to *hy1* and *hy2* and disrupt phytochromobilin synthesis, leading to haeme accumulation and feedback

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 207

the cryptochromes, establishing GUN4 as a link between the phytochromes and the regulation of MgCH activity (Stephenson & Terry, 2008). *GUN5* encodes the H subunit of MgCH, known as CHLH (Nott *et al.*, 2006). The expression of *CHLH* is regulated at the mRNA level by light/dark cycles and by the circadian clock. Interestingly, this gene is co-regulated with *HEMA1*, *lhcb, Mg-protoporphyrin IX monomethyl estercyclase (MGCy)* and the gene encoding the chlorophyll(ide) a oxygenase (*CAO*) (Matsumoto *et al.*, 2004). On the other hand, *GUN1* encodes a pentatricopeptide repeat–containing protein that does not affect chlorophyll synthesis. GUN1 was proposed to generate a signal in chloroplast that represses nuclear photosynthetic gene expression; this repression on *lhcb* genes seems to be mediated by direct

Another connection between light signalling and the retrograde signalling was recently established. A sensitive genetic screening for the *gun* phenotype uncovered new cry1 alleles. These results establish that cry1 is necessary for maximal repression of *lhcb* genes, when

One of the latest steps in chlorophyll synthesis is the reduction of 3,8-divinyl protochlorophyllide to 3,8-divinyl chlorophyllide. This protochlorophyllide to chlorophyllide conversion is catalysed by the POR enzyme. In angiosperms, POR is light-dependent and it is likely the source of red-light promoted chlorophyll synthesis in the absence of phytochromes (Strasser *et al.*, 2010). Angiosperms carry three POR-encoding genes, *PorA, PorB* and *PorC*, which are differentially regulated by both light and developmental stage. *PORA* expression is high in etiolated seedlings and rapidly becomes undetectable after illumination with FR, a HIR response mediated by phyA, whereas *PORB* expression persists throughout greening and in adult plants (Runge *et al.*, 1996). PORC is expressed during the adult life and together with PORB is responsible for bulk chlorophyll synthesis in green plants (Paddock *et al.*, 2010). It has been recently shown that *PORC* expression is directly activated by PIF1 binding to a G-box in *PORC* promoter, whereas *PORA* and *PORB* are also induced by PIF1, presumably in an

During the last twenty years, plant biologists have witnessed major advances in our understanding of how plants use light as a source of information. These advances were possible thanks to the adoption of Arabidopsis as a model system. During these twenty years, 13 Arabidopsis photoreceptors were characterised In molecular terms and these findings extended to other species as well. A high number of signal transduction components were also characterised. With the advent of "omics" technologies, the networks that work downstream photoreceptors and their targets started to surface. However, with all these advances, we still do not know in detail how a single light responsive promoter works. How many transcription factors are sitting there? Which are their identities? How do they interact to fine tune expression under the diverse light conditions found in nature? If we multiply these questions by the number of light responsive promoters we can just have a

Akhilesh, K. T. &Gaur, T. (2003). Light regulation of nuclear photosynthetic genes in higher

plants. *Critical Reviews in Plant Sciences* 22(5): 417-452.

binding of *ABI4*, an AP2–type transcription factor (Koussevitzky *et al.*, 2007).

chloroplast biogenesis is blocked (Ruckle *et al.*, 2007).

indirect manner (Moon *et al.*, 2008).

hint of the enormous task ahead.

**7. Conclusion**

**8. References**

inhibition of Glutamyl-tRNA reductase (Nott *et al.*, 2006). The product of the *GUN4* gene, a 22 kD protein localized to Chloroplasts, promotes Magnesium chelatase (MgCH) activity which catalyses the insertion of Mg2+ into protoporphyrin IX (Tanaka & Tanaka, 2007). The *GUN4* gene is also under circadian clock regulation and is repressed by PIF1 and PIF3 suggesting a similar regulatory mechanism to HEMA1 (Stephenson *et al.*, 2009). The expression of GUN4 is primarily under the control of phyA and phyB with some input from

Fig. 4. Simplified chlorophyll biosynthesis pathway and light regulated steps. We emphasise how the light regulate directly the activity of NADPH:protochlorophyllide oxidoreductase (POR); or indirectly, through phytochrome and PIFs the expression of genes encoding the Glutamyl-tRNA reductases (HEMAs), Ferrum chelatase (FeCH), Magnesium chelatase (MgCH), NADPH:protochlorophyllide oxidoreductase (POR), Chlorophyllide a oxygenase (CAO), Mg-protoporphyrin IX methyltransferase (MgMT), and Mg-protoporphyrin IX monomethyl estercyclase (MgCy). The ATP/ADP ratio, the Mg2+ concentration and the thioredoxin levels also affect the MgCH activity, furthermore, these factors are light regulated (Tanaka & Tanaka, 2007). LHCs attach *chlorophyll a*, and CAO converts the *chlorophyll* a to *b* on the LHC apoprotein (Tanaka & Tanaka, 2007).

the cryptochromes, establishing GUN4 as a link between the phytochromes and the regulation of MgCH activity (Stephenson & Terry, 2008). *GUN5* encodes the H subunit of MgCH, known as CHLH (Nott *et al.*, 2006). The expression of *CHLH* is regulated at the mRNA level by light/dark cycles and by the circadian clock. Interestingly, this gene is co-regulated with *HEMA1*, *lhcb, Mg-protoporphyrin IX monomethyl estercyclase (MGCy)* and the gene encoding the chlorophyll(ide) a oxygenase (*CAO*) (Matsumoto *et al.*, 2004). On the other hand, *GUN1* encodes a pentatricopeptide repeat–containing protein that does not affect chlorophyll synthesis. GUN1 was proposed to generate a signal in chloroplast that represses nuclear photosynthetic gene expression; this repression on *lhcb* genes seems to be mediated by direct binding of *ABI4*, an AP2–type transcription factor (Koussevitzky *et al.*, 2007).

Another connection between light signalling and the retrograde signalling was recently established. A sensitive genetic screening for the *gun* phenotype uncovered new cry1 alleles. These results establish that cry1 is necessary for maximal repression of *lhcb* genes, when chloroplast biogenesis is blocked (Ruckle *et al.*, 2007).

One of the latest steps in chlorophyll synthesis is the reduction of 3,8-divinyl protochlorophyllide to 3,8-divinyl chlorophyllide. This protochlorophyllide to chlorophyllide conversion is catalysed by the POR enzyme. In angiosperms, POR is light-dependent and it is likely the source of red-light promoted chlorophyll synthesis in the absence of phytochromes (Strasser *et al.*, 2010). Angiosperms carry three POR-encoding genes, *PorA, PorB* and *PorC*, which are differentially regulated by both light and developmental stage. *PORA* expression is high in etiolated seedlings and rapidly becomes undetectable after illumination with FR, a HIR response mediated by phyA, whereas *PORB* expression persists throughout greening and in adult plants (Runge *et al.*, 1996). PORC is expressed during the adult life and together with PORB is responsible for bulk chlorophyll synthesis in green plants (Paddock *et al.*, 2010). It has been recently shown that *PORC* expression is directly activated by PIF1 binding to a G-box in *PORC* promoter, whereas *PORA* and *PORB* are also induced by PIF1, presumably in an indirect manner (Moon *et al.*, 2008).

#### **7. Conclusion**

206 Advances in Photosynthesis – Fundamental Aspects

inhibition of Glutamyl-tRNA reductase (Nott *et al.*, 2006). The product of the *GUN4* gene, a 22 kD protein localized to Chloroplasts, promotes Magnesium chelatase (MgCH) activity which catalyses the insertion of Mg2+ into protoporphyrin IX (Tanaka & Tanaka, 2007). The *GUN4* gene is also under circadian clock regulation and is repressed by PIF1 and PIF3 suggesting a similar regulatory mechanism to HEMA1 (Stephenson *et al.*, 2009). The expression of GUN4 is primarily under the control of phyA and phyB with some input from

Glutamate

**Light**

Phytochromes

PIFs

Protoheme

FeCH

Heme oxidases

Phytochromebilin

**Light**

Other Hemes

Glutamyl-tRNA

HemA

5-Aminoleculinic acid

Protoporphyrinogen IX

ATP Mg2+ Thiredoxin GUN4

PIFs Phytochromes

Chlorophyll b

Fig. 4. Simplified chlorophyll biosynthesis pathway and light regulated steps. We emphasise how the light regulate directly the activity of NADPH:protochlorophyllide oxidoreductase (POR); or indirectly, through phytochrome and PIFs the expression of genes encoding the Glutamyl-tRNA reductases (HEMAs), Ferrum chelatase (FeCH), Magnesium chelatase (MgCH), NADPH:protochlorophyllide oxidoreductase (POR), Chlorophyllide a oxygenase (CAO), Mg-protoporphyrin IX methyltransferase (MgMT), and Mg-protoporphyrin IX monomethyl estercyclase (MgCy). The ATP/ADP ratio, the Mg2+ concentration and the thioredoxin levels also affect the MgCH activity, furthermore, these factors are light regulated (Tanaka & Tanaka, 2007). LHCs attach *chlorophyll a*, and CAO converts the

Lchb1

Mg- protoporphyrin IX

FLU

Divinyl protochlorofilide a

**Light**

Divinyl chlorofilide a

Chlorophyll a

MgCH

MgCY MgMT

POR

CAO

*chlorophyll* a to *b* on the LHC apoprotein (Tanaka & Tanaka, 2007).

During the last twenty years, plant biologists have witnessed major advances in our understanding of how plants use light as a source of information. These advances were possible thanks to the adoption of Arabidopsis as a model system. During these twenty years, 13 Arabidopsis photoreceptors were characterised In molecular terms and these findings extended to other species as well. A high number of signal transduction components were also characterised. With the advent of "omics" technologies, the networks that work downstream photoreceptors and their targets started to surface. However, with all these advances, we still do not know in detail how a single light responsive promoter works. How many transcription factors are sitting there? Which are their identities? How do they interact to fine tune expression under the diverse light conditions found in nature? If we multiply these questions by the number of light responsive promoters we can just have a hint of the enormous task ahead.

#### **8. References**

Akhilesh, K. T. &Gaur, T. (2003). Light regulation of nuclear photosynthetic genes in higher plants. *Critical Reviews in Plant Sciences* 22(5): 417-452.

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 209

Chen, M., Galvao, R. M., Li, M., Burger, B., Bugea, J., Bolado, J. &Chory, J. (2010).

Deng, X. W., Matsui, M., Wei, N., Wagner, D., Chu, A. M., Feldmann, K. A. &Quail, P. H.

Dixon, L. E., Knox, K., Kozma-Bognar, L., Southern, M. M., Pokhilko, A. &Millar, A. J.

Edwards, K. D., Anderson, P. E., Hall, A., Salathia, N. S., Locke, J. C., Lynn, J. R., Straume,

Fankhauser, C. &Chen, M. (2008). Transposing phytochrome into the nucleus. *Trends Plant*

Farre, E. M., Harmer, S. L., Harmon, F. G., Yanovsky, M. J. &Kay, S. A. (2005). Overlapping

Favory, J. J., Stec, A., Gruber, H., Rizzini, L., Oravecz, A., Funk, M., Albert, A., Cloix, C.,

Franklin, K. A. &Quail, P. H. (2009). Phytochrome functions in Arabidopsis development. *J*

Fukushima, A., Kusano, M., Nakamichi, N., Kobayashi, M., Hayashi, N., Sakakibara, H.,

Gruber, H., Heijde, M., Heller, W., Albert, A., Seidlitz, H. K. &Ulm, R. (2010). Negative

Harmer, S. L. (2009). The circadian system in higher plants. *Annu Rev Plant Biol* 60: 357-377. Harmer, S. L., Hogenesch, J. B., Straume, M., Chang, H. S., Han, B., Zhu, T., Wang, X., Kreps,

Harmer, S. L. &Kay, S. A. (2005). Positive and negative factors confer phase-specific circadian regulation of transcription in Arabidopsis. *Plant Cell* 17(7): 1926-1940. Heim, M. A., Jakoby, M., Werber, M., Martin, C., Weisshaar, B. &Bailey, P. C. (2003). The

protein structure and functional diversity. *Mol Biol Evol* 20(5): 735-747. Henriques, R., Jang, I. C. &Chua, N. H. (2009). Regulated proteolysis in light-related

in Arabidopsis. *Proc Natl Acad Sci U S A* 107(46): 20132-20137.

by the circadian clock. *Science* 290(5499): 2110-2113.

signaling pathways. *Curr Opin Plant Biol* 12(1): 49-56.

binding motif and a G beta homologous domain. *Cell* 71(5): 791-801. Devlin, P. F. &Kay, S. A. (2000). Cryptochromes are required for phytochrome signaling to the circadian clock but not for rhythmicity. *Plant Cell* 12(12): 2499-2510. Ding, Z., Millar, A. J., Davis, A. M. &Davis, S. J. (2007). TIME FOR COFFEE encodes a

FLOWERING 3 in Arabidopsis. *Curr Biol* 21(2): 120-125.

acclimation in Arabidopsis. *Embo J* 28(5): 591-601.

*Cell* 141(7): 1230-1240.

*Cell* 18(3): 639-650.

*Sci* 13(11): 596-601.

*Exp Bot* 61(1): 11-24.

15(1): 47-54.

7251-7256.

1536.

Arabidopsis HEMERA/pTAC12 initiates photomorphogenesis by phytochromes.

(1992). COP1, an Arabidopsis regulatory gene, encodes a protein with both a zinc-

nuclear regulator in the Arabidopsis thaliana circadian clock. *Plant Cell* 19(5): 1522-

(2011). Temporal repression of core circadian genes is mediated through EARLY

M., Smith, J. Q. &Millar, A. J. (2006). FLOWERING LOCUS C mediates natural variation in the high-temperature response of the Arabidopsis circadian clock. *Plant*

and distinct roles of PRR7 and PRR9 in the Arabidopsis circadian clock. *Curr Biol*

Jenkins, G. I., Oakeley, E. J., Seidlitz, H. K., Nagy, F. &Ulm, R. (2009). Interaction of COP1 and UVR8 regulates UV-B-induced photomorphogenesis and stress

Mizuno, T. &Saito, K. (2009). Impact of clock-associated Arabidopsis pseudoresponse regulators in metabolic coordination. *Proc Natl Acad Sci U S A* 106(17):

feedback regulation of UV-B-induced photomorphogenesis and stress acclimation

J. A. &Kay, S. A. (2000). Orchestrated transcription of key pathways in Arabidopsis

basic helix-loop-helix transcription factor family in plants: a genome-wide study of

Alabadi, D., Oyama, T., Yanovsky, M. J., Harmon, F. G., Mas, P. &Kay, S. A. (2001).

Anderson, S. L., Teakle, G. R., Martino-Catt, S. J. &Kay, S. A. (1994). Circadian clock- and

Andronis, C., Barak, S., Knowles, S. M., Sugano, S. &Tobin, E. M. (2008). The clock protein

Bae, G. &Choi, G. (2008). Decoding of light signals by plant phytochromes and their

Ballaré, C. L. S., A.L.; Sánchez, R.A.; Radosevich, S. (1992). Photomorphogenic process in the

Baudry, A., Ito, S., Song, Y. H., Strait, A. A., Kiba, T., Lu, S., Henriques, R., Pruneda-Paz, J.

Brewer, P. B., Howles, P. A., Dorian, K., Griffith, M. E., Ishida, T., Kaplan-Levy, R. N., Kilinc,

Casal, J. J., Cerdan, P. D., Staneloni, R. J. &Cattaneo, L. (1998). Different phototransduction

Casal, J. J., Luccioni, L. G., Oliverio, K. A. &Boccalandro, H. E. (2003). Light, phytochrome

Cazzonelli, C. I. &Pogson, B. J. (2010). Source to sink: regulation of carotenoid biosynthesis

Cerdan, P. D., Staneloni, R. J., Casal, J. J. &Sanchez, R. A. (1997). A 146 bp fragment of the

Cerdan, P. D., Staneloni, R. J., Ortega, J., Bunge, M. M., Rodriguez-Batiller, M. J., Sanchez, R.

Cerdan, P. D., Yanovsky, M. J., Reymundo, F. C., Nagatani, A., Staneloni, R. J., Whitelam, G.

Covington, M. F., Maloof, J. N., Straume, M., Kay, S. A. &Harmer, S. L. (2008). Global

Chattopadhyay, S., Puente, P., Deng, X. W. &Wei, N. (1998). Combinatorial interaction of

and FHY1 in Arabidopsis thaliana. *Plant J* 18(5): 499-507.

growth and development. *Genome Biol* 9(8): R130.

agricultural environment. *Photochemistry and Photobiology* 56: 12.

circadian clock. *Science* 293(5531): 880-883.

the Arabidopsis CAB2 promoter. *Plant J* 6(4): 457-470.

expression in Arabidopsis. *Mol Plant* 1(1): 58-67.

*Plant Cell* 22(3): 606-622.

*Physiol* 116(4): 1533-1538.

*Biol* 33(2): 245-255.

12(7): 1203-1211.

in plants. *Trends Plant Sci* 15(5): 266-274.

12066-12074.

625-636.

interacting proteins. *Annu Rev Plant Biol* 59: 281-311.

Reciprocal regulation between TOC1 and LHY/CCA1 within the Arabidopsis

phytochrome-regulated transcription is conferred by a 78 bp cis-acting domain of

CCA1 and the bZIP transcription factor HY5 physically interact to regulate gene

L., Chua, N. H., Tobin, E. M., Kay, S. A. &Imaizumi, T. (2010). F-box proteins FKF1 and LKP2 act in concert with ZEITLUPE to control Arabidopsis clock progression.

A. &Smyth, D. R. (2004). PETAL LOSS, a trihelix transcription factor gene, regulates perianth architecture in the Arabidopsis flower. *Development* 131(16): 4035-4045. Bu, Q., Zhu, L., Dennis, M. D., Yu, L., Lu, S. X., Person, M. D., Tobin, E. M., Browning, K. S.

&Huq, E. (2011). Phosphorylation by CK2 enhances the rapid light-induced degradation of phytochrome interacting factor 1 in Arabidopsis. *J Biol Chem* 286(14):

kinetics of phytochrome A and phytochrome B in Arabidopsis thaliana. *Plant*

signalling and photomorphogenesis in Arabidopsis. *Photochem Photobiol Sci* 2(6):

tobacco Lhcb1\*2 promoter confers very-low-fluence, low-fluence and highirradiance responses of phytochrome to a minimal CaMV 35S promoter. *Plant Mol*

A. &Casal, J. J. (2000). Sustained but not transient phytochrome A signaling targets a region of an Lhcb1\*2 promoter not necessary for phytochrome B action. *Plant Cell*

C. &Casal, J. J. (1999). Regulation of phytochrome B signaling by phytochrome A

transcriptome analysis reveals circadian regulation of key pathways in plant

light-responsive elements plays a critical role in determining the response characteristics of light-regulated promoters in Arabidopsis. *Plant J* 15(1): 69-77.

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 211

Lee, J., He, K., Stolc, V., Lee, H., Figueroa, P., Gao, Y., Tongprasit, W., Zhao, H., Lee, I.

Leivar, P. &Quail, P. H. (2011). PIFs: pivotal components in a cellular signaling hub. *Trends*

Leivar, P., Tepperman, J. M., Monte, E., Calderon, R. H., Liu, T. L. &Quail, P. H. (2009).

Li, Q. H. &Yang, H. Q. (2007). Cryptochrome signaling in plants. *Photochem Photobiol* 83(1):

Li, G., Siddiqui, H., Teng, Y., Lin, R., Wan, X. Y., Li, J., Lau, O. S., Ouyang, X., Dai, M., Wan,

Lian, H. L., He, S. B., Zhang, Y. C., Zhu, D. M., Zhang, J. Y., Jia, K. P., Sun, S. X., Li, L.

Lin, R., Ding, L., Casola, C., Ripoll, D. R., Feschotte, C. &Wang, H. (2007). Transposase-

Liu, B., Zuo, Z., Liu, H., Liu, X. &Lin, C. (2011). Arabidopsis cryptochrome1 interacts with

Locke, J. C., Southern, M. M., Kozma-Bognar, L., Hibberd, V., Brown, P. E., Turner, M. S.

Maddonni, G. A., Otegui, M. E., Andrieu, B., Chelle, M. &Casal, J. J. (2002). Maize leaves

Martin-Tryon, E. L., Kreps, J. A. &Harmer, S. L. (2007). GIGANTEA acts in blue light

Martinez-Garcia, J. F., Huq, E. &Quail, P. H. (2000). Direct targeting of light signals to a promoter element-bound transcription factor. *Science* 288(5467): 859-863. Mas, P. (2005). Circadian clock signaling in Arabidopsis thaliana: from gene expression to

Mas, P., Kim, W. Y., Somers, D. E. &Kay, S. A. (2003). Targeted degradation of TOC1 by ZTL modulates circadian function in Arabidopsis thaliana. *Nature* 426(6966): 567-570. Matsumoto, F., Obayashi, T., Sasaki-Sekimoto, Y., Ohta, H., Takamiya, K. &Masuda, T.

Arabidopsis with a mini-array system. *Plant Physiol* 135(4): 2379-2391. Maxwell, B. B., Andersson, C. R., Poole, D. S., Kay, S. A. &Chory, J. (2003). HY5, Circadian

experimentation and mathematical analysis. *Mol Syst Biol* 1: 2005 0013. Ma, L., Gao, Y., Qu, L., Chen, Z., Li, J., Zhao, H. &Deng, X. W. (2002). Genomic evidence for

turn away from neighbors. *Plant Physiol* 130(3): 1181-1189.

physiology and development.  *Genes Dev* 25(10): 1023-1028.

Lin, C. &Todo, T. (2005). The cryptochromes. *Genome Biol* 6(5): 220.

Arabidopsis. *Plant Cell* 14(10): 2383-2398.

time regulation. *Plant Physiol* 143(1): 473-486.

731-749.

*Plant Cell*.

94-101.

622.

1034.

*Plant Sci* 16(1): 19-28.

318(5854): 1302-1305.

&Deng, X. W. (2007). Analysis of transcription factor HY5 genomic binding sites revealed its hierarchical role in light regulation of development. *Plant Cell* 19(3):

Definition of Early Transcriptional Circuitry Involved in Light-Induced Reversal of PIF-Imposed Repression of Photomorphogenesis in Young Arabidopsis Seedlings.

J., Devlin, P. F., Deng, X. W. &Wang, H. (2011). Coordinated transcriptional regulation underlying the circadian clock in Arabidopsis. *Nat Cell Biol* 13(5): 616-

&Yang, H.Q. (2011). Blue-light-dependent interaction of cryptochrome 1 with SPA1

derived transcription factors regulate light signaling in Arabidopsis. *Science*

SPA1 to suppress COP1 activity in response to blue light. *Genes Dev* 25(10): 1029-

&Millar, A. J. (2005). Extension of a genetic network model by iterative

COP1 as a repressor of light-regulated gene expression and development in

signaling and has biochemically separable roles in circadian clock and flowering

(2004). Gene expression profiling of the tetrapyrrole metabolic pathway in

Clock-Associated 1, and a cis-element, DET1 dark response element, mediate DET1

Holm, M., Ma, L. G., Qu, L. J. &Deng, X. W. (2002). Two interacting bZIP proteins are direct

Hornitschek, P., Lorrain, S., Zoete, V., Michielin, O. &Fankhauser, C. (2009). Inhibition of the

Huala, E., Oeller, P. W., Liscum, E., Han, I. S., Larsen, E. &Briggs, W. R. (1997). Arabidopsis

Imaizumi, T. (2010). Arabidopsis circadian clock and photoperiodism: time to think about

Ito, S., Kawamura, H., Niwa, Y., Nakamichi, N., Yamashino, T. &Mizuno, T. (2009). A

Jang, I. C., Henriques, R., Seo, H. S., Nagatani, A. &Chua, N. H. (2010). Arabidopsis

Kiba, T., Henriques, R., Sakakibara, H. &Chua, N. H. (2007). Targeted degradation of

Kikis, E. A., Khanna, R. &Quail, P. H. (2005). ELF4 is a phytochrome-regulated component

Kim, J., Kim, Y., Yeom, M., Kim, J. H. &Nam, H. G. (2008). FIONA1 is essential for

Kim, K., Shin, J., Lee, S. H., Kweon, H. S., Maloof, J. N. &Choi, G. (2011). Phytochromes

Kim, W. Y., Fujiwara, S., Suh, S. S., Kim, J., Kim, Y., Han, L., David, K., Putterill, J., Nam, H.

Kodama, Y., Suetsugu, N. &Wada, M. (2011). Novel protein-protein interaction family

Koussevitzky, S., Nott, A., Mockler, T. C., Hong, F., Sachetto-Martins, G., Surpin, M., Lim, J.,

Laubinger, S., Fittinghoff, K. &Hoecker, U. (2004). The SPA quartet: a family of WD-repeat

GIGANTEA in blue light. *Nature* 449(7160): 356-360.

gene expression. *Science* 316(5825): 715-719.

CAB EXPRESSION 1 (TOC1) gene. *Plant Cell Physiol* 50(2): 290-303. Ito, S., Nakamichi, N., Kiba, T., Yamashino, T. &Mizuno, T. (2007). Rhythmic and light-

Arabidopsis. *Genes Dev* 16(10): 1247-1259.

location. *Curr Opin Plant Biol* 13(1): 83-89.

*EMBO J* 28(24): 3893-3902.

*Physiol* 48(11): 1644-1651.

LHY. *Plant J* 44(2): 300-313.

*Sci U S A* 108(4): 1729-1734.

*Plant Cell* 16(9): 2293-2306.

2120-2123.

60: 407-431.

2530.

319.

490.

targets of COP1-mediated control of light-dependent gene expression in

shade avoidance response by formation of non-DNA binding bHLH heterodimers.

NPH1: a protein kinase with a putative redox-sensing domain. *Science* 278(5346):

genetic study of the Arabidopsis circadian clock with reference to the TIMING OF

inducible appearance of clock-associated pseudo-response regulator protein PRR9 through programmed degradation in the dark in Arabidopsis thaliana. *Plant Cell*

PHYTOCHROME INTERACTING FACTOR proteins promote phytochrome B polyubiquitination by COP1 E3 ligase in the nucleus. *Plant Cell* 22(7): 2370-2383. Jenkins, G. I. (2009). Signal transduction in responses to UV-B radiation. *Annu Rev Plant Biol*

PSEUDO-RESPONSE REGULATOR5 by an SCFZTL complex regulates clock function and photomorphogenesis in Arabidopsis thaliana. *Plant Cell* 19(8): 2516-

of a negative-feedback loop involving the central oscillator components CCA1 and

regulating period length in the Arabidopsis circadian clock. *Plant Cell* 20(2): 307-

inhibit hypocotyl negative gravitropism by regulating the development of endodermal amyloplasts through phytochrome-interacting factors. *Proc Natl Acad*

G. &Somers, D. E. (2007). ZEITLUPE is a circadian photoreceptor stabilized by

proteins involved in chloroplast movement response. *Plant Signal Behav* 6(4): 483-

Mittler, R. &Chory, J. (2007). Signals from chloroplasts converge to regulate nuclear

proteins with a central role in suppression of photomorphogenesis in arabidopsis.

The Photomorphogenic Signal: An Essential Component of Photoautotrophic Life 213

Rizzini, L., Favory, J. J., Cloix, C., Faggionato, D., O'Hara, A., Kaiserli, E., Baumeister, R.,

Robson, P. R., McCormac, A. C., Irvine, A. S. &Smith, H. (1996). Genetic engineering of

Rockwell, N. C., Su, Y. S. &Lagarias, J. C. (2006). Phytochrome structure and signaling

Ruckle, M. E., DeMarco, S. M. &Larkin, R. M. (2007). Plastid signals remodel light signaling

Runge, S., Sperling, U., Frick, G., Apel, K. &Armstrong, G. A. (1996). Distinct roles for light-

Sharrock, R. A. (2008). The phytochrome red/far-red photoreceptor superfamily. *Genome*

Shen, H., Zhu, L., Castillon, A., Majee, M., Downie, B. &Huq, E. (2008). Light-induced

interactions with photoactivated phytochromes. *Plant Cell* 20(6): 1586-1602. Somers, D. E., Devlin, P. F. &Kay, S. A. (1998). Phytochromes and cryptochromes in the entrainment of the Arabidopsis circadian clock. *Science* 282(5393): 1488-1490. Srivastava, A., Lake, V., Nogaj, L. A., Mayer, S. M., Willows, R. D. &Beale, S. I. (2005). The

Staneloni, R. J., Rodriguez-Batiller, M. J., Legisa, D., Scarpin, M. R., Agalou, A., Cerdan, P.

Stephenson, P. G., Fankhauser, C. &Terry, M. J. (2009). PIF3 is a repressor of chloroplast

Stephenson, P. G. &Terry, M. J. (2008). Light signalling pathways regulating the Mg-

Strasser, B., Sanchez-Lamas, M., Yanovsky, M. J., Casal, J. J. &Cerdan, P. D. (2010).

Strayer, C., Oyama, T., Schultz, T. F., Raman, R., Somers, D. E., Mas, P., Panda, S., Kreps, J.

Tanaka, R. &Tanaka, A. (2007). Tetrapyrrole biosynthesis in higher plants. *Annu Rev Plant*

Toledo-Ortiz, G., Huq, E. &Quail, P. H. (2003). The Arabidopsis basic/helix-loop-helix

autoregulatory response regulator homolog. *Science* 289(5480): 768-771. Takano, M., Inagaki, N., Xie, X., Kiyota, S., Baba-Kasai, A., Tanabata, T. &Shinomura, T.

Arabidopsis UVR8 protein. *Science* 332(6025): 103-106.

mechanisms. *Annu Rev Plant Biol* 57: 837-858.

greening in higher plants. *Plant J* 9(4): 513-523.

expressed enzyme. *Plant Mol Biol* 58(5): 643-658.

thaliana. *Photochem Photobiol Sci* 7(10): 1243-1252.

rice. *Proc Natl Acad Sci U S A* 106(34): 14705-14710.

transcription factor family. *Plant Cell* 15(8): 1749-1770.

development. *Proc Natl Acad Sci U S A* 106(18): 7654-7659.

*Sci U S A* 106(32): 13624-13629.

*Biotechnol* 14(8): 995-998.

*Cell* 19(12): 3944-3960.

*Biol* 9(8): 230.

4776-4781.

*Biol* 58: 321-346.

Schafer, E., Nagy, F., Jenkins, G. I. &Ulm, R. (2011). Perception of UV-B by the

harvest index in tobacco through overexpression of a phytochrome gene. *Nat*

networks and are essential for efficient chloroplast biogenesis in Arabidopsis. *Plant*

dependent NADPH:protochlorophyllide oxidoreductases (POR) A and B during

phosphorylation and degradation of the negative regulator PHYTOCHROME-INTERACTING FACTOR1 from Arabidopsis depend upon its direct physical

Chlamydomonas reinhardtii gtr gene encoding the tetrapyrrole biosynthetic enzyme glutamyl-trna reductase: structure of the gene and properties of the

D., Meijer, A. H., Ouwerkerk, P. B. &Casal, J. J. (2009). Bell-like homeodomain selectively regulates the high-irradiance response of phytochrome A. *Proc Natl Acad*

chelatase branchpoint of chlorophyll synthesis during de-etiolation in Arabidopsis

Arabidopsis thaliana life without phytochromes. *Proc Natl Acad Sci U S A* 107(10):

A. &Kay, S. A. (2000). Cloning of the Arabidopsis clock gene TOC1, an

(2009). Phytochromes are the sole photoreceptors for perceiving red/far-red light in

regulation of chlorophyll a/b-binding protein 2 expression. *Plant Physiol* 133(4): 1565-1577.

Mazzella, M. A., Cerdan, P. D., Staneloni, R. J. &Casal, J. J. (2001). Hierarchical coupling of

McCormac, A. C., Fischer, A., Kumar, A. M., Soll, D. &Terry, M. J. (2001). Regulation of

McCormac, A. C. &Terry, M. J. (2002a). Light-signalling pathways leading to the co-

McCormac, A. C. &Terry, M. J. (2002b). Loss of nuclear gene expression during the

Moglich, A., Yang, X., Ayers, R. A. &Moffat, K. (2010). Structure and function of plant

Moon, J., Zhu, L., Shen, H. &Huq, E. (2008). PIF1 directly and indirectly regulates

Nakamichi, N., Kita, M., Ito, S., Sato, E., Yamashino, T. &Mizuno, T. (2005a). The

essential roles for circadian clock function. *Plant Cell Physiol* 46(4): 609-619. Nakamichi, N., Kita, M., Ito, S., Yamashino, T. &Mizuno, T. (2005b). PSEUDO-RESPONSE

circadian clock of Arabidopsis thaliana. *Plant Cell Physiol* 46(5): 686-698. Ni, M., Tepperman, J. M. &Quail, P. H. (1998). PIF3, a phytochrome-interacting factor

Nott, A., Jung, H. S., Koussevitzky, S. &Chory, J. (2006). Plastid-to-nucleus retrograde

Osterlund, M. T., Hardtke, C. S., Wei, N. &Deng, X. W. (2000). Targeted destabilization of

Paddock, T. N., Mason, M. E., Lima, D. F. &Armstrong, G. A. (2010). Arabidopsis

Pagnussat, G. C., Yu, H. J., Ngo, Q. A., Rajani, S., Mayalagu, S., Johnson, C. S., Capron, A.,

Para, A., Farre, E. M., Imaizumi, T., Pruneda-Paz, J. L., Harmon, F. G. &Kay, S. A. (2007).

Pruneda-Paz, J. L., Breton, G., Para, A. &Kay, S. A. (2009). A functional genomics approach

Puente, P., Wei, N. &Deng, X. W. (1996). Combinatorial interplay of promoter elements

expression in Arabidopsis. *Embo J* 15(14): 3732-3743.

Arabidopsis development. *Development* 128(12): 2291-2299.

Arabidopsis thaliana. *Plant J* 25(5): 549-561.

Arabidopsis thaliana. *Plant J* 32(4): 549-559.

photoreceptors. *Annu Rev Plant Biol* 61: 21-47.

*Acad Sci U S A* 105(27): 9433-9438.

helix protein. *Cell* 95(5): 657-667.

*Development* 132(3): 603-614.

*Plant Cell* 19(11): 3462-3473.

signaling. *Annu Rev Plant Biol* 57: 739-759.

1565-1577.

402-414.

457.

1481-1485.

regulation of chlorophyll a/b-binding protein 2 expression. *Plant Physiol* 133(4):

phytochromes and cryptochromes reconciles stability and light modulation of

HEMA1 expression by phytochrome and a plastid signal during de-etiolation in

ordinated expression of HEMA1 and Lhcb during chloroplast development in

phytochrome A-mediated far-red block of greening response. *Plant Physiol* 130(1):

chlorophyll biosynthesis to optimize the greening process in Arabidopsis. *Proc Natl*

Arabidopsis pseudo-response regulators, PRR5 and PRR7, coordinately play

REGULATORS, PRR9, PRR7 and PRR5, together play essential roles close to the

necessary for normal photoinduced signal transduction, is a novel basic helix-loop-

HY5 during light-regulated development of Arabidopsis. *Nature* 405(6785): 462-466.

protochlorophyllide oxidoreductase A (PORA) restores bulk chlorophyll synthesis and normal development to a porB porC double mutant. *Plant Mol Biol* 72(4-5): 445-

Xie, L. F., Ye, D. &Sundaresan, V. (2005). Genetic and molecular identification of genes required for female gametophyte development and function in Arabidopsis.

PRR3 Is a vascular regulator of TOC1 stability in the Arabidopsis circadian clock.

reveals CHE as a component of the Arabidopsis circadian clock. *Science* 323(5920):

constitutes the minimal determinants for light and developmental control of gene

**11**

*Japan*

*Kyushu University*

**Chloroplast Photorelocation Movement:**

**to Perform Efficient Photosynthesis**

Noriyuki Suetsugu and Masamitsu Wada

**A Sophisticated Strategy for Chloroplasts**

Chloroplasts move to weak light so that they can perceive light efficiently (the accumulation response), whereas they escape from strong light to avoid photodamage (the avoidance

Fig. 1. Typical intracellular distribution pattern of chloroplasts by their photorelocation movement. In darkness, chloroplasts are located on the cell bottom in *Arabidopsis thaliana*. Note that the dark position varies among plant species. Weak light induces the chloroplast accumulation response along the peliclinal walls so that chloroplasts can perceive light efficiently. Strong light induces the chloroplast avoidance response toward the anticlinal

The phototropin photoreceptor family of proteins, which includes phototropin (phot) and neochrome (neo), mediate chloroplast photorelocation movement in green plants (reviewed

**1. Introduction**

response) (Fig. 1).

walls to reduce photodamage.

### **Chloroplast Photorelocation Movement: A Sophisticated Strategy for Chloroplasts to Perform Efficient Photosynthesis**

Noriyuki Suetsugu and Masamitsu Wada *Kyushu University Japan*

#### **1. Introduction**

214 Advances in Photosynthesis – Fundamental Aspects

Viczian, A., Kircher, S., Fejes, E., Millar, A. J., Schafer, E., Kozma-Bognar, L. &Nagy, F.

Wang, Z. Y., Kenigsbuch, D., Sun, L., Harel, E., Ong, M. S. &Tobin, E. M. (1997). A Myb-

Wu, J. F., Wang, Y. &Wu, S. H. (2008). Two new clock proteins, LWD1 and LWD2, regulate

Yamashino, T., Matsushika, A., Fujimori, T., Sato, S., Kato, T., Tabata, S. &Mizuno, T. (2003).

Yanovsky, M. J., Mazzella, M. A. &Casal, J. J. (2000). A quadruple photoreceptor mutant still

Yanovsky, M. J., Mazzella, M. A., Whitelam, G. C. &Casal, J. J. (2001). Resetting of the

Yanovsky, M. J. Casal., J.J & Whitelam, G.C. (1995). Phytochrome A, phytochrome B and

Yu, J. W., Rubio, V., Lee, N. Y., Bai, S., Lee, S. Y., Kim, S. S., Liu, L., Zhang, Y., Irigoyen, M.

Zeilinger, M. N., Farre, E. M., Taylor, S. R., Kay, S. A. &Doyle, F. J., 3rd (2006). A novel

Zhang, H., He, H., Wang, X., Yang, X., Li, L. &Deng, X. W. (2011). Genome-wide mapping of

Zhu, D., Maier, A., Lee, J. H., Laubinger, S., Saijo, Y., Wang, H., Qu, L. J., Hoecker, U.

and post-transcriptional regulation. *Plant J* 65(3): 346-358.

Arabidopsis photoperiodic flowering. *Plant Physiol* 148(2): 948-959.

in Arabidopsis thaliana. *Plant Cell Physiol* 44(6): 619-629.

Arabidopsis Lhcb gene. *Plant Cell* 9(4): 491-507.

keeps track of time. *Curr Biol* 10(16): 1013-1015.

498.

16(6): 523-530.

*Cell & Environment* 18: 788-794.

and PRR9. *Mol Syst Biol* 2: 58.

*Mol Cell* 32(5): 617-630.

21(10): 841-847.

(2005). Functional characterization of phytochrome interacting factor 3 for the Arabidopsis thaliana circadian clockwork. *Plant Cell Physiol* 46(10): 1591-1602. Wang, Y., Wu, J. F., Nakamichi, N., Sakakibara, H., Nam, H. G. &Wu, S. H. (2011). LIGHT-

REGULATED WD1 and PSEUDO-RESPONSE REGULATOR9 form a positive feedback regulatory loop in the Arabidopsis circadian clock. *Plant Cell* 23(2): 486-

related transcription factor is involved in the phytochrome regulation of an

A Link between circadian-controlled bHLH factors and the APRR1/TOC1 quintet

circadian clock by phytochromes and cryptochromes in Arabidopsis. *J Biol Rhythms*

HY4 are involved in hypocotyl growth responses to natural radiation in Arabidopsis: weak de-etiolation of the phyA mutant under dense canopies. *Plant*

L., Sullivan, J. A., Lee, I., Xie, Q., Paek, N. C. &Deng, X. W. (2008). COP1 and ELF3 control circadian function and photoperiodic flowering by regulating GI stability.

computational model of the circadian clock in Arabidopsis that incorporates PRR7

the HY5-mediated gene networks in Arabidopsis that involve both transcriptional

&Deng, X. W. (2008). Biochemical characterization of Arabidopsis complexes containing CONSTITUTIVELY PHOTOMORPHOGENIC1 and SUPPRESSOR OF PHYA proteins in light control of plant development. *Plant Cell* 20(9): 2307-2323. Zuo, Z., Liu, H., Liu, B., Liu, X. &Lin, C. (2011). Blue-light dependent interaction of CRY2

with SPA1 regulates COP1 activity and floral initiation in Arabidopsis. *Curr Biol*

Chloroplasts move to weak light so that they can perceive light efficiently (the accumulation response), whereas they escape from strong light to avoid photodamage (the avoidance response) (Fig. 1).

Fig. 1. Typical intracellular distribution pattern of chloroplasts by their photorelocation movement. In darkness, chloroplasts are located on the cell bottom in *Arabidopsis thaliana*. Note that the dark position varies among plant species. Weak light induces the chloroplast accumulation response along the peliclinal walls so that chloroplasts can perceive light efficiently. Strong light induces the chloroplast avoidance response toward the anticlinal walls to reduce photodamage.

The phototropin photoreceptor family of proteins, which includes phototropin (phot) and neochrome (neo), mediate chloroplast photorelocation movement in green plants (reviewed

Chloroplast Photorelocation Movement: A Sophisticated

mechanism and (iii) the contribution to photosynthesis.

discussed.

**detailed photobiological analyses**

Strategy for Chloroplaststo Perform Efficient Photosynthesis 217

facilitate leaf photosynthesis by allowing greater light penetration to lower parts within the leaf (reviewed by Suetsugu & Wada, 2009) (Fig. 4b & 4c). However, these hypotheses are

In this chapter, we review three topics of chloroplast photorelocation movement: (i) the insights gained from physiological and photobiological analyses, (ii) the molecular

Light-induced chloroplast movement (chloroplast photorelocation movement) has fascinated plant biologists since its discovery in the mid-nineteenth century (Böhm, 1856). Comprehensive analyses by Gustav Senn (1875-1945) of chloroplast movement in various plant species revealed the general responses of chloroplasts to light intensity and direction; chloroplasts are distributed at a position that ensures more efficient light absorption under weak light conditions, and they are positioned away from strong light, as if they had escaped (Senn, 1908). In land plant species, which generally bear multiple small chloroplasts in a cell, low light induces chloroplast movement and distribution toward the periclinal walls (the accumulation response), whereas strong light induces chloroplast avoidance toward the anticlinal walls (the avoidance response) (Fig. 1). Blue light is most effective at inducing chloroplast movement in most plant species, but in some cryptogam plant species, red light as well as blue light is effective. This red light effect exhibits red/far-red light reversibility, suggestive of the involvement of a red/far-red light receptor phytochrome. Detailed photobiological analyses, especially by the research groups of Wolfgang Haupt (1921-2005) and Jan Zurzycki (1925-1984), have provided many important insights on putative photoreceptor molecules that regulate chloroplast movement (Haupt, 1999; Zurzycki, 1980). They found that membrane-bound blue light photoreceptors other than phytochrome mediate blue-light-induced chloroplast movement in most plant species and that membrane-bound phytochromes mediate red-light induced chloroplast movement in some cryptogam plants. These predictions were demonstrated by the recent identification of photoreceptor genes in various plant species. The blue light receptor phot mediates bluelight-induced chloroplast movement in various plant species (Jarillo et al., 2001; Kagawa et al., 2001, 2004; Kasahara et al., 2004; Sakai et al., 2001). Neo, the chimeric photoreceptor that is a fusion of phytochrome and phototropin, regulates red-light-induced chloroplast movement in ferns and some green alga (Kawai et al., 2003; Suetsugu et al., 2005b). The photoreceptors are not discussed here; for a comprehensive review, see Suetsugu & Wada, 2005, 2007a, 2007b, 2009. First, we show our attempts to elucidate the mechanism of chloroplast photorelocation movement by detailed photobiological analyses. Second, we review recent molecular biological analyses of chloroplast photorelocation movement. Finally, the contribution of chloroplast movement and positioning to photosynthesis is

**2.1 Elucidation of the mechanism of chloroplast photorelocation movement by**

The underlying processes of chloroplast photorelocation movement can be categorized into three parts: photoperception, signal transduction and the motility system. Most of the photobiological analyses of chloroplast photorelocation movement were performed to

controversial and have not yet been clearly demonstrated experimentally.

**2. From physiological analyses to molecular genetic analyses**

by Suetsugu & Wada, 2007b, 2009). Phot mediates blue-light-induced chloroplast movement in most green plant species, and neo mediates red-light-induced chloroplast movement in ferns and some green alga (reviewed by Suetsugu & Wada, 2005, 2007a). Like other plant organelle movement responses, chloroplast photorelocation movement depends on actin filaments (reviewed by Wada & Suetsugu, 2004). Detailed physiological and photobiological analyses revealed that chloroplasts could move in any direction without turning or rolling within a short lag time during both the accumulation and the avoidance responses (Tsuboi et al., 2009; Tsuboi & Wada, 2011a). This fact argued that chloroplasts move by utilizing preexisting actin filaments and myosins. However, recent detailed microscopic analyses in the flowering plant *A. thaliana* (Kadota et al., 2009), the fern *Adiantum capillus-veneris* (Tsuboi & Wada, 2011b) and the moss *Physcomitrella patens* (Yamashita et al., 2011) have revealed that short actin filaments around the periphery of chloroplasts (called as cp-actin filaments) but not cytoplasmic actin cables are involved in chloroplast photorelocation movement and in the attachment to the plasma membrane (Fig. 2). Furthermore, chloroplast photorelocation movement was normal in all of the examined multiple-myosin knockout plants and even in myosin mutant plants severely defective in movements of the mitochondria, Golgi bodies, peroxisomes, endoplasmic reticulum and cytoplasm (Suetsugu et al., 2010b). Molecular genetic analyses using *A. thaliana* are of a great benefit to the study of chloroplast movement. First, various molecular factors that regulate cp-actin filament generation and reorganization during chloroplast movement can be identified (Fig. 3). Two phototropins, phot1 and phot2, mediate chloroplast photorelocation movement (Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001) by reorganizing cp-actin filaments (Kadota et al., 2009, Ichikawa et al., 2011). Two interacting coiled-coil proteins, WEB1 (weak chloroplast movement under blue light 1) and PMI2 (plastid movement impaired 2), regulate the velocity of chloroplast movement via light-induced cp-actin filament reorganization, possibly by suppressing JAC1 (J-domain protein required for chloroplast accumulation response 1) (Kodama et al., 2010, 2011; Luesse et al., 2006; Suetsugu et al., 2005a). A chloroplast outer envelope protein, CHUP1 (chloroplast unusual positioning 1), and two kinesin-like proteins, KAC1 (kinesin-like protein for actin-based chloroplast movement 1) and KAC2, are indispensable for cp-actin filament formation (Kadota et al., 2009; Oikawa et al., 2003; Suetsugu et al., 2010a). CHUP1 and KAC1 showed in vitro F-actin binding activity, and CHUP1 also interacted with G-actin and profilin in vitro (Oikawa et al., 2003; Schmidt von Braun & Schleiff, 2008; Suetsugu et al., 2010a), suggesting the direct involvement of these proteins in cp-actin filament generation and regulation. Most of these components are highly conserved in land plants from bryophytes to angiosperms (reviewed by Suetsugu et al., 2010b), suggesting that cp-actin filament-mediated chloroplast movement may facilitate the explosive evolution of land plants when in a fluctuating, ambient light environment. Second, the availability of mutants deficient in chloroplast movement encouraged us to verify a long-standing hypothesis that chloroplast movement is required for efficient photosynthesis in fluctuating light conditions. Experiments using mutants deficient in avoidance movement showed that the avoidance response is necessary for reducing photodamage under strong light conditions (Kasahara et al., 2002)(Fig. 4a). Some reports have suggested that the avoidance response (i.e. the distribution of chloroplasts on the anticlinal walls) affects CO2 diffusion by changing the chloroplast surface that is exposed to intercellular air spaces and that the avoidance response in the upper part of the leaf could

by Suetsugu & Wada, 2007b, 2009). Phot mediates blue-light-induced chloroplast movement in most green plant species, and neo mediates red-light-induced chloroplast movement in ferns and some green alga (reviewed by Suetsugu & Wada, 2005, 2007a). Like other plant organelle movement responses, chloroplast photorelocation movement depends on actin filaments (reviewed by Wada & Suetsugu, 2004). Detailed physiological and photobiological analyses revealed that chloroplasts could move in any direction without turning or rolling within a short lag time during both the accumulation and the avoidance responses (Tsuboi et al., 2009; Tsuboi & Wada, 2011a). This fact argued that chloroplasts move by utilizing preexisting actin filaments and myosins. However, recent detailed microscopic analyses in the flowering plant *A. thaliana* (Kadota et al., 2009), the fern *Adiantum capillus-veneris* (Tsuboi & Wada, 2011b) and the moss *Physcomitrella patens* (Yamashita et al., 2011) have revealed that short actin filaments around the periphery of chloroplasts (called as cp-actin filaments) but not cytoplasmic actin cables are involved in chloroplast photorelocation movement and in the attachment to the plasma membrane (Fig. 2). Furthermore, chloroplast photorelocation movement was normal in all of the examined multiple-myosin knockout plants and even in myosin mutant plants severely defective in movements of the mitochondria, Golgi bodies, peroxisomes, endoplasmic reticulum and cytoplasm (Suetsugu et al., 2010b). Molecular genetic analyses using *A. thaliana* are of a great benefit to the study of chloroplast movement. First, various molecular factors that regulate cp-actin filament generation and reorganization during chloroplast movement can be identified (Fig. 3). Two phototropins, phot1 and phot2, mediate chloroplast photorelocation movement (Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001) by reorganizing cp-actin filaments (Kadota et al., 2009, Ichikawa et al., 2011). Two interacting coiled-coil proteins, WEB1 (weak chloroplast movement under blue light 1) and PMI2 (plastid movement impaired 2), regulate the velocity of chloroplast movement via light-induced cp-actin filament reorganization, possibly by suppressing JAC1 (J-domain protein required for chloroplast accumulation response 1) (Kodama et al., 2010, 2011; Luesse et al., 2006; Suetsugu et al., 2005a). A chloroplast outer envelope protein, CHUP1 (chloroplast unusual positioning 1), and two kinesin-like proteins, KAC1 (kinesin-like protein for actin-based chloroplast movement 1) and KAC2, are indispensable for cp-actin filament formation (Kadota et al., 2009; Oikawa et al., 2003; Suetsugu et al., 2010a). CHUP1 and KAC1 showed in vitro F-actin binding activity, and CHUP1 also interacted with G-actin and profilin in vitro (Oikawa et al., 2003; Schmidt von Braun & Schleiff, 2008; Suetsugu et al., 2010a), suggesting the direct involvement of these proteins in cp-actin filament generation and regulation. Most of these components are highly conserved in land plants from bryophytes to angiosperms (reviewed by Suetsugu et al., 2010b), suggesting that cp-actin filament-mediated chloroplast movement may facilitate the explosive evolution of land plants when in a fluctuating, ambient light environment. Second, the availability of mutants deficient in chloroplast movement encouraged us to verify a long-standing hypothesis that chloroplast movement is required for efficient photosynthesis in fluctuating light conditions. Experiments using mutants deficient in avoidance movement showed that the avoidance response is necessary for reducing photodamage under strong light conditions (Kasahara et al., 2002)(Fig. 4a). Some reports have suggested that the avoidance response (i.e. the distribution of chloroplasts on the anticlinal walls) affects CO2 diffusion by changing the chloroplast surface that is exposed to intercellular air spaces and that the avoidance response in the upper part of the leaf could facilitate leaf photosynthesis by allowing greater light penetration to lower parts within the leaf (reviewed by Suetsugu & Wada, 2009) (Fig. 4b & 4c). However, these hypotheses are controversial and have not yet been clearly demonstrated experimentally.

In this chapter, we review three topics of chloroplast photorelocation movement: (i) the insights gained from physiological and photobiological analyses, (ii) the molecular mechanism and (iii) the contribution to photosynthesis.

#### **2. From physiological analyses to molecular genetic analyses**

Light-induced chloroplast movement (chloroplast photorelocation movement) has fascinated plant biologists since its discovery in the mid-nineteenth century (Böhm, 1856). Comprehensive analyses by Gustav Senn (1875-1945) of chloroplast movement in various plant species revealed the general responses of chloroplasts to light intensity and direction; chloroplasts are distributed at a position that ensures more efficient light absorption under weak light conditions, and they are positioned away from strong light, as if they had escaped (Senn, 1908). In land plant species, which generally bear multiple small chloroplasts in a cell, low light induces chloroplast movement and distribution toward the periclinal walls (the accumulation response), whereas strong light induces chloroplast avoidance toward the anticlinal walls (the avoidance response) (Fig. 1). Blue light is most effective at inducing chloroplast movement in most plant species, but in some cryptogam plant species, red light as well as blue light is effective. This red light effect exhibits red/far-red light reversibility, suggestive of the involvement of a red/far-red light receptor phytochrome. Detailed photobiological analyses, especially by the research groups of Wolfgang Haupt (1921-2005) and Jan Zurzycki (1925-1984), have provided many important insights on putative photoreceptor molecules that regulate chloroplast movement (Haupt, 1999; Zurzycki, 1980). They found that membrane-bound blue light photoreceptors other than phytochrome mediate blue-light-induced chloroplast movement in most plant species and that membrane-bound phytochromes mediate red-light induced chloroplast movement in some cryptogam plants. These predictions were demonstrated by the recent identification of photoreceptor genes in various plant species. The blue light receptor phot mediates bluelight-induced chloroplast movement in various plant species (Jarillo et al., 2001; Kagawa et al., 2001, 2004; Kasahara et al., 2004; Sakai et al., 2001). Neo, the chimeric photoreceptor that is a fusion of phytochrome and phototropin, regulates red-light-induced chloroplast movement in ferns and some green alga (Kawai et al., 2003; Suetsugu et al., 2005b). The photoreceptors are not discussed here; for a comprehensive review, see Suetsugu & Wada, 2005, 2007a, 2007b, 2009. First, we show our attempts to elucidate the mechanism of chloroplast photorelocation movement by detailed photobiological analyses. Second, we review recent molecular biological analyses of chloroplast photorelocation movement. Finally, the contribution of chloroplast movement and positioning to photosynthesis is discussed.

#### **2.1 Elucidation of the mechanism of chloroplast photorelocation movement by detailed photobiological analyses**

The underlying processes of chloroplast photorelocation movement can be categorized into three parts: photoperception, signal transduction and the motility system. Most of the photobiological analyses of chloroplast photorelocation movement were performed to

Chloroplast Photorelocation Movement: A Sophisticated

actual signals.

Strategy for Chloroplaststo Perform Efficient Photosynthesis 219

accumulation response are long-lived. (iv) Signals for the avoidance response can be override those for the accumulation response, at least in the irradiated area. Alternatively, it is possible that the signals for the avoidance response can be generated only when chloroplasts are directly irradiated with strong light. In this case, it is likely that the photoreceptor is localized on chloroplasts or that a plasma membrane-localized receptor

However, it is clear that the signals for the accumulation response are generated by the activation of photoreceptors in the irradiated area and are subsequently transferred toward chloroplasts. If measuring the speed of signal transfer for the accumulation response were possible, we could guess as to what is the putative signal by comparing the speeds between the putative signal and the known signaling molecules. When the chloroplast accumulation response was induced by microbeam irradiation in *A. capillus-veneris* protonemal cells, the onset of the accumulation movement lagged in proportion to the increase in the distance between the irradiated area and the chloroplasts, suggesting that the speed of the signal can be calculated as the lag time before the onset of movement (Tsuboi & Wada, 2010a, b). Similar calculations were also performed in *A. capillus-veneris* prothallial and *A. thaliana* mesophyll cells (Tsuboi & Wada, 2010a, b). These analyses revealed three interesting features of the putative signals. First, in protonemal cells, the speed of the signals in the basal-to-apical directions (about 2.3-2.4 µm min-1) was about three times faster than that in the apical-to-basal direction (about 0.6-0.9 µm min-1). However, the speed of the signals was almost equal in each cell type (about 0.9-1.1 µm min-1 in *A. capillus-veneris* prothallial cells and about 0.7 µm min-1 in *A. thaliana* mesophyll cells) (Tsuboi & Wada, 2010a, b). This difference in speed could result from the difference in the cell growth pattern of the cells, i.e. polarized (protonemal cells) and diffusive (prothallial and mesophyll cells). Second, in fern gametophytic cells, the speed of the signal and the maximum distance over which the signals could be transferred were almost equal irrespective of the intensity of the red or blue light microbeam, although in this case, more chloroplasts that were located farther away responded under continuous irradiation, compared to those submitted to a 1 min pulse of irradiation (Tsuboi & Wada, 2010a, b). Interestingly, the velocity of chloroplast accumulation movement was constant, regardless of the intensity of the microbeam placed on the prothallial cells (Kagawa & Wada, 1996; Tsuboi & Wada, 2011a). These results suggested that the properties of the signal, such as the speed, the amount and the activity, do not change in proportion to the change of light intensity. However, chloroplasts in the protonemal cells accumulated in the area that had been irradiated by a beam with a higher fluence rate, compared to adjacent areas that had been irradiated with beam of lower fluence rates of blue or red light. This result suggests that the amount or activity of the signal was increased when exposed to a beam with a higher fluence rate (Yatsuhashi et al., 1987; Yatsuhashi, 1996). Third and most importantly, the speed of signals (about 0.6-2.4 µm min-1) was much slower than that caused by calcium ion spiking or waves known to occur in plant and animal systems (about several µm sec-1 to 100 µm sec-1) (Tsuboi & Wada, 2010a, b). Furthermore, the signal transfer must not be actomyosin-dependent because the transfer of the signal still occurred when actin filaments were disrupted by treatment with inhibitor (Sato et al., 2001). Collectively, although the signals for chloroplast movement remained to be determined, our detailed physiological analyses will provide the clue to identify the

generates the signals only when it exists in close proximity to the chloroplasts.

identify the photoreceptor molecules (reviewed by Haupt, 1999; Zurzycki, 1980; Wada et al., 1993). Many pharmacological (i.e. treatment with chemicals and inhibitors) and microscopic (i.e. staining of the cytoskeleton) analyses have provided valuable insights, such as the possible involvement of calcium ions in the signal transduction pathway and the actin filament-dependency of the motility system (reviewed by Suetsugu & Wada, 2007b, 2009). However, the data from pharmacological treatment and microscopic observation of fixed samples should be carefully considered because of possible artifactual results. Thus, we decided to analyze chloroplast relocation movement using detailed physiological and photobiological analyses of the gametophytic cells of a fern *A. capillus-veneris* as a model system (reviewed by Wada, 2007). By changing light conditions, we can easily obtain two types of gametophytes: a filamentous protonemal cell or a two-dimensional prothallus, which is a cell sheet made of a one-cell layer. This fern regulates chloroplast movement by utilizing phot family proteins and actin filaments, like *A. thaliana* (Kadota & Wada, 1992; Kagawa et al., 2004; Tsuboi & Wada, 2011b). Using a microbeam irradiation system, we analyzed chloroplast photorelocation movement in protonemal and phothallial cells and elucidated several aspects of chloroplast movement, especially putative signaling molecules and movement.

#### **2.1.1 Physiological properties of putative signals in chloroplast photorelocation movement**

Blue light mediates the influx of calcium ions (Ca2+) into the cytosol and this influx is dependent upon phots in *A. thaliana* and *P. patens* (reviewed by Harada & Shimazaki, 2007). Importantly, a Ca2+ chelator inhibited chloroplast movement and external Ca2+ ions and Ca2+ ionophores changed the distribution of chloroplasts when placed in darkness (reviewed by Suetsugu & Wada, 2009). However, plasma membrane Ca2+ channel blockers, which effectively inhibited phot-dependent blue light-mediated Ca2+ influx (reviewed by Harada & Shimazaki, 2007), were totally ineffective in suppressing chloroplast photorelocation movement in various plant species (reviewed by Suetsugu & Wada, 2009). Thus, the putative signals that control chloroplast movement remain to be determined. To characterize the properties of these putative signals, chloroplast photorelocation movement was induced by partial cell irradiation with a microbeam irradiator and analyzed in detail (Kagawa & Wada, 1999, 2000; Tsuboi & Wada, 2010a, b).

An open question was whether the signals were different between the accumulation and avoidance responses. When a dark-adapted cell of an *A. capillus-veneris* prothallus (Kagawa & Wada, 1999) and an *A. thaliana* leaf (Kagawa & Wada, 2000) (in this situation, a few chloroplasts were on the upper periclinal walls) were partially irradiated with strong blue light, chloroplasts moved to the irradiated area but could not enter the beam area. Immediately after the light was turned off, the chloroplasts moved into the formerly irradiated area. Similar responses were also found in filamentous protonemal cells in *A. capillus-veneris* (Yatsuhashi et al., 1985) and *P. patens* (Kadota et al., 2000; Sato et al., 2001). These results suggested several characteristics of putative signals (reviewed by Suetsugu & Wada, 2009): (i) Signals for both the accumulation and the avoidance responses are simultaneously generated by strong light. (ii) Signals for the avoidance response function only at the irradiated area whereas those for the accumulation response can be transferred toward chloroplasts when located far from the irradiated area. (iii) Signals for the avoidance response disappear immediately after the light is turned off, whereas those for the

identify the photoreceptor molecules (reviewed by Haupt, 1999; Zurzycki, 1980; Wada et al., 1993). Many pharmacological (i.e. treatment with chemicals and inhibitors) and microscopic (i.e. staining of the cytoskeleton) analyses have provided valuable insights, such as the possible involvement of calcium ions in the signal transduction pathway and the actin filament-dependency of the motility system (reviewed by Suetsugu & Wada, 2007b, 2009). However, the data from pharmacological treatment and microscopic observation of fixed samples should be carefully considered because of possible artifactual results. Thus, we decided to analyze chloroplast relocation movement using detailed physiological and photobiological analyses of the gametophytic cells of a fern *A. capillus-veneris* as a model system (reviewed by Wada, 2007). By changing light conditions, we can easily obtain two types of gametophytes: a filamentous protonemal cell or a two-dimensional prothallus, which is a cell sheet made of a one-cell layer. This fern regulates chloroplast movement by utilizing phot family proteins and actin filaments, like *A. thaliana* (Kadota & Wada, 1992; Kagawa et al., 2004; Tsuboi & Wada, 2011b). Using a microbeam irradiation system, we analyzed chloroplast photorelocation movement in protonemal and phothallial cells and elucidated several aspects of chloroplast movement, especially putative signaling molecules

**2.1.1 Physiological properties of putative signals in chloroplast photorelocation**

(Kagawa & Wada, 1999, 2000; Tsuboi & Wada, 2010a, b).

Blue light mediates the influx of calcium ions (Ca2+) into the cytosol and this influx is dependent upon phots in *A. thaliana* and *P. patens* (reviewed by Harada & Shimazaki, 2007). Importantly, a Ca2+ chelator inhibited chloroplast movement and external Ca2+ ions and Ca2+ ionophores changed the distribution of chloroplasts when placed in darkness (reviewed by Suetsugu & Wada, 2009). However, plasma membrane Ca2+ channel blockers, which effectively inhibited phot-dependent blue light-mediated Ca2+ influx (reviewed by Harada & Shimazaki, 2007), were totally ineffective in suppressing chloroplast photorelocation movement in various plant species (reviewed by Suetsugu & Wada, 2009). Thus, the putative signals that control chloroplast movement remain to be determined. To characterize the properties of these putative signals, chloroplast photorelocation movement was induced by partial cell irradiation with a microbeam irradiator and analyzed in detail

An open question was whether the signals were different between the accumulation and avoidance responses. When a dark-adapted cell of an *A. capillus-veneris* prothallus (Kagawa & Wada, 1999) and an *A. thaliana* leaf (Kagawa & Wada, 2000) (in this situation, a few chloroplasts were on the upper periclinal walls) were partially irradiated with strong blue light, chloroplasts moved to the irradiated area but could not enter the beam area. Immediately after the light was turned off, the chloroplasts moved into the formerly irradiated area. Similar responses were also found in filamentous protonemal cells in *A. capillus-veneris* (Yatsuhashi et al., 1985) and *P. patens* (Kadota et al., 2000; Sato et al., 2001). These results suggested several characteristics of putative signals (reviewed by Suetsugu & Wada, 2009): (i) Signals for both the accumulation and the avoidance responses are simultaneously generated by strong light. (ii) Signals for the avoidance response function only at the irradiated area whereas those for the accumulation response can be transferred toward chloroplasts when located far from the irradiated area. (iii) Signals for the avoidance response disappear immediately after the light is turned off, whereas those for the

and movement.

**movement**

accumulation response are long-lived. (iv) Signals for the avoidance response can be override those for the accumulation response, at least in the irradiated area. Alternatively, it is possible that the signals for the avoidance response can be generated only when chloroplasts are directly irradiated with strong light. In this case, it is likely that the photoreceptor is localized on chloroplasts or that a plasma membrane-localized receptor generates the signals only when it exists in close proximity to the chloroplasts.

However, it is clear that the signals for the accumulation response are generated by the activation of photoreceptors in the irradiated area and are subsequently transferred toward chloroplasts. If measuring the speed of signal transfer for the accumulation response were possible, we could guess as to what is the putative signal by comparing the speeds between the putative signal and the known signaling molecules. When the chloroplast accumulation response was induced by microbeam irradiation in *A. capillus-veneris* protonemal cells, the onset of the accumulation movement lagged in proportion to the increase in the distance between the irradiated area and the chloroplasts, suggesting that the speed of the signal can be calculated as the lag time before the onset of movement (Tsuboi & Wada, 2010a, b). Similar calculations were also performed in *A. capillus-veneris* prothallial and *A. thaliana* mesophyll cells (Tsuboi & Wada, 2010a, b). These analyses revealed three interesting features of the putative signals. First, in protonemal cells, the speed of the signals in the basal-to-apical directions (about 2.3-2.4 µm min-1) was about three times faster than that in the apical-to-basal direction (about 0.6-0.9 µm min-1). However, the speed of the signals was almost equal in each cell type (about 0.9-1.1 µm min-1 in *A. capillus-veneris* prothallial cells and about 0.7 µm min-1 in *A. thaliana* mesophyll cells) (Tsuboi & Wada, 2010a, b). This difference in speed could result from the difference in the cell growth pattern of the cells, i.e. polarized (protonemal cells) and diffusive (prothallial and mesophyll cells). Second, in fern gametophytic cells, the speed of the signal and the maximum distance over which the signals could be transferred were almost equal irrespective of the intensity of the red or blue light microbeam, although in this case, more chloroplasts that were located farther away responded under continuous irradiation, compared to those submitted to a 1 min pulse of irradiation (Tsuboi & Wada, 2010a, b). Interestingly, the velocity of chloroplast accumulation movement was constant, regardless of the intensity of the microbeam placed on the prothallial cells (Kagawa & Wada, 1996; Tsuboi & Wada, 2011a). These results suggested that the properties of the signal, such as the speed, the amount and the activity, do not change in proportion to the change of light intensity. However, chloroplasts in the protonemal cells accumulated in the area that had been irradiated by a beam with a higher fluence rate, compared to adjacent areas that had been irradiated with beam of lower fluence rates of blue or red light. This result suggests that the amount or activity of the signal was increased when exposed to a beam with a higher fluence rate (Yatsuhashi et al., 1987; Yatsuhashi, 1996). Third and most importantly, the speed of signals (about 0.6-2.4 µm min-1) was much slower than that caused by calcium ion spiking or waves known to occur in plant and animal systems (about several µm sec-1 to 100 µm sec-1) (Tsuboi & Wada, 2010a, b). Furthermore, the signal transfer must not be actomyosin-dependent because the transfer of the signal still occurred when actin filaments were disrupted by treatment with inhibitor (Sato et al., 2001). Collectively, although the signals for chloroplast movement remained to be determined, our detailed physiological analyses will provide the clue to identify the actual signals.

Chloroplast Photorelocation Movement: A Sophisticated

photorelocation movement.

Strategy for Chloroplaststo Perform Efficient Photosynthesis 221

chloroplast movement is mediated by neo in several ferns and probably in some green algae (reviewed by Suetsugu & Wada, 2005, 2007a, 2007b, 2009). Regardless of significant advances in photoreceptor identification, the molecular mechanism of signal transduction and the identity of the motility system for chloroplast movement have been obscure. However, molecular genetic analyses using *A. thaliana* have identified several components that regulate chloroplast movement. Furthermore, recent imaging analyses have revealed that a novel actin-based mechanism governs chloroplast photorelocation and positioning. By combining these results, we could imagine the molecular framework of chloroplast

**2.2.1 Unique actin-based mechanism for chloroplast movement in land plants**

utilize an actin-based mechanism that is different from those of other organelles.

For many years, it was thought that chloroplasts moved along long cytosolic actin cables by myosin motor proteins, similar to the movements of other organelles. However, the aforementioned studies (Tsuboi & Wada, 2009, 2011a) suggested that chloroplasts could

To find the actin-based mechanism for chloroplast movement, we utilized Arabidopsis transgenic lines in which actin filaments could be visualized by various fusions of fluorescent proteins and actin binding proteins (such as GFP-talin and tdTomato-fimbrin) and analyzed the behavior of the actin filaments during chloroplast movement using a custom-made microscope and a confocal microscope (Kadota et al., 2009). Although cytoplasmic actin cables and filaments were associated with chloroplasts, they did not change much in response to light irradiation, and their behavior did not associate with directional chloroplast movement. Instead, we found that short actin filaments found around chloroplasts dynamically changed their structure in response to light irradiation and that their dynamics correlated with the direction and speed of chloroplast movement. We have named these actin filaments chloroplast-actin filaments (abbreviated as cp-actin filaments) (Kadota et al., 2009)(Fig. 2). When chloroplasts were stationary, cp-actin filaments were distributed around the chloroplast periphery. In response to strong blue light, cp-actin filaments transiently disappeared within about 30 seconds and then reappeared at the one side of the chloroplasts, which would eventually be the front region of the moving chloroplasts (Fig. 2). We called this pattern of localization of cp-actin filaments at the front region "biased" (Fig. 2). After biased cp-actin filaments were fully formed, chloroplasts moved toward the side where the cp-actin filaments accumulated (Kadota et al., 2009). The generation of biased cp-actin filaments was also found during the accumulation response that had been induced by weak blue light, but this was not accompanied by a transient disappearance of cp-actin filaments, unlike what occurred during the avoidance response (Kadota et al., 2009). Thus, the light-induced generation of biased cpactin filaments is a prerequisite for both the avoidance and the accumulation responses. Possibly, a transient disappearance of cp-actin filaments induced by strong blue light facilitated an acceleration of chloroplast avoidance movement. As more cp-actin filaments accumulated at the front halves of the chloroplasts, in relation to the rear halves, the velocity of chloroplast avoidance also increased. When irradiated with a higher fluence of blue light, even more cp-actin filaments accumulated at the front halves, and chloroplasts moved even faster (Kadota et al., 2009). Thus, strong light caused a greater difference in the amount of cp-actin filaments at certain locations on the chloroplasts because cp-actin filaments located at the rear halves of the chloroplasts did not increase after transient disappearance. Conversely, weak light could not induce transient disappearance of cp-actin filaments, so a greater difference in

#### **2.1.2 An actin-based motility system deduced by detailed observation of chloroplast movement**

For a long time, it was believed that the actomyosin system mediated chloroplast movement in various species. Many analyses using several kinds of techniques (such as inhibitor treatment, immunocytochemistry and observation of the in vivo dynamics of actin filaments) clarified the involvement of actin filaments in chloroplast movement in various plant species (reviewed by Suetsugu & Wada, 2009; Suetsugu et al., 2010b). However, the involvement of myosin motor proteins was still controversial (reviewed by Suetsugu et al., 2010b). If the actomyosin system is involved in chloroplast movement, it is expected that chloroplasts move along long actin cables that preexist or elongate in the direction of movement immediately after light exposure and, thus, that chloroplast movement should be polarized (i.e. parallel to actin cables). However, this was not the case with chloroplast photorelocation movement at least in *A. capillus-veneris* prothallial and *A. thaliana* mesophyll cells (Tsuboi & Wada, 2009, 2011a). Importantly, chloroplasts moved by sliding but not rolling during both the accumulation and the avoidance responses (Tsuboi & Wada, 2009, 2011a), suggesting that chloroplasts moved by attaching one side to the plasma membrane via actin filaments that spanned between the chloroplasts and the plasma membrane. When observed with a microscope, chloroplasts look elliptic (or dumbbell-shaped for dividing chloroplasts) but not completely round. Therefore, it is plausible that chloroplasts keep their long axis in parallel with the moving direction so that they can take the path of least resistance. If that is the case, then they should turn at an angle formed by an imaginary line spanning their long axis and a second imaginary line that connects the center of the chloroplast, at the original position, to the center of the irradiated area. However, chloroplasts were capable of moving in any direction even without turning. Even if chloroplasts turned immediately before or while they moved, the extent of their turning was so small (Tsuboi & Wada, 2009, 2011a). Exceptionally, chloroplasts of Arabidopsis mesophyll cells tended to adjust their short axis to be parallel with the moving direction during the avoidance movement, although they started to move without turning (Tsuboi & Wada, 2011a). Importantly, chloroplasts escaped from strong light by taking the shortest route, suggesting that they are capable of determining the location of the closest area that is out of the strong light (Tsuboi & Wada, 2011a). Moreover, when sequentially irradiated with weak or strong light, chloroplasts could change their moving direction according to the position of subsequent irradiated beam, with a short lag time (Tsuboi & Wada, 2009, 2011a). Collectively, these detailed microscopic analyses argued against the hypothesis that chloroplasts utilize of pre-existing actin filaments for photomovement and suggested that they move using actin filaments that dynamically reorganize in response to light irradiation.

#### **2.2 Conserved molecular mechanism of chloroplast photorelocation movement in land plants**

Generally, blue light is most effective in inducing chloroplast photorelocation movement, although red light can also induce the movement in some cryptogam plants (green algae, mosses and ferns). Phot is the blue light receptor for chloroplast movement and also mediates phototropism and stomatal opening (reviewed by Christie, 2007). Phototropins were identified in green plants, from green alga to seed plants, and were shown to regulate blue-light-induced chloroplast movement at least in *A. thaliana*, *A. capillus-veneris* and *P. patens* (reviewed by Suetsugu & Wada, 2005, 2007a, 2007b, 2009). Red-light-induced

**2.1.2 An actin-based motility system deduced by detailed observation of chloroplast**

**2.2 Conserved molecular mechanism of chloroplast photorelocation movement in**

Generally, blue light is most effective in inducing chloroplast photorelocation movement, although red light can also induce the movement in some cryptogam plants (green algae, mosses and ferns). Phot is the blue light receptor for chloroplast movement and also mediates phototropism and stomatal opening (reviewed by Christie, 2007). Phototropins were identified in green plants, from green alga to seed plants, and were shown to regulate blue-light-induced chloroplast movement at least in *A. thaliana*, *A. capillus-veneris* and *P. patens* (reviewed by Suetsugu & Wada, 2005, 2007a, 2007b, 2009). Red-light-induced

For a long time, it was believed that the actomyosin system mediated chloroplast movement in various species. Many analyses using several kinds of techniques (such as inhibitor treatment, immunocytochemistry and observation of the in vivo dynamics of actin filaments) clarified the involvement of actin filaments in chloroplast movement in various plant species (reviewed by Suetsugu & Wada, 2009; Suetsugu et al., 2010b). However, the involvement of myosin motor proteins was still controversial (reviewed by Suetsugu et al., 2010b). If the actomyosin system is involved in chloroplast movement, it is expected that chloroplasts move along long actin cables that preexist or elongate in the direction of movement immediately after light exposure and, thus, that chloroplast movement should be polarized (i.e. parallel to actin cables). However, this was not the case with chloroplast photorelocation movement at least in *A. capillus-veneris* prothallial and *A. thaliana* mesophyll cells (Tsuboi & Wada, 2009, 2011a). Importantly, chloroplasts moved by sliding but not rolling during both the accumulation and the avoidance responses (Tsuboi & Wada, 2009, 2011a), suggesting that chloroplasts moved by attaching one side to the plasma membrane via actin filaments that spanned between the chloroplasts and the plasma membrane. When observed with a microscope, chloroplasts look elliptic (or dumbbell-shaped for dividing chloroplasts) but not completely round. Therefore, it is plausible that chloroplasts keep their long axis in parallel with the moving direction so that they can take the path of least resistance. If that is the case, then they should turn at an angle formed by an imaginary line spanning their long axis and a second imaginary line that connects the center of the chloroplast, at the original position, to the center of the irradiated area. However, chloroplasts were capable of moving in any direction even without turning. Even if chloroplasts turned immediately before or while they moved, the extent of their turning was so small (Tsuboi & Wada, 2009, 2011a). Exceptionally, chloroplasts of Arabidopsis mesophyll cells tended to adjust their short axis to be parallel with the moving direction during the avoidance movement, although they started to move without turning (Tsuboi & Wada, 2011a). Importantly, chloroplasts escaped from strong light by taking the shortest route, suggesting that they are capable of determining the location of the closest area that is out of the strong light (Tsuboi & Wada, 2011a). Moreover, when sequentially irradiated with weak or strong light, chloroplasts could change their moving direction according to the position of subsequent irradiated beam, with a short lag time (Tsuboi & Wada, 2009, 2011a). Collectively, these detailed microscopic analyses argued against the hypothesis that chloroplasts utilize of pre-existing actin filaments for photomovement and suggested that they move using actin filaments that dynamically

**movement**

reorganize in response to light irradiation.

**land plants**

chloroplast movement is mediated by neo in several ferns and probably in some green algae (reviewed by Suetsugu & Wada, 2005, 2007a, 2007b, 2009). Regardless of significant advances in photoreceptor identification, the molecular mechanism of signal transduction and the identity of the motility system for chloroplast movement have been obscure. However, molecular genetic analyses using *A. thaliana* have identified several components that regulate chloroplast movement. Furthermore, recent imaging analyses have revealed that a novel actin-based mechanism governs chloroplast photorelocation and positioning. By combining these results, we could imagine the molecular framework of chloroplast photorelocation movement.

#### **2.2.1 Unique actin-based mechanism for chloroplast movement in land plants**

For many years, it was thought that chloroplasts moved along long cytosolic actin cables by myosin motor proteins, similar to the movements of other organelles. However, the aforementioned studies (Tsuboi & Wada, 2009, 2011a) suggested that chloroplasts could utilize an actin-based mechanism that is different from those of other organelles.

To find the actin-based mechanism for chloroplast movement, we utilized Arabidopsis transgenic lines in which actin filaments could be visualized by various fusions of fluorescent proteins and actin binding proteins (such as GFP-talin and tdTomato-fimbrin) and analyzed the behavior of the actin filaments during chloroplast movement using a custom-made microscope and a confocal microscope (Kadota et al., 2009). Although cytoplasmic actin cables and filaments were associated with chloroplasts, they did not change much in response to light irradiation, and their behavior did not associate with directional chloroplast movement. Instead, we found that short actin filaments found around chloroplasts dynamically changed their structure in response to light irradiation and that their dynamics correlated with the direction and speed of chloroplast movement. We have named these actin filaments chloroplast-actin filaments (abbreviated as cp-actin filaments) (Kadota et al., 2009)(Fig. 2). When chloroplasts were stationary, cp-actin filaments were distributed around the chloroplast periphery. In response to strong blue light, cp-actin filaments transiently disappeared within about 30 seconds and then reappeared at the one side of the chloroplasts, which would eventually be the front region of the moving chloroplasts (Fig. 2). We called this pattern of localization of cp-actin filaments at the front region "biased" (Fig. 2). After biased cp-actin filaments were fully formed, chloroplasts moved toward the side where the cp-actin filaments accumulated (Kadota et al., 2009). The generation of biased cp-actin filaments was also found during the accumulation response that had been induced by weak blue light, but this was not accompanied by a transient disappearance of cp-actin filaments, unlike what occurred during the avoidance response (Kadota et al., 2009). Thus, the light-induced generation of biased cpactin filaments is a prerequisite for both the avoidance and the accumulation responses. Possibly, a transient disappearance of cp-actin filaments induced by strong blue light facilitated an acceleration of chloroplast avoidance movement. As more cp-actin filaments accumulated at the front halves of the chloroplasts, in relation to the rear halves, the velocity of chloroplast avoidance also increased. When irradiated with a higher fluence of blue light, even more cp-actin filaments accumulated at the front halves, and chloroplasts moved even faster (Kadota et al., 2009). Thus, strong light caused a greater difference in the amount of cp-actin filaments at certain locations on the chloroplasts because cp-actin filaments located at the rear halves of the chloroplasts did not increase after transient disappearance. Conversely, weak light could not induce transient disappearance of cp-actin filaments, so a greater difference in

Chloroplast Photorelocation Movement: A Sophisticated

**cp-actin filaments**

Strategy for Chloroplaststo Perform Efficient Photosynthesis 223

We identified various Arabidopsis mutants deficient in chloroplast photorelocation movement (Kagawa et al., 2001; Kodama et al., 2010; Oikawa et al., 2003; Suetsugu et al., 2005, 2010a), and thus, the analyses of cp-actin filament behavior in these mutants have shed light on the molecular mechanism of cp-actin filament-mediated chloroplast movement (Kadota et al., 2009; Kodama et al., 2010; Suetsugu et al., 2010a; Ichikawa et al., 2011)(Fig.3). Phototropin is a blue light receptor bearing two photosensory LOV (light, oxygen and voltage) domains at its N-terminus and a C-terminal serine/threonine kinase domain (reviewed by Christie, 2007). In *A. thaliana*, phot1 and phot2 redundantly mediated the chloroplast accumulation response (Sakai et al., 2001), and phot2 alone regulated the avoidance response (Kagawa et al., 2001; Jarillo et al., 2001). In *phot1phot2* double mutant plants, which are completely defective in chloroplast photorelocation movement (Sakai et al., 2001), blue-light-induced cp-actin filament reorganization did not occur, indicating that phototropins mediated chloroplast movement via the regulation of cp-actin filaments (Kadota et al., 2009; Ichikawa et al., 2011). The *phot1phot2* double mutant plants also did not change their amounts of cp-actin filaments in response to both weak and strong blue light and thus showed no light-induced motility changes (Kadota et al., 2009). This outcome indicated that phototropins mediated anchoring of the chloroplast to the plasma membrane via regulation of the amounts of cp-actin filaments. The strong-blue-light-induced transient disappearance of cp-actin filaments did not occurr at all in *phot2* mutant plants, which were impaired in the avoidance response. However, they showed normal biased cp-actin filament formation during the accumulation response (Kadota et al., 2009; Ichikawa et al., 2011), indicating that phot2 mediated the strong-blue-light-induced transient disappearance of cpactin filaments (Fig.3) and that this reorganization of cp-actin filaments could be a prerequisite for the avoidance response. In *phot1* mutant plants, chloroplast photorelocation movement was only slightly impaired in the accumulation response (Kagawa & Wada, 2000), and therefore light-induced reorganization of cp-actin filaments in these plants was mostly normal (Kadota et al., 2009). However, in response to strong blue light, the onset of biased cp-actin formation and the avoidance movement in *phot1* mutants occurred earlier than in wild-type plants (Ichikawa et al., 2011), suggesting a small inhibition of cp-actin

**2.2.2 Molecular components regulating the generation and/or reorganization of**

filament reorganization by phot1 during the avoidance movement.

JAC1 has a J-domain at the C-terminus and is similar to a clathrin uncoating factor, auxilin (Suetsugu et al., 2005). The J-domain of JAC1 is necessary for JAC1 function and the crystal structure showed high similarity between that domain and that of the bovine auxilin J-domain (Takano et al., 2010; Suetsugu et al., 2010c). *jac1* mutant plants were completely defective in the accumulation response but retain the avoidance response (Suetsugu et al., 2005). In response to weak blue light, the reorganization of cp-actin filaments did not occur in most chloroplasts of *jac1* mutant plants, but a few chloroplasts that avoided weak light formed biased cp-actin filaments (Ichikawa et al., 2011), indicating that JAC1 is essential for the reorganization of cp-actin filaments during the accumulation response but not for biased cp-actin filament formation (Fig. 3). Interestingly, in *jac1* mutant plants, whole cell irradiation with strong blue light did not induce the disappearance and subsequent biased localization of cp-actin filaments and thus the avoidance movement did not occur. However, when part of a cell was irradiated, chloroplasts that were close to the beam edge showed the avoidance movement with biased cp-actin filament formation, although cp-actin filaments on chloroplasts inside the

the amount of cp-actin filaments at certain locations was not made. Actually, the velocity of chloroplast accumulation movement was constant irrespective of light intensity (Kagawa & Wada, 1996; Tsuboi & Wada, 2011a). Cp-actin filaments localized at the interface between the chloroplast and the plasma membrane, elongated from the edge of the chloroplast and shortened toward the chloroplast periphery, suggesting that the nucleation site of cp-actin filaments might exist on the chloroplast edge and that the force for chloroplast movement by cp-actin filaments might be generated there (Kadota et al., 2009). Cp-actin filaments mediated the anchoring of chloroplasts to the plasma membrane as well as their directional movement (Kadota et al., 2009). The strong-light-induced disappearance of cp-actin filaments was accompanied by increased chloroplast motility in random directions before avoidance movement, suggestive of the detachment of chloroplasts from the plasma membrane. Conversely, weak blue light induced the increase of cp-actin filaments around the chloroplast periphery and accompanied a decrease in chloroplast motility, likely facilitating chloroplast anchoring to the plasma membrane. In summary, there are three types of blue-light-induced rearrangements of cp-actin filaments that mediate both directional movement and the anchoring of chloroplasts to the plasma membrane: (i) the formation of biased cp-actin filaments during both the accumulation and the avoidance responses; (ii) a strong-blue-lightinduced transient disappearance of cp-actin filaments; (iii) a weak-blue-light-induced increase in cp-actin filaments. Importantly, cp-actin filament-mediated chloroplast movement is conserved in a fern, *A. capillus-veneris*, and in a moss, *P. patens* (Tsuboi & Wada, 2011b; Yamashita et al., 2011). Thus, the regulation of chloroplast movement by cp-actin filaments was likely to be utilized during the early stages of land plant evolution.

Fig. 2. Light-induced cp-actin filament reorganization during the chloroplast avoidance response. The chloroplast avoidance response was induced by scanning circular regions of interest (diameter 15 µm indicated by a shaded circle) with 2.8 mW of a 458 nm laser and using a confocal microscope (SP5, Leica). Time-lapse images of chloroplast movement and the associated cp-actin filament dynamics were captured at the indicated times (sec). Detailed dynamics of the cp-actin filaments of a chloroplast marked with a white circle are indicated below. After 40 sec of light irradiation, cp-actin filaments disappeared and then reappeared at the front region of the chloroplast, which had moved to the upper left side in this figure via the avoidance response.

the amount of cp-actin filaments at certain locations was not made. Actually, the velocity of chloroplast accumulation movement was constant irrespective of light intensity (Kagawa & Wada, 1996; Tsuboi & Wada, 2011a). Cp-actin filaments localized at the interface between the chloroplast and the plasma membrane, elongated from the edge of the chloroplast and shortened toward the chloroplast periphery, suggesting that the nucleation site of cp-actin filaments might exist on the chloroplast edge and that the force for chloroplast movement by cp-actin filaments might be generated there (Kadota et al., 2009). Cp-actin filaments mediated the anchoring of chloroplasts to the plasma membrane as well as their directional movement (Kadota et al., 2009). The strong-light-induced disappearance of cp-actin filaments was accompanied by increased chloroplast motility in random directions before avoidance movement, suggestive of the detachment of chloroplasts from the plasma membrane. Conversely, weak blue light induced the increase of cp-actin filaments around the chloroplast periphery and accompanied a decrease in chloroplast motility, likely facilitating chloroplast anchoring to the plasma membrane. In summary, there are three types of blue-light-induced rearrangements of cp-actin filaments that mediate both directional movement and the anchoring of chloroplasts to the plasma membrane: (i) the formation of biased cp-actin filaments during both the accumulation and the avoidance responses; (ii) a strong-blue-lightinduced transient disappearance of cp-actin filaments; (iii) a weak-blue-light-induced increase in cp-actin filaments. Importantly, cp-actin filament-mediated chloroplast movement is conserved in a fern, *A. capillus-veneris*, and in a moss, *P. patens* (Tsuboi & Wada, 2011b; Yamashita et al., 2011). Thus, the regulation of chloroplast movement by cp-actin filaments

was likely to be utilized during the early stages of land plant evolution.