Transduction mechanisms of photoreceptor signals in plant cells

Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

Contents lists available at ScienceDirect

Journal of Photochemistry and Photobiology C: Photochemistry Reviews journal homepage: www.elsevier.com/locate/jphotochemrev

Review

Transduction mechanisms of photoreceptor signals in plant cells Vladimir D. Kreslavski a , Robert Carpentier b , Vyacheslav V. Klimov a , Suleyman I. Allakhverdiev a,∗ a b

Institute of Basic Biological Problems, Russian Academy of Sciences, Pushchino, Moscow Region 142290, Russia Groupe de Recherche en Biologie Végétale, Université du Québec à Trois-Rivières, C.P. 500, Québec, Canada G9A 5H7

a r t i c l e

i n f o

Article history: Received 13 January 2009 Received in revised form 28 April 2009 Accepted 30 April 2009 Available online 12 May 2009 Keywords: Ion exchange Phytochrome Plant Photoreceptors Signal transduction Transcriptional factors

a b s t r a c t During a plant life, light is necessary not only as a source of energy, but also as a regulatory factor of plant metabolism with information signal function. In this review we consider basic links of primary stages of light signal transduction in higher plants. The transformation circuits and possible pathways of photoreceptor light signal transduction, as well as possible roles of photoreceptor-interacting proteins, secondary messengers and some transcriptional factors are discussed. The review is also focused on examination of rapid signaling events such as activation of ion exchange systems as well as interaction of photoreceptors in signaling pathways. © 2009 Elsevier B.V. All rights reserved.

Contents 1. 2.

3. 4.

5. 6. 7. 8. 9.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Photoreceptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Plants possess multiple photoreceptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Types of phytochrome reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Cellular localization of photoreceptors: a key to understanding their functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Photoreceptors and systems of ion exchange . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Second messengers and effector proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Mechanisms of ion exchange and Ca2+ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Targets of calcium signals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Other messengers: cAMP (cGMP), IP3 and DAG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1. The role of G-proteins and effectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Phosphoinositide cycle and protein kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell structure and signal transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The regulation of gene activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Interaction of photoreceptor signaling pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nucleus, chloroplast and mitochondrial signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

64 66 66 67 68 69 70 70 71 72 72 73 73 74 75 75 77 77 77

Abbreviations: cAMP, adenosine-3 ,5 -cyclophosphate or 3 ,5 -cyclo-AMP; AMP, adenosine-5 -monophosphate; GMP, guanosine monophosphate; GTP, guanosine triphosphate; GC, guanylyl cyclase; IP3 , inositol-1,4,5-triphosphate; PIP2, phosphatidylinositol-4,5-diphosphate; phy, phytochrome; PDE, phosphodiesterase; RL, red light; FRL, far-red light. ∗ Corresponding author. Tel.: +7 496 77 31 837; fax: +7 496 73 30 532. E-mail addresses: [email protected] (V.D. Kreslavski), [email protected] (R. Carpentier), [email protected] (V.V. Klimov), [email protected] (S.I. Allakhverdiev). 1389-5567/$20.00 © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.jphotochemrev.2009.04.001

64

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80 Vladimir D. Kreslavski is a senior researcher. He is the head of group in Laboratory of Ecology and Physiology of phototrophic organisms at Institute of Basic Biological Problems, RAS, Pushchino, Moscow Region, Russia. The field of interests: photoreceptor signaling, molecular mechanisms of plant stress resistance and acclimation of photosynthetic apparatus as well as the pathways of photosynthetic improvement.

Robert Carpentier is professor at Université du Québec à Trois- Rivières, Québec, Canada. He obtained his Ph.D. in Biochemistry from Laval University (1983, Québec). He is editor of Journal of Photochemistry and Photobiology B: Biology, associate editor of the journal Photosynthesis Research and was the chair of the XIIIth International Congress on Photosynthesis (Montréal, 2004). His research interests concern the influence of environmental stresses on electron transport pathways in photosystems I and II and energy dissipation in photosynthesis.

Suleyman I. Allakhverdiev is the Chief Research Scientist at the Institute of Basic Biological Problems, RAS, Pushchino, Russia. He received Dr.Sci. degree in Photochemistry, Photobiology, and Plant Physiology (in 2002, Moscow); and Ph.D. in Physics and Mathematics, (in 1984, Pushchino). He graduated from Azerbaijan State University, Department of Physics (Baku). He has been guest-editor and is a member of the Editorial Board of several international journals. His research interests include the structure and function of photosystem II, water oxidizing complex, artificial photosynthesis, hydrogen photoproduction, catalytic conversion of solar energy, plant under environmental stresses, photoreceptor signaling. He has been cited ca. 3000.

1. Introduction Light is known to regulate almost all physiological and biochemical processes in plants. Together with photosynthesis, photomorphogenesis, phototaxis, phototropism, etc. play an important role in vital activity of different plant species. These processes are triggered by light and transformed into a cell response via a system of transduction of the light signal [1–13]. The mechanisms and components of these processes have not been adequately studied in plant cells. This is primarily true for the rather fast first stages of the light signal transduction chain, when light-induced proteins have no time to be synthesized. The term “signal transduction” became popular in the early 1980s, therefore the problem of light signal transduction in plant cells belongs to a fairly new area of biochemistry designated as “cell signaling”. In biology, signal transduction refers to any process by which a cell converts one kind of signal or stimulus into another. Most often, this involves ordered sequences of biochemical reactions inside the cell that are carried out by enzymes and linked through second messengers resulting in what is thought of as a “second messenger pathway”. The concept of “cell signaling” implies not only the signals’ transduction, but the entire set of events connected to it, including signal multiplication, depression, and suppression (or switching off) [4,8,11]. Such processes are usually rapid, lasting on the order of milliseconds in the case of ion flux, to minutes for the activation of protein and lipid mediated kinase cascades. Thus, sensing both external and internal environments at the cellular level relies on signal transduction. Signal transduction usually involves the binding of small extracellular signaling molecules to receptors that face outwards from the plasma membrane and trigger events inside the cell. Between them, steroids represent an example of extracellular signalling molecules that may cross the plasma membrane due to their lipophilic or hydrophobic nature [14].

Environmental stimuli may be both molecular in nature (as above) or more physical, such as light absorbed by plant photoreceptors. In this case light may affect photoreceptor molecules localized in the whole cell. In that case, inner membranes are most likely to be involved in the signal transduction chain, whereas the receptors of the hormones are mainly located on the plasma membrane. Various effector proteins (the effectors), for example adenylate cyclase (ADC) and GTP-binding proteins (G-proteins) are involved in the transduction chain from receptors to subsequent compartments of the cell. Activation of the effectors may be initiated by the detachment of the ␣-subunit of the heterotrimeric Gproteins. The activated G protein subunits can initiate the signaling for many downstream effector proteins, including phosphodiesterases and adenylyl cyclases, phospholipases, and ion channels that permit the release of second messenger molecules such as cyclic nucleotides (cyclic guanosine 3 ,5 -monophosphate—cGMP and cyclic adenosine-3 ,5 -monophosphate—cAMP), the components of the phosphatidylinositol signaling system (inositol 1,4,5-triphosphate–IP3 and 1,2-diacylglycerol—DAG), as well as Ca2+ [10,15–20]. The central position in the system of intracellular signalling is occupied by these three important messengers: Ca2+ , IP3 and DAG. The origin of the two last compounds merits special notice. They are formed from the plasma membrane component phosphatidyl-4,5-bisphosphate (PIP2 ) with the participation of phosphoinositide-specific phospholipase C. Several reactive oxygen species such as H2 O2 , O2 − , as well as NO, cADP-ribose and nicotinamide adenine dinucleotide phosphate also belong to the important family of secondary messengers. The increasing concentration of free cytosolic Ca2+ is the most widespread mechanism of transduction signaling chain independently from the nature of various signals. In this case Ca2+ -sensitive proteins, in particular calmodulin (CaM) can be the targets of free cytosolic calcium. A certain amount of evidence of the participation of Ca2+ , cAMP, cGMP, CaM, as well as G-proteins and the components of the phosphatidylinositol signaling system in the transduction of the phytochrome signal has been obtained [7,10,11,19–25]. The activated receptors located in the plasma membrane transmit the signal to the intracellular targets. If a target, or the effector protein, is an enzyme, then the signal modulates its catalytic activity. If an ion channel serves as effector protein, then the conductivity and lifetime of this channel is modulated. Nevertheless, it is important to note that a small quantity of hormonal molecules or light quanta, affecting the corresponding receptors, can produce a great number of messenger molecules activating synthesis of several proteins. The adenylate kinase system, catalyzing the formation of cellular cAMP, functions in such manner [19,26]. In that case a significant amplification of the signal is observed as a result of the interaction of an external signal with a receptor. Another mechanism of signal amplification involves regulation of the expression of light-controlled genes. A cell receptor involved in light signal transduction can interact with different cell components. Hence, several signal transduction pathways are possible. Second messengers such as Ca2+ and cyclic nucleotides are involved in the most widespread pathways of signal transduction (Fig. 1). Some receptors can also activate directly (without involvement of second messengers) protein kinases, for example the enzyme tyrosine kinase, which phosphorylates the residues of tyrosine in the proteins [27]. In this case a cascade of cytosolic protein kinases is triggered. They phosphorylate various proteins that cause subsequent physiological effects. Regulation of gene expression can result from the transduction of signals of various natures. The signal can be transduced into the cell nucleus by translocation of cytosolic protein kinases or activating transcription factors. Transcription factors produced

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

65

Fig. 1. Schematic diagram of primary stages of transduction of the light signaling in plant cells (modified scheme from [11]). ABPs, actine-binding proteins; AMP, adenosine5 -monophosphate; cAMP, 3 ,5 -cyclo-AMP; ADC, adenylate cyclase; DAG, diacylglycerin, IP3 , inositol-1,4,5-triphosphate; CaM, calmodulin; EPs, effector proteins; PIP2, phosphatidylinositol-4,5-diphosphate; PK, a protein kinase; SIE, the systems of ion exchange in the cytoplasmic membrane; PDE, phosphodiesterase; PLC, phospholipase C; PKC, protein kinase C; PKA, protein kinase A; P, photoreceptor; P*, an active form of the photoreceptor; ER, endoplasmic reticulum; EC, the elements of cytoskeleton; X and Y, the unknown elements.

as a result of a signal transduction cascade can in turn activate yet more genes [13]. Therefore an initial stimulus can trigger the expression of an entire cohort of genes leading to the activation of any number of complex physiological events. Photophosphorylation of proteins plays a key role in this process. Phosphorylation of nuclear proteins was proposed to be involved in phytochrome signal transduction [28]. It has been shown that light activates the phytochrome, which penetrates into the nucleus to interact with nuclear proteins such as the phytochrome interacting factor 3, PIF3 [29,30]. Red light promotes the formation of the Pfr-active form of phytochrome, which translocates into the nucleus. Pfr induces the rapid PIF3 phosphorylation prior to proteasome-mediated degradation [31,32]. PIF3 induces the expression of skotomorphogenesis genes, thereby acting as a negative regulator of photomorphogenesis [13]. Like phytochromes, the cryptochromes cry1 and cry2 appear to be localized to the nucleus following illumination [31]. Thus, the phytochrome is able to regulate the transcription of nuclear genes directly involved in growth reactions [3,8,13,33]. PhyA and phyB can bind a rather large number of proteins localized

in the cytoplasm and the nucleus that participate in the transduction of the phytochrome signal [2,3,8,13,33]. A group of PIF transcription factors interact directly with phytochromes and function mainly as repressors of photomorphogenesis. Phytochrome interacting factors such as PIF3, PIF4, PIF5 (PIL6) and PIF6 interact mainly with phyB, whereas PIF1 (PIL5) can bind to both phyA and phyB [13,29,34–36]. It is important to note that transcription factors, which are downstream of the light-signaling pathways, are regulated at the level of protein degradation by COP1 and other proteins (for a review see [37], Fig. 2). Thus, photomorphogenesis is also governed by proteolysis. Some similarities are supposed to exist between the pathways and mechanisms of transduction of various signals that can be realized via G-proteins, cyclic nucleotides (cAMP, cGMP), Ca2+ , Ca2+ binding proteins and protein kinases, as well as the components of the phosphatidylinositol signaling system and/or the elements of the cytoskeleton [7–11,16,18,19,22,24,26,38]. We have attempted to examine the pathways of light signal transduction in plant cells. The major elements that are considered will include ion exchange systems and messengers and effector proteins of the primary stages of

66

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

Fig. 2. A model of phytochrome signal transduction. In response to absorption of RL, the phytochrome in the form of Pr transfers into an active form Pfr, the formation of which triggers migration of the greater part of Pfr into the nucleus. After formation of Pfr, the phytochrome is phosphorylated (marked as P), as well as can be bond to certain cytoplasmic proteins like SPKA (substrate of phytochrome kinase activity) and transcription factors like CPRF1, CPRF2 and GBF1, where both CPRF1 and CPRF2 are members of the common promoter-binding transcription factor family (CPRF) and GBF1 is G-box binding factor 1. Phosphorylation also leads to migration of Pfr and subsequent proteins into the nucleus. Formation of Pfr is also followed by activation of heterotrimeric G-proteins by an unknown mechanism and leads to changes of the content of cytoplasmic cGMP and Ca2+ with calmodulin (CaM). This leads to activation of transcription factors (X and Y) belonging to transcription complexes required for light-dependent expression of various genes. Pfr is proposed to regulate transcription through several parallel pathways. A rapid response involves Pfr translocation to the nucleus where it binds transcription factors of the bHLH family (in particular PIF1, PIF3; PIF4; PIF5; PIF6, PIF7 [13,194]. Key regulatory transcription factors (RTFs) that are responsible for inducing a range of light-regulated genes are subsequently activated. In a second nuclear-localized pathway, the phytochromes are proposed to bind response regulators (RR), which stabilize them in the activated form and can induce light-regulated gene expression by inhibiting COP1-, COP10- and CSN-dependent proteolysis of the HY5 transcription factor and by binding to activated cryptochromes (cry). X, Y and Z, the unknown elements. In the cytoplasm, the phytochromes may activate gene expression through G-proteins (G), calcium and cGMP-dependent pathways, which are regulated by cytoplasmic protein SUB1. In addition, the phytochromes may be sequestered away from the signalling-competent pool by PKS1. Elements involved in signalling from specific photoreceptors or controlling specific responses have not been included. The presented scheme with modifications was taken from [3,33]. For more details see [11,13,40,195,196].

such signal transduction chain as presented on schematic diagrams (see Figs. 1 and 2). 2. Photoreceptors 2.1. Plants possess multiple photoreceptors Light is one of the most important environmental factors both providing the source of energy for plants and controling all stages of plant development, from seeds germination up to the final stages of the life cycle. Seed germination, growth and inhibition of hypocotyls, cotyledon expansion, chloroplast development, plant architecture, cell disposition in a leaf, and ion exchange in cells are all light-regulated processes [8,12,13,39]. At least three families of photoreceptors are supposed to take part in the response of plants to light: phytochromes, blue-light receptors and UV-B photoreceptor(s) [5,8,12,13,39,40]. Comprehensive up-to-date overviews of plant photoreceptors are provided

[8,12]. In Arabidopsis there are five phytochromes and several bluelight photoreceptors, but this list can be extended [8,41–43]. For example, Folta [42] suggested the availability of a novel green lightactivated sensor, which promotes early stem elongation in etiolated Arabidopsis seedlings. High-resolution analyses of early growth kinetics has shown that irradiation of plants with green light causes a rapid increase in early stem elongation rate, a response that is contrary to that induced by all other light conditions studied. According to the data available, the transient growth promotion could not be completely attributed genetically or photophysiologically to the action of known photoreceptors. Interestingly, supplemental green light antagonized long-term blue and red light effects on stem growth. Phytochromes, chromoproteins with a molecular weight of about 240 kDa, are homodimeric complexes. Each polypeptide contains a N-terminal chromophore-binding domain (CBD) that autocatalytically attaches, via a thioether linkage, a single linear tetrapyrrole (or bilin) chromophore, a PHY domain that is

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

Fig. 3. Absorption spectra of phytochrome in 50 mM Tris–C1, 25% (v/v) ethylene glycol buffer pH 7.8 at 4 ◦ C (c = 5.0 × 10−6 M; 1-cm path length). (–) Pr ; (- - -) Pfr . Pphytochrome was isolated from etiolated Avena sativa L. seedlings. Adapted from [197].

important for spectral integrity, and a C-terminal domain that promotes dimerization and often signal transmission [44]. The ability of phytochromes to sense the light environment is due to the covalently attached tetrapyrrole chromophore (phytochromobilin). The photoreceptor exists in two photoconvertible conformations, denoted red light-absorbing (Pr form) and far-red light-absorbing (Pfr form). Photoconvertion between Pr and Pfr constitutes a unique light-regulated molecular switch that operates in various signal transduction cascades. Phytochromes are thus photoswitchable photosensors; canonical phytochromes have a conserved N-terminal photosensory core and a C-terminal regulatory region, which typically includes a histidine-kinase-related domain [45] (for a detailed description of phytochrome structure see the review of Rockwell et al. [44]). There are five types of phytochromes (A, B, C, D and E) in Arabidopsis thaliana [13,46,47]. The physiological functions of phyA and phyB are the most characterized ones and only recently has some information been obtained about the roles of phyC–E [48]. As mentioned, each type of phytochrome can exist in two major relatively stable photochromic forms: with m = 660 nm (Pr) or m = 730 nm (Pfr) in the region of visible spectrum and several other relatively unstable forms [47,49] (Fig. 3). It should be stressed that the initially inactive form Pr is synthesized in the dark, at absorption of light quantum with m = 660 nm this form converts into the physiologically active form, Pfr [40,50]. Absorption of m = 730 nm light converts the active form into Pr again. PhyA seems to be the primary photoreceptor for mediating farred light regulation of gene expression [51]. PhyA is prevalent in etiolated seedlings grown in the dark and very photolabile. This form of the phytochrome is easily exposed to proteolytical degradation following transition into an active form, whereas other types of phytochrome, in particular phyB, are relatively stable in the light. Since phyA degrades in fact completely in sunlight, phyB is dominant under these conditions. During deetiolation, continuous far-red light (FRL) affects only via phyA, whereas red light (RL) has an influence mainly via phyB. Note that the later becomes a major photoreceptor during seedling deetiolation. Photoreceptors similar to the phytochromes of higher plants, including two photoreversibile states, were also found in green algae and cyanobacteria [52–54]. One phytochrome-like protein has been characterized in Synechococcus 7942 [55] and other possible phytochromes were detected in the Synechococcus 7942 genome sequence (U.S. Department of Energy Joint Genome Institute: http://genome.jgi-sf.org/draft microbes/synel/synel.home.html). However, the genome of the non-sulphur purple bacterium Rhodopseudomonas palustris is unusual in containing six bacteriophytochromes (Bphs), compared with only one or two Bphs in other bacterial genomes sequenced to date. It even exceeds the five phytochromes found in the model plant species Arabidopsis thaliana. It is known that Bph3 is active in its red absorbing Pr form and a working hypothesis suggests that Bph4 is active in the Pfr state [56]. The phytochromes of the cyanobacterium Synechocystis have sites, which are highly homologous to the phytochromes of higher

67

plants. Their C-terminal homologous amino acid sequence can be considered as sensor hystidine kinase [54]. In vitro it phosphorylates the cytoplasmic protein PKS1 (phytochrome kinase substrate), the substrate of phytochrome kinase activity [57]. PhyA works as untypical serine-treonine kinase, which phosphorylates the protein PKS1 in a light-dependent way. Along with phytochromes, another type of photoreceptors, blue-light receptors, such as cryptochromes [58], phototropins [59,60] and FKF1 family members, were also found [61]. Arabidopsis contains two phototropins, phot1 and phot2, that exhibit overlapping functions in addition to having unique physiological roles, and two cryptochromes (cry1 and cry2) [60,61]. Several forms of cryptochrome have been found at present that slightly differ in composition and have absorption maxima within the range of wavelengths up to 500 nm. These photoreceptors are involved in light signal transduction regulating phototropism, chloroplast movements and light-induced stomatal opening (for review see [43,60]). Little is known about the UV-B photoreceptor(s) [62]. However, recently two signaling components required for normal UV-B responses have been identified [8]. A “chimeric” photoreceptor, homologous both to phytochrome and phototropin (NPH1), has been isolated from the fern Adiantum [63]. The N-terminal site of this photoreceptor is similar to the phytochrome N-terminal, and the C-terminal is similar to phototropin. In this case, red and blue light can effect on phototropism via one and the same photoreceptor. More recently, this photoreceptor was investigated in more detail and named Phy3 [64]. Since Phy3 greatly enhances the sensitivity of leaves and chloroplasts orientation to white light and Phy3 homologues exist between different fern species, in the author’s opinion, it is likely to play an important role in the divergence and proliferation of fern species under lowlight conditions. An analogous photoreceptor and its “chimeric” genes (neochrome) have been recently found in the green algae Mougeotia scalaris [65]. 2.2. Types of phytochrome reactions Many reactions controlled by phytochromes are saturated at rather low intensity of red light (m = 660 nm). These reactions are characterized by the reversibility of the short-term RL effect by subsequent application of short-term FRL (m = 730 nm), which can convert Pfr back into the inactive Pr form (Fig. 1; Fig. 4A). Thus, red/far-red reversibility and reciprocity are the hallmarks of the classic phytochrome responses. This class of phytochrome responses, known as low fluence responses (LFR), was already studied in 1952 [66] and has been described in many different plant systems [6,40] (Fig. 4B). The last figure demonstrates the processes of transformation of two phytochrome forms (A) and the different phytochrome response modes (B). Leaf and stem growth, gene expression during etiolation, seed germination, leaf movement, and rotation of the chloroplasts in the green algae Mougeotia belong to LFRs [6,40,67]. The growth and development of the first leaf and coleoptile in wheat seedlings studied by one of us are also among these reactions [68], as well as the release of ethylene in the green algae Chlorella [69]. Besides LFRs, verylow-fluence responses (VLFRs) without any RL/FRL reversibility and high-irradiance responses (HIRs) were also reported [3,6,49]. Studies with phytochrome-deficient mutants grown in defined VLFR, LFR and HIR light environments have revealed that specific phytochromes can be ascribed to individual responses (Fig. 5B), although significant redundancy exists [3,48]. These three response modes (Fig. 5) can be distinguished by the amount of light required, which varies over eight orders of magnitude. In a typical VLFR, plants respond between 0.1 and 1 ␮mol/m2 of light, whereas the LFRs are typically between 1 and 1000 ␮mol/m2 of light. HIRs are now further subdivided into red

68

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

Fig. 4. Phytochrome response modes. (A) The phytochrome photocycle. Pr and Pfr denote the red and far-red light-absorbing conformations of phytochrome, respectively, which are reversible depending on light conditions. Pfr can also be converted to Pr in a light-independent process known as dark reversion. (B) The different phytochrome response modes. The influence of red and far-red light on each response mode is shown, together with the phytochrome principally involved in initiating the response. Adapted from [3].

(R)- and far-red (FR)-HIRs [3]. It should be also stressed that HIRs are observed during irradiation with far-red light and VLFRs are controlled by phyA [3,6]. This is in line with the fact that the bestknown mutant, the goodish tomato mutant (aurea, au), deficient in phyA (not in phyB) [70] is insensitive to far-red light responsible for HIRs [6]. 2.3. Cellular localization of photoreceptors: a key to understanding their functions Localization of photoreceptors is of current interest to study light signal transduction in plant cells. According to data obtained with narrow microbeams of red and far-red light [71,72], at least a fraction of the phytochrome appeared to be bound to the cytoplasmic membranes. However, with the help of biochemical methods it has been ascertained further that phytochromes are mainly localized in the cytoplasm, though a fraction can be in organelles—mitochondrions, chloroplasts, or nucleus [3,48,73].

Fig. 5. Effect of low intensity red light (m = 660 nm) and following far-red light (m = 730 nm) on ethylene production in green alga Chlorella. Initial cell density (no ) = 70.5 × 106 cells ml−1 . Adapted from [69].

For the purpose of studying phytochrome localization in intact tissues, transgenic Arabidopsis plants expressing fusion proteins consisting of phyB and other types of phytochrome with the green fluorescent protein (GFP) were obtained [74]. Their physiological activity remained practically the same relative to that in tissues with common phytochrome. The intracellular localization of these “specific” photoreceptors was studied. Fluorescent microscopy and immunological analysis confirmed that in tissues growing under illumination, the phytochromes were mainly localized in the nucleus, and in dark seedlings, in the cytosol. The early results obtained with cells of the filamentous algae Mougeotia [67,75] suggested that the phytochromes in the cytosol were bound with elements of the cytoskeleton. This is in agreement with the amino acid sequence of the C-terminal phytochrome domain being similar to the amino acid sequence of proteins bound with microtubules [76]. The early experiments with plane-polarized red light showed that Pr was directed parallel to the long cell axis, and Pfr was directed perpendicular [67,75] that is in line with the idea of phytochrome ‘association” to the cytoskeleton. A fraction of the phytochrome molecules is likely bound with the cytoskeleton near to the plasma membrane [67,77]. Experiments with different types of phytochromes were carried out under various conditions using different plant species expressing protein fusion complexes of phytochrome and GFP, in particular in transgenic Arabidopsis plants [74] and tobacco [78]. In these experiments both fluorescent spectroscopy and immunological analysis with antibodies to phytochrome and fluorescent protein were used. Red light irradiation of tobacco seedlings grown in dark resulted in shifting of the complex phyB:GFP (but not the phyA:GFP) into the cell nuclei and this effect was reversible by farred light revealing the red light/far-red light reversibility typical for LFRs [78]. Hence, only far-red light stimulated the translocation of the phyA:GFP complex into the nuclei of tobacco and Arabidopsis cells [79] that indicated the possible involvement of the low fluence response of the phytochrome. In Arabidopsis, translocation of the phyA:GFP complex into the nuclei under continuous light was maximal after 2 h. Rather long-term irradiation with red light resulted in marked accumulation of phyB:GFP in the cell nuclei and development (maximal after 6–8 h) of new structures with clearly visible green spots due to GFP fluorescence [74,78]. In many cases, formation of these spots represented an integral functional feature of physiologically active phytochrome [80]. It was induced by the short-term red light accumulation of phyB:GFP in tobacco cell nuclei. This accumulation was released by far-red light demonstrating the red light/far-red light reversibility belonging to phytochrome-controlled LFRs [78,79]. It is known that phytochrome translocation is rather slow, except for phyA, and that the majority of the intracellular Pfr pool is not translocated to the nucleus [3,33]. Thus, phytochrome translocation depends on its state and type and is controlled by light: in dark it is in inactive form and localized mainly in the cytosol, whereas after light activation it migrates into the nucleus [81]. These and other observations (see below) suggest that phytochromes may activate signalling pathways in both the cytoplasm and the nucleus. Cryptochromes, like phytochromes, were found in every compartment of the plant cells. The main functions of different photoreceptors have been studied using photoreceptor mutants of Arabidopsis and other plants [2,6,13,82]. The results of all these studies suggest heterogeneity of phytochromes and differences in their localization. For example, one and the same form of phytochrome (A, B, C, D, E) can be localized in different cell compartments [74]. Moreover, its localization also depends on light and other physiological conditions, the form of phytochrome (Pr or Pfr), type of tissue, and the type of proteins bound to the phytochrome. The same refers to the cryptochromes. These receptors are ideally suited to detect the quantity and quality of light [2,4,5,60]. Experiements

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

studying the light-dependent rotation of chloroplasts in the cells of Mougeotia showed that the blue light receptors are mainly involved in cell response to light intensity, whereas direction of the incident light is monitored by phytochromes [67]. Nevertheless, many processes controlled by phytochromes are regulated by the Pr/Pfr ratio and subsequently by the RL/FRL ratio [2,83]. Thus, the RL/FRL ratio is a good indicator of time and place [2]. Light-regulated protein phosphorylation is often involved in photoreceptor signaling (Fig. 2). For example, the protein CPRF2 from parsley is localized in the cytosol in the dark and treatment with light causes an import in the nucleus [84]. Light-dependent in vivo phosphorylation of CPRF2 is probably the key event that triggers its nuclear import [85]. However, association between the kinase activity of a photoreceptor and a system of transduction has not been specified [13,40,43]. Finally, although we have a good understanding of how plants distinguish differences in light quality, it is still unclear how plants distinguish differences in light quantity, duration and direction. Such regulation could be at both the photoreceptor level and at the level of downstream signalling networks. Resolving these questions requires the identification of key steps in signalling under specific conditions through precisely designed genetic and genome-wide transcriptomic studies. 3. Photoreceptors and systems of ion exchange Whereas it is clear that phytochromes can regulate transcription events, it has long been known that many responses both in lower and higher plants (reviewed in [77]) are too fast to be explained in this way. Protoplast swelling [86–88], light-dependent rotation of chloroplasts in the cells of Mougeotia, which leads to the orientation of the chloroplast perpendicular to the incident beam of red light [67,89], change of membrane potential [18,90,91], mobility of the cytoplasm in cells of Vallisneria [92] belong to such fast processes. Phototropins control fast reactions as well: phototropism, chloroplast photorelocation movements, stomatal opening [5,43,60], and the level of free Ca2+ in the cytoplasm [93]. This is evidenced by many observations, for example, the transmembrane potential in Salmanea pulvini cells responds to the formation of Pfr within 2 min [94]. The first fast phase of inhibition of Arabidopsis seedlings, which is controlled by a UV-A/blue photoreceptor is mediated by an unknown UVA/blue-specific photoreceptor after a lag time of approximately 30 s [95]. The mobility of cytoplasm in the cells of Vallisneria has been stimulated markedly even after 2.5 s upon formation of Pfr [92]. This is in line with the study of Mittmann et al. [77], which revealed direct actions of phytochrome in the cytoplasm of Physcomitrella cells. An example of type of rapid response is an enhancement of ethylene production induced by RL impulse. Ethylene production in Chlorella grown at green light was quickly enhanced by low intensity short-time RL and abolished by FRL (Fig. 5). This evidences that ethylene production belongs to LFRs. For details see [69]. Ion channels are one of the most efficient ways of redistribution of the ion flows. They play an important role in the mechanism of intracellular signaling both in animal and plant cells [10,96–99]. Several types of channels are designated depending on their opening mechanism: receptor-opened ion channels, voltage-dependent ion channels, ligand-gated channels [10,98]. The opening of channels following membrane depolarization constitutes a fairly universal mechanism [18]. It is known that blue and red light as well as hormone signalling can promote the opening of Ca2+ -permeable slow vacuolar channels due to rapid depolarization of vacuolar membranes [10,98]. Early observations have demonstrated that large plasma-membrane depolarization precedes rapid blue-light-induced growth inhibition in cucumber [100]. Depo-

69

larization of the cytoplasmic membrane induced by blue light probably via a cryptochrome has been found in hypocotyls of Arabidopsis [95,101]. It was concluded that activation of anion channels (Cl− ) contributed to membrane depolarization. A Ca2+ -dependent depolarization of the plasma membrane induced by phytochromes was found in the cells of moss protonema Physcomitrella patens as well [90,102]. Nevertheless, the role of ion fluxes and membrane depolarization in phytochrome or blue-light photoreceptor control of long-time processes such as growth or morphogenetic responses remains elusive. Both the channels activated by the products of the phosphoinositide cycle IP3 [103] and cAMP-gated Ca2+ channels [104], are among ligand-gated Ca2+ channels of the tonoplast. The activity of ion channels can be regulated as well by the interaction with elements of the cytoskeleton, by phosphorylation and dephosphorylation and many other factors, including G-proteins, Ca2+ -binding proteins, etc. [99,105,106]. In principle, a photoreceptor can regulate membrane permeability without secondary mediators via modulation of the activity of ion channels [27]. This can be realized in several ways, either as a primary event or via some effector molecules. One cannot exclude that not only ligand-gated but also photo-gated channels, i.e. controlled directly via photoreceptor conformation changes, are available in plant cells. Moreover, the process can be rather quick, as the conformation of a photoreceptor can be changed in a short period of time. The activity of a channel depends on the conformational state of the protein forming the channel, which can be modulated by the change in the state of a photoreceptor. The concept viewing the functioning of the receptors, the “gates” of ion channels and membranes, as allosteric proteins having several conformations is progressing actively [11]. Finally, photo-potential-gated channels can be found, that are channels opened both by photoreceptor activation and by membrane potential changes. The assumption that photo-gated channels may involve a direct effect of photoreceptors on membrane permeability is acknowledged in a number of works. For example, early experiments on artificial bilayer phospholipid membranes showed that exogenic phytochrome could change the conductivity of the membranes in relation to its form (Pr or Pfr) [107]. It has been also studied how RL and RL with FRL affect the membrane potential of Characeae cells [108]. Depolarization due to RL was significantly lower in the presence of FRL; the decrease was maximal at low intensity of RL (about 50%). Reversibility of RL effect with additional FRL was maximal at low intensity of RL that testified in favor of the fact that membrane depolarization was at least partially regulated by phytochromes and not only by photosynthesis [108]. It has been also shown in some cells that light-induced depolarization has a lagphase of 0.4 s. In the cells of moss protonema Physcomitrella patens, RL caused a short-term (within 2–15 s) phytochrome-dependent electric depolarization of the cytoplasma membrane, which was abolished by cation (tetraethylammonium, TEA) and anion channel blockers (niflumic acid) [90]. At maximal depolarization, both cation (Ca2+ and K+ ) and anion channels were opened. Such short period of depolarization means that phytochromes can act directly on ion exchange components: channels, transporters, exchangers and pumps. This agrees well with the fact that a fraction of the phytochrome is likely to be bound to the plasma membrane [71,72,109]. Membrane-bound phytochrome acts most probably via G-proteins with further phosphorylation that may lead to opening of the ion channels [106]. However, a phytochrome-specific G-protein has not been found yet [27]. Recent studies using recombinant DNA technology have added much convincing evidence to the longstanding view of phytochromes as light-regulated kinases that phosphorylate ion channels to stimulate their opening [6,110,111]. Nevertheless, we need additional information to make final conclusions on the

70

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

mechanisms connecting the phytochrome with the systems of ion exchange as primary components of light signaling transduction chains. One of the pathways regulating the activity of ion channels may be the increase of concentration of free calcium ions, of components of phosphatidylinositol exchange and of a number of other activators of ion channels in the cytoplasm [18,105,112]. K+ channels may also participate in light signal transduction [96]. It should be noted that Ca2+ -activated potassium channels were found in the cytoplasmic membrane of the protoplasts of Mougeotia. They seem to regulate the flow of K+ ions in relation to the state of the phytochrome [113]. The opening of these channels can be triggered by the accumulation of cytosolic Ca2+ . Indeed, in the presence of the calcium channel antagonist 3,4,5-trimethoxybenzoic acid 8(diethylamino)octyl ester (TMB-8), thus retarding accumulation of Ca2+ , an activation of the channels with RL was not observed. The process of Ca2+ accumulation can be induced by the phytochromes either directly or via the secondary messenger IP3 [113], initiating an increase in the concentration of free Ca2+ in the cytosol [114]. There is evidence that the introduction of IP3 into the cells can activate Ca2+ channels of the tonoplast, inducing the release of calcium ions from vesicles and vacuoles [15,103,115]. This mechanism was supposed to take part in activation of Ca2+ -sensitive K+ -permeable channels of plasma membrane of the green alga Eremosphaera virdis [114]. An alternative to the mechanism of increasing [Ca2+ ]cyt could be the system of transduction described in [33] where it is supposed that formation of Pfr can be accompanied by the activation of heterotrimeric G-proteins leading to changes of the content of [cGMP]cyt and [Ca2+ ]cyt (Fig. 2). Let us note that approximately twenty genes coding ion channels that are gated by cyclic nucleotides were found in the Arabidopsis genome [112,116]. Some of these cyclic nucleotide-gated ion channels have domains for binding both with cAMP and cGMP and with calmodulin that may be important for various transduction pathways [112,117]. G-proteins are known to participate in the chain of light signals transduction [25]. Several observations are in the line with this concept. Okamoto et al. [24] designed a transgenic approach to examine whether heterotrimeric G-proteins are involved in the phytochrome signal transduction pathway. Arabidopsis is known to have a single gene encoding a canonical alpha subunit of a heterotrimeric G-protein (GPA1; [51]) and a single beta subunit (AGB1). They have generated transgenic Arabidopsis plants that are inducible overexpressors of the heterotrimeric G-protein, AtGPA1. The transgenes were placed under a glucocorticoid-inducible promoter [118]. This study showed that overexpression of the heterotrimeric G-protein ␣-subunit enhanced phytochrome-mediated inhibition of hypocotyl elongation in Arabidopsis and provided physiological evidence of the involvement of the heterotrimeric G-protein in light-regulated seedling development. G-proteins seem to influence the activity of ion channels. Aharon et al. [119] introduced an exogenic ␣-subunit of G-protein into tomato cells and found activation of the Ca2+ channels in plasmalemma membranes. Hence, it follows that G-proteins can participate in the regulation of the activity of Ca2+ channels. Ca2+ based signaling appears to be a common if not universal mechanism in plant G-protein responses [25]. However, a recent study [120] reexamined the role of the single canonical heterotrimeric G protein in R and FR control of hypocotyl growth using a loss-of-function approach. Single- and double-null mutants for the GPA1, AGB1 genes encoding the alpha and beta subunit of the heterotrimeric G protein, respectively, had wild-type sensitivity to R and FR. Ectopic overexpression of wild type with a constitutive active form of the alpha subunit and of the wild-type beta subunit had no effect that can be unequivocally attributed to altered R and FR responsiveness. These results, in author’s opinion, preclude a direct role for the het-

erotrimeric G complex in R and FR signal transduction in Arabidopsis participating in the control of the hypocotyl growth. Hence, future studies are required in this field for further comprehension of the mechanism of G-protein participation in light signal transduction. In addition, it should be marked that many light sensitive processes that involve phytochromes and cryptochromes occur within several seconds to minutes. They are associated most likely with the regulation of ion channels and pumps, and accordingly, with redistribution of ions’ flows, i.e. with changes at the membrane level. However, the relationship between these fast processes, activation of transcription and following growth and photomorphogenetic processes is often unclear. 4. Second messengers and effector proteins Various effector-proteins and some small molecules, which serve as secondary messengers modulating enzymatic activity, ion exchange and other cell functions, are involved in the transduction chains of various signals. The peculiarity of a second messenger is its relatively small molecular mass, quick degradation and in the case of calcium, fast removal of excess cytosolic calcium (a disturbance of the removing mechanism can lead to cell intoxication). The most important secondary messengers are cAMP, cGMP, IP3 , DAG and Ca2+ . The important role of secondary messengers in cell signaling was first determined in animal cells [121,122]. However, the role of secondary messengers in plants is now well studied. It has been shown, for example, that the Ca2+ -calmodulin system participates in the transduction of phytochrome signals [1] and is involved in the role of IP3 [15] as well as cAMP and cGMP [23] in increasing free Ca2+ . Three pathways of light signal transduction that involve cAMP and/or Ca2+ have been postulated [7,21,38]. An example of the first pathway is the expression of chalconsynthetase (chs gene) and biosynthesis of cAMP- regulated anthocyanins. The regulation of Ca2+ -dependent synthesis of chlorophyll a/b-binding protein (cab gene) constitutes a good example of the second pathway. The third pathway involves the participation of both cAMP and Ca2+ [21] as the case for the activation of gene expression for the enzyme ferredoxin – NADP – oxidoreductase (fnr gene). 4.1. Mechanisms of ion exchange and Ca2+ Calcium ion is one of the key secondary messengers in the transduction chains of various signals both in animal and plant cells [10,17,18,99,106]. The concentration gradient of calcium ions between the intracellular and extracellular compartments is 3–4 orders exceeding the gradients of other ions. In addition, calcium, like magnesium, is able to form several coordination links (six to eight) that are important for binding with the targets. The flow of calcium ions is regulated by Ca2+ channels, calcium exchangers and pumps, as well as by the level of Ca2+ -sensitive proteins. Ca2+ channels provide entering of calcium according to the concentration gradient; pumps and exchangers (or carriers) work against the concentration gradient using the ATP energy. Ca2+ -permeable channels were found in the plasma membrane, tonoplast, endoplasmic reticulum, chloroplasts, and nuclear membranes of plant cells [10,98]. Usually two major types of Ca2+ channles are considered in the plasma membranes: rather selective (voltage-dependent cation channel) and weakly selective (maxication channel) [17]. It has been found that voltage-dependent Ca2+ -permeable channels are activated by hyperpolarization and depolarization, IP3 , cAMP, or cADP-ribose [98,103–105]. The activity of Ca2+ channels can be suppressed by direct or indirect interaction with microtubes [123] that emphasizes the importance of cytoskeleton elements in the regulation of the activity of ion channels.

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

It should be stressed that the coordinated function of passive and active transport of Ca2+ is required to provide a so called shortterm increase of Ca2+ concentration, i.e. the ability of cytoplasmic calcium to return to the basal level regardless of whether the signal that caused entering of Ca2+ into the cell is still acting. For this purpose, an active ejection of excess calcium from the cytoplasm by Ca2+ -pumps and exchangers is required. Biochemical experiments using inhibitors of calcium channels, Ca2+ -ionophores and regulation of the level of external calcium in the medium have shown the participation of calcium ions in many fast phytochrome-controlled processes [1,21,124,125]. Thus, blocking Ca2+ channels or removal of extracellular Ca2+ resulted in inhibition of phytochrome-dependent effects. Conversely, introduction of Ca2+ ionophores stimulated these effects. In particular, treatment with Ca2+ -ionophores such as A23187 mimics the effect of RL, activating the phytochrome. Both, irradiation with RL, or local treatment of the cell with the Ca2+ -ionophore A23187 in the presence of Ca2+ lead to rotation of the chloroplast in Mougeotia [126]. In contrast, in the presence of calcium channel blockers, verapamil or La3+ , the phytochrome-dependent effects were not revealed. Similar effects of Ca2+ -ionophore were observed in wheat protoplasts [124]. It has been also established that the accumulation of Ca2+ induced with RL in the cytoplasm is released following irradiation with FRL, i.e. it relates to low fluence phytochrome-controlled responses. The studies using activators and inhibitors of cytosolic calcium and Ca2+ channels have received further development. It has been shown [127] that both RL and blue light result in decreasing pH (acidification) in epidermis segments obtained from leaves of mutant pea plants: the blockers of Ca2+ channels inhibited acidification and the Ca2+ ionophores, ionomycin and A23187, reduced the segment responses to light. Moreover, the antagonists of CaM-W-7 and trifluoroperazine (TFP) inhibited completely the light-dependent acidification, but not in the case of blue light where the inhibition was made up to 60–70% only. The above indicates that not only Ca2+ but also Ca2+ out can be involved in the chain of in light signal transduction. On the other hand, there are a number of contradictory reports where the participation of calcium ions and Ca2+ channels in the chain of light signal transduction has not been demonstrated. As example, no marked effect of the blockers of Ca2+ entrance (ruthenium red and La3+ ) on chloroplast rotation induced by low- and high-energy illumination and their adhesion has been revealed [67]. These data along with some evidence of the influence of phytochrome on the activity of ion channels in protoplasts where the potential was locally fixed (patch-clamp experiments) [113], are in agreement with the conclusion on non-involvement of calcium channels in rotation of Mougeotia chloroplasts [67]. Shrinking of Arabidopsis protoplasts induced by blue light pulses did not depend on the concentration of external Ca2+ either [128]. Calcium ions also serve as secondary messengers in reactions controlled by blue light receptors, for example, in the cryptochrome-dependent process of stomata closing [5]. In experiments using the fluorescent dye indo-1, which indicates [Ca2+ ], rather quick (3–6 s) increases of Ca2+ concentration were observed in cells of Mougeotia after their illumination with UV-A and blue light at high irradiances, causing orientation movements of the chloroplasts [129]. Also, in a mechanism that involves phototropins (phot 1, phot 2), blue light induced a short-term increase of cytosolic Ca2+ in the leaves of Arabidopsis [93]. Thus, an increase of the level of [Ca2+ ]cyt is, obviously, one of the leading links in the chain of light signal transduction. Environmental signals, for example, mechanical stimuli, cold, light, etc., induce a single impulse of Ca2+ release in the cytosol of plant cells. In this case one can observe definite correlation between the intensity of the signal and the concentration of Ca2+ [130]. The increase of [Ca2+ ]cyt may be caused by entering of calcium ions

71

from the intercellular medium and/or by the release from intracellular depots (calcium vesicles, mitochondria, vacuoles, etc., where the concentration of Ca2+ can exceed 1 mM) [10]. The main depots of Ca2+ in plant cells are vacuoles and chloroplasts, and in animal cells, the endoplasmatic reticulum, mitochondria and the nuclei. An alternative origin of increasing [Ca2+ ]cyt can be the inhibition of the activity of Ca2+ pumps or a modification of the activity of Ca2+ /nH+ exchangers in the plasma membrane. It has been also proposed that besides Ca2+ pumps phytochrome could control Na+ /Ca2+ exchange; in protoplasts, activation of phytochrome was shown to lead to the entering of Ca2+ as well [131]. In plant cells, repeating releases of calcium ions, the so-called calcium oscillations, also play an important role in calcium signaling [132]. Thus, an increased concentration of free Ca2+ in the cytoplasm is one of the general links in the light signal transduction chain. However, the ratio between the increasing levels of cytosolic Ca2+ due to its release from the inner depots to that due to entering from the intercellular medium has not been established frequently. It is supposed that a release of Ca2+ can be stimulated first from the intracellular depots, and then the increased level of Ca2+ triggers the opening of calcium and other channels of the plasma membrane via which calcium ions enter. For example, the early research of Lew et al. [113] on calcium activation of K+ -channels of Mougeotia protoplasts suggested that Pfr stimulated the release of Ca2+ from calcium-containing vesicles, probably through a mechanism associated with formation of IP3 . Indeed, IP3 stimulated the release of intracellular calcium stores into the cytoplasm and induced short-term hyperpolarization of the cytoplasmic membrane due to activation of Ca2+ -sensitive K+ -permeable channels in the algal cells of Eremosphaera viridis [114]. In so doing, the increased concentration of free Ca2+ ions was obviously principal for hyperpolarization. Indeed, an artificial increase of [Ca2+ ] due to introduction of Ca2+ or its ionophores caused a short-term hyperpolarization of the cytoplasmic membrane in this alga [133]. An increase of free Ca2+ can activate K+ channels in the cytoplasmic membrane. Actually, evidences have been received that the level of free Ca2+ in the cytoplasm can control the activity of plasma membrane channels [18], blocking or activating these channels and, thus, regulating the distribution of ionic flows. Hence, an increase of Ca2+ level in the cytoplasm of Mougeotia cells was proposed to activate K+ channels, causing entering of external calcium and potassium ions via K+ channels inside the cells [113]. It is possible to think that in many cases, a gradual increase of the concentration of free Ca2+ in the cytoplasm causes partial or complete opening of calcium and potassium ion channels that stimulates the flow of calcium ions into the cell as discussed above. On the other hand, it has been shown that the extracellular concentration of Ca2+ is not always important for transduction of light signals. In particular, shrinking of Arabidopsis protoplasts induced by a blue light pulse did not depend on the concentration of external Ca2+ [128]. 4.2. Targets of calcium signals Calcium ions are able to bind (the number of coordinative links is 6–8) with a great number of cytosolic proteins that work as adaptors of Ca2+ -signal [10,17]. Binding of Ca2+ with Ca2+ -sensitive proteins is one of the key processes in the system of signaling both in animal and plant cells [99]. Calcium-binding protein analyses revealed that many of the muscle- and nerve-tissue-specific genes are not present in plants. Calcium-dependent protein kinase (CDPK), CDPKrelated kinase (CRK), and many plant-specific modified genes are involved in plant intracellular signal transduction systems [99]. There are primary sensors of calcium signals (Ca2+ -sensors), which include CaM and/or Ca2+ -dependent protein kinases and other proteins having the so-called “EF-hand” [10]. Most of the Ca2+ -binding proteins have this unique sequence of 12 amino acid residues

72

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

with high affinity to Ca2+ . The C2-domain sensitive to Ca2+ was found in many animal proteins. In plants the C2-domain is part of well-known signal components such as protein kinase C, as well as PIP2-phospholipase C and phospholipase D involved in lipid exchange [10]. CaM is a unique eukaryotic receptor of calcium, which can bind a lot of protein targets, mainly, upon binding with four calcium ions [134,135]. Note that CaM is an allosteric protein that reveals its activity only as a complex with calcium ions. To transmit the Ca2+ signal, CaM and other Ca2+ sensors such as calcineurin B-like proteins interact with target proteins and regulate their activity. CaM target proteins have been identified in higher plants and include protein kinases, metabolic enzymes, cytoskeleton-associated proteins, and others [134,135]. CaM and other Ca2+ -binding proteins are important for the light signal transduction. This was evidenced from a number of experiments, in particular from the disappearance of phytochromedependent rotation of the chloroplasts of Mougeotia with the inhibition of CaM activity [136]. In a tomato mutant deficient in phyA, light did not influence the formation of chlorophyll and anthocyanins [1]. Moreover, the light-activated synthesis of photosynthetic proteins was not induced. As a consequence, the formation of chloroplasts was suppressed in the mutant grown in the light. However, the introduction of oat phyA into the cells of hypocotyls of the mutant plants reestablished the expression of light-induced genes and induction of anthocyanins and chlorophyll synthesis was observed. As a result, mature chloroplasts that did not differ from the chloroplasts of the wild type, developed in the hypocotyls. Introduction of Ca2+ -CaM into cells also improved chloroplast formation. However, the chloroplasts formed upon the introduction of Ca2+ -CaM, were functionally incomplete because some photosynthetic proteins were still missing. For example, no synthesis of ferredoxin and cytochrome b6 f was observed. CaM participates in the transduction of light signals effecting on various CaM-binding proteins. Among them are the multifunctional Ca2+ -CaM-binding protein kinases (calmodulin-dependent protein kinases, CaMKs), with which the system Ca2+ -CaM, via transcription factors, influences gene expression [137]. Thus, the system Ca2+ CaM participates in the transduction of the phytochrome signal [33,125] (Fig. 2). 4.3. Other messengers: cAMP (cGMP), IP3 and DAG 4.3.1. The role of G-proteins and effectors Along with calcium ions the components of the phosphatidylinositol cycle, first of all IP3 and DAG [138], cyclic nucleotides [19,104,139,140], as well as G-proteins [25] play an important role in the processes of cell signals transduction. In this connection it has been suggested [16] that the active form of phytochrome can interact with G-proteins that have an influence on the level of cGMP and cytosolic Ca2+ , which could activate guanylate cyclase or membrane-bound phospholipase C. Phospholipase C can split phosphatidylinositol-4,5-diphosphate (PIP2) into two components: IP3 and DAG. IP3 favors the release of calcium ions from intracellular calcium stores. In particular, treatment with hydrophilic IP3 can induce the release of calcium ions from vacuoles and vesicles of the tonoplast, as well as from membrane vesicles of the endoplasmic reticulum [15,115,141]. DAG, the other product of the cycle, stimulates the activity of protein kinase C in the cytoplasmic membrane that is induced by calcium ions. The activated protein kinase C phosphorylates proteins of ion channels, regulating their conductivity. Viewing the possible role of cyclic nucleotides in the chain of transduction of photoreceptor signals, it should be stressed that all the components of the system of cyclic nucleotides are

found in plant cells: cAMP, cGMP, ADC, guanylyl cyclase (GC), phosphodiesterase (PDE), members of cyclic nucleotide-gated channel (CNGC)1 family, proteins binding cyclic nucleotides, and CaM [92,104,112,140,141,142]. In particular, cGMP and cAMP were detected in tobacco plant cells (BY2) by use of radioimmunological and mass-spectrometric methods [140]. Upon activation of ADC, catalyzing cAMP biosynthesis, or by inhibition of phosphodiesterase, which decomposes this nucleotide, the concentration of cAMP in the cell increases rapidly. Hence, the concentration of cAMP in the cell is determined by the ratio between the activities of these two enzymes. The action mechanism of cyclic nucleotides was first studied in animal systems. It was shown that the mediator, for example a hormone, stimulates the formation of cAMP probably via the system of chemoreceptor-G-protein-ADC, which could phosphorylate the proteins involved in the regulation of Ca2+ channels via cAMP-dependent protein kinases [122]. In plant cells, early observations indicated the influence of cAMP on the activity of Ca2+ channels in Characeae cell plasmalemma [105]. The regulatory effect of cAMP on Ca2+ -permeable channels in the plasma membrane of Arabidopsis leaf guard and mesophyll cells was also studied [104]. Local potential fixation in plasma membranes allowed determining the presence of Ca2+ -permeable channels activated directly by cAMP [104]. The early observations of Bowler et al. [21] showed the important role of cGMP in the phytochrome transduction chain. They found that cGMP introduced together with Ca2+ in a cell suspension of phytochrome mutants with disturbed formation of chloroplasts could restore the development of the chloroplasts up to mature ones, containing all the components of the photosynthetic apparatus. Moreover, it has been shown that cGMP together with Ca2+ acted by modulation of gene expression. It has been found further that cGMP concentration in the extracts of tissues from etiolated seedlings of Avena sativa increased at irradiation with a 5-min RL pulse and decreased at irradiation with RL with further irradiation with FRL, i.e. exhibited RL/FRL photoreversibility, demonstrating phytochrome control [142]. It is obvious that the enzymes involved in cGMP metabolism also play an important role in phytochrome signal transduction. The effect of light on cGMP level is thought to be basically due to modulation of the activity of GC rather than PDE. The above is supported by the suppression of red light-induced effect by GC inhibitor LY83583 while PDE inhibitor theophylline is ineffective [142]. The activity of GC responsible for cGMP synthesis was found to be predominantly located in the fraction of plasma membranes in cells of Avena sativa seedlings [143]. The reversible regulation of GC activity in etiolated seedlings by red/far-red light was also shown. The authors thus suggested that photoregulation of GC activity is mediated by the phytochrome. The data on activation of Ca2+ channels in algae [105] and Arabidopsis plants [104] indicate that exogenic cAMP activates protein kinases, which phosphorylate proteins regulating the conductivity of Ca2+ channels. However, no study to date has demonstrated the presence of a cyclic nucleotide-activated inward Ca2+ -conducting ion channel in plant cell membranes [104]. Evidently, changing the phytochrome state has an influence on the activity of the enzymes responsible for synthesis and hydrolysis of cAMP such as ADC and phosphodiesterase. For example, the phytochrome has been established to control the activity of ADC. The relationship between cAMP and cGMP and the concentration of free cytosolic Ca2+ has been investigated [23]. It has been shown that the introduction of dibutyryl analogs of cAMP and cGMP in a suspension of tobacco protoplasts leads to increasing levels of free Ca2+ in the cytosol. Dibutyryl analogs, unlike cAMP and cGMP, penetrate rather easily through the membrane. The increase of free Ca2+ was blocked by the inhibitor of Ca2+ channels, verapamil. The genes coding ion channels and controlled by cyclic nucleotides have been identified. For

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

example, about 20 types of such genes were found in the genome of Arabidopsis plants [112,116]. In principle, the phytochrome can interact with heterotrimeric G-proteins, though, as mentioned, a phytochrome-specific Gprotein has not been found yet [27]. A study using transgenic plants of Arabidopsis with high levels of G-protein expression suggested the participation of G-proteins in the transduction of light signal controlled by phyA [24]. In line with this study are the data of Volotovski et al. [143], which showed that photoregulation of GC activity is mediated by the phytochrome in cells of oat seedlings. In contrast to that however, Jones et al. [120] have found that single- and double-null mutants for the GPA1, AGB1 genes encoding the alpha

and beta subunits of heterotrimeric G-protein, respectively, have wild-type sensitivity to R and FR in Arabidopsis hypocotyl growth. Taking the above into account, it can be proposed that the sequence of events following transduction of the photoreceptor signal would be: photoreceptor → activated photoreceptor → G-protein → ADC → cAMP → cAMP-dependent protein kinases → phosphorylation of the effector proteins (EPs) → activation of the systems of ion exchange → the increase in free [Ca2+ ]cyt .

4.4. Phosphoinositide cycle and protein kinases It is suggested that both Ca2+ and cyclic nucleotides like cGMP are secondary messengers in the system of phytochrome signaling [7,23,38,125]. However, the question arises: how the phytochrome modulates the concentration of cyclic nucleotides or Ca2+ ? One of the possibilities concerns the interaction of the phytochrome with G-proteins [25] that could activate further such enzymes as guanylate cyclase or phospholipase C. As mentioned, phospholipase C catalyzes the hydrolysis of PIP2 into two secondary messengers, IP3 and DAG. Formation of IP3 results in an increase of cytosolic calcium, and DAG activates protein kinases [15,16]. Protein kinases play important roles in many regulatory processes in eukaryotes. Among the protein kinases, AGC kinases (cAMP-dependent protein kinase A, cGMP-dependent protein kinase G and phospholipid dependent protein kinase C) form a major superfamily and are known to play crucial roles in cell growth, apoptosis and many other important cellular processes in animals by regulating protein synthesis or gene transcription [10,27,144]. Protein kinases are supposed to play an important role in the transduction of light signals in plant cells [27]. For example, early observations [28] indicated the involvement of protein phosphorylation in the transduction of phytochrome signals. Also, many secondary messengers like cAMP, cGMP, Ca2+ , DAG affect on protein kinases, activating them. The most wide spread plant proteins controlled by calcium ions are Ca2+ dependent (CaMindependent) protein kinases [134]. At the same time CaM-specific protein kinases have been found [145]. Ca2+ -CaM-dependent protein kinase isolated from coleoptiles of etiolated maize plants is autophosphorylated and activated with orange light of 600 nm [146]. There are evidences that at least some phytochromes function as kinases or are bound with kinases physically [27]. Hence, a light quantum, without secondary messenger, may activate protein kinases. Protein kinase C was found both in animal and plant cells [147,148]. As mentioned above, protein kinase C can activate ion channels. Hence, examination of different factors having an influ-

73

ence on the activity of Ca2+ channels in Characeae cell plasmalemma suggested that protein kinase C can regulate the activity of the channels. Early studies of the influence of G-proteins, the modulators of phosphoinositide metabolism, neomycine and LaCl3 , as well as protein kinase C on phytochrome-dependent swelling of the protoplasts in etiolated wheat leaves [87] demonstrated similar effects as the ones observed during signal transduction in animal cells [121]. Hydrolysis of membrane phospholipid PIP2, the predecessor of DAG and IP3 , is also probably controlled by the phytochrome [23]. We suggest the following sequence of transduction of the light signal with the participation of the phosphatidylinositol signaling system and protein kinase C (Fig. 1):

where P is a ptotoreceptor, P* is the active form of the photoreceptor, PLC is phospholipase C, PKC is protein kinase C, SIE is the systems of ion exchange, CR is a cell response. In this case Pfr is associated with the plasma membrane or localized near it. 5. Cell structure and signal transduction Elements of the cytoskeleton such as microtubes and microfibrilles can play an important role in the mechanism of transduction as they influence many links in the light signal transduction chain of light signals, in particular in regards to the activity of ion channels. For example, the functioning of Ca2+ -activated Cl-channels of Characeae cell plasmalemma depends on microtubes [105]. Destruction of microtubes resulted in the activation of potentialdependent Ca2+ -permeable channels in the plasma membrane of carrot cells [123]. Obviously, this is connected with two factors: the viscous-elastic properties of the cytoskeleton that provides a mechanical link between the cell elements and the large negatively charged surface of the cytoskeleton elements [149]. In response to the formation of the active form of a photoreceptor, the availability of the charges leads to the localization of different proteins, in particular lipid kinases, phospholipases and G-proteins, on the surface of the cytoskeleton. Based on such views, a schematic diagram of light signal transduction has been developed for the cells Mougeotia, where light can induce chloroplasts rotation [67,71]. Mougeotia is a filamentous green alga that contains a single ribbon like chloroplast, which can rotate about its long axis in response to light. The sequence of transduction of the light signal with some minor modification can be as follows: P + hv → P* → Ca2+ → CaM → modification of the microtubes (microfilaments) → rotation of the chloroplast [150,151]. In contrast, Wagner [67] concluded that for the chloroplasts rotation in Mougeotia, the transduction of the light signal via the phytochrome occurred without direct participation of calcium ions. It is supposed that actin-binding proteins (ABPs), protein kinases, as well as various secondary messengers like Ca2+ and phosphoinositides [152] are involved in signal transduction with the participation of the actin cytoskeleton (Fig. 1). By analogy with animal cells the sequence of light signal transduction in plant cells could be as follows: P + hv → P* → G-protein → ADC→ cAMP → protein kinases → elements of the cytoskeleton → Ca2+ channels → increasing concentration of free cytosolic Ca2+ → metabolic response. In this case, the elements of the cytoskeleton that are involved are microtubes and/or microfilaments.

74

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

6. The regulation of gene activity The role of membrane changes triggered by the activation of photoreceptors and secondary messengers in light signal transduction is often rather clear. However, the relationship of these processes with the regulation of gene activity is not clear in many respects. One of the difficulties derives from the fact that the phytochrome in vitro bounds easily with many proteins [153]. Binding of the phytochrome with certain proteins and their participation in the transduction of phytochrome signals has been shown by mutation of genes that encode such proteins as PKS 1 [57], nucleoside-diphosphate kinase 2 (NDPK2) [2,154] or a phytochrome interacting factor (PIF3) [30] (Fig. 2) and many other ones [6,8,13]. Mutant plants overexpressing PKS1 have elongated hypocotyls, which indicate this protein belongs to negative regulators of the stem elongation controlled by PhyB [57]. Conversely, NDPK2 is likely to be a positive regulator of the signal chain with participation of both types of phytochrome, phyB and phyA. The phytochrome is able to interact with NDPK2 both in the cytoplasm and nucleus [154], as well as to stimulate phosphorylation of this protein [155]. Nevertheless, the mechanism affecting NDPK in plants is poorly studied. Presently, many phytochrometranscription factors and phytochrome-binding proteins involved in photomorphogenesis were found (for review see [13]). However, the biochemical function of most known phyA, such as PAT1, FAR1, FHY3 and SPA1 or phyB signaling molecules such as PIF4 are not well understood and is likely to be a primary focus of future research [6,13,34]. Early pharmacological studies and microinjections led to the suggestion of an important role of heterotrimeric G-proteins, cGMP and calcium ions as secondary messengers in phytochrome signaling [7,21,125]. Simultaneously, the question arises on the identification of the proteins interacting with the phytochrome. Three main proteins have been detailed: PIF 3 [29,30], PKS1 [57] and NDPK2 [154], which interact not only with phyA, but also with phyB. During the last decade, both phyA and phyB as well as blue-light photoreceptors were found to be bound with many transcription factors and have an influence on the expression of several genes regulating growth and photomorphogenesis [6,8,13]. The activation of photoreceptors, mainly phytochromes and cryptochromes, can significantly affect transcription through signal transduction pathways, especially by direct phytochrome effects on transcription factors [13]. For example, PIF3, a basic helix–loop–helix (bHLH) transcription factor that has been shown to bind G-boxes [156] (G-boxes being functionally important cis-elements within the promoters of some phytochrome sensitive genes [153,157]), can interact directly with both phyA and phyB, thus suggesting a phytochrome regulation of nuclear gene expression via direct interaction with transcription factors [13,36]. In such cases, the chain of light-induced signaling between nuclear-localized phytochrome and transcriptional regulation can be rather short [3,30,156] (Fig. 2). Nowadays, possible phytochrome binding with a rather great quantity of various phytochrome-interacting proteins has been proposed [13,36,157]. In particular, the protein PIL5 constitutes a key negative regulator of seeds germination in Arabidopsis [35]. A large number of transcription factors and phytochrome (photoreceptor)binding proteins forming a light-regulated transcriptional network were discovered and discussed in relevant reviews (for review see [6,13]). Phytochrome translocation into the nucleus has been demonstrated using mutants of phyA and phyB, which are incapable of binding with PIF3 in vitro [30]. However, no typical speckles formed in the nucleus and revealed by GFP fluorescence have been found. It is supposed that phytochrome signaling can include light-regulated gene expression by inhibiting COP1-, COP10- and CSN-dependent proteolysis of the HY5 transcription factor and by binding to activated cryptochromes (cry) (Fig. 2). Protein HY5 is

the transcription factor, key for photomorphogenesis that serves as signal integration point of major branches downstream of all photoreceptors (Fig. 2). HY5 is required for dissociation of the repressor complex, which hinders the development of a number of growth and morphogenesis processes in plants. The molecular mechanism by which photoactivated Phy induces altered expression of target genes is unknown, but there is evidence that intranuclear binding of the photoreceptor to PIF1 and PIF3 leads to rapid degradation of these bHLH proteins via the ubiquitin-proteosome system [32,158]. These observations suggest the possibility that Phy may regulate expression by altering the abundance of these transcription factors at the promoters of target genes. There are also many observations that the phosphorylation of transcription factors is a common modification that can influence their ability to bind to promoters [13]. Thus, phytochrome interacts both with nucleous proteins such as PIFs and cytoplasmic proteins such as phytochrome kinase substrate 1, PKS1 [2,13,57] shown in Fig. 2. Perhaps PKS1 is the best-characterized cytoplasmic phytochrome-signaling component. PKS1 interacts with the histidine kinase-related domain of phytochrome (HKRD) of both phyA and phyB, can be phosphorylated by phyA in vitro, and is a phosphoprotein in vivo [57]. Based on reverse genetic studies it has been proposed that PKS1 acts as a negative regulator of phyB and phyA signaling, which inhibits nuclear translocation of phytochromes [111]. The transcription factor PIF3 interacts only with Pfr and this complex dissociates when the active form of phytochrome turns into inactive one (PR ) following irradiation with a far-red light pulse [41]. There is evidence that intranuclear binding of the photoreceptor to PIF1 and PIF3 leads to rapid degradation of these bHLH proteins via the ubiquitinproteosome system [32,158]. However, the question of whether this postulated signal transfer mechanism is specific for PIF3 or occurs more generally across other target proteins has remained unanswered [159]. It is obvious that interaction of the phytochrome with PIF3 is controlled by light and by the daily rhythm [74]. However, this process can occur differently in various plants, in particular in transgenic plants of Arabidopsis and tobacco [79]. In some types of cells, additional regulation factors both cytoplasmic and nuclear, for example NDPK2, may be required for the function of the phytochrome transduction chain [160]. Light might also regulate the subcellular localization of transcription factors through phosphorylation [13,161]. For example, the protein CPRF2 (Fig. 2), the transcription factor from the G-boxbinding transcription factors (CBF), is localized in the cytosol in the dark and treatment with RL causes its import into the nucleus [84,85]. This process is reversible upon illumination with FRL that evidences the involvement of the phytochrome. Light induced interaction of photoreceptors with transcription factors is one of the key elements in light signal transduction in plant cells. Light-responsive transcription factors have been identified through screens for light-responsive cis-element (LRE)-binding proteins (Fig. 2) and through genetic analyses of mutants that are deficient in their response to specific types of light [6,13,36]. Protein kinases, phosphorylating many transcription factors, participate, as a rule, in this interaction [3,27]. However, the multiplicity of phytochrome forms, factors of protein modification interacting with the phytochrome, as well as the interaction of definite proteins with specific types of cells should be taken into account. Thus, one should not construct a simplified model, for example, PIF3 and PKS1 are active in hypocotyl, and NDPK2, in the region of hypocotyl loop [2]. The biochemical function of most known photoreceptor signaling molecules, such as PAT1, FAR1, FHY3 and SPA1 [13] remains largely unresolved and is likely to be a primary focus of future research. Finally, although many transcription factors have been defined as key factors in the light-regulated transcriptional network, their downstream targets are mostly undefined.

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

From the data available, a model for phytochrome signaling with modifications has been presented in Fig. 2. Key aspects of the model are [3,6,13,36]: (1) The phytochrome can initiate signal transduction from both the nucleus and the cytoplasm; (2) signaling can involve pathways: Pfr → G-protein → cGMP → X → light-induced transcription; Pfr → G-protein → Ca2+ -CaM → Y → light-induced transcription; (3) activated nuclear-localized phytochromes can interact with bHLH transcription family members and thereby rapidly activate transcription; (4) HY5 is a key transcription factor for all photoreceptors that activates light-responsive genes; (5) proteasome-dependent degradation of positively acting regulatory factors represses light-responsive gene expression in the dark; (6) phytochromes and cryptochromes directly interact to regulate co-action of these two classes of photoreceptors; (7) chromatin remodelling via nucleosome-binding complex DET1/DDB1 provides epigenetic control of light-responsive gene expression; and (8) light-regulated transcriptional control converges most frequently on G-boxes. 7. Interaction of photoreceptor signaling pathways Genetic studies in Arabidopsis thaliana have shown that phytochrome signaling involves a complex web of interactions [2,3,6,13,162]. The phytochromes sometimes act independently of one another, but in certain growth conditions and periods of development, they may also act redundantly or antagonistically. The genetic complexity of phytochrome signaling has been underscored by biochemical studies in which diverse proteins have been found that interact directly with various domains of the phytochrome molecule. In early studies, much attention was given to three of these proteins PIF3, PKS1, and nucleoside diphosphate kinase 2, whose activity is regulated by the phytochrome [2,57,162]. These three proteins are not structurally or functionally related and appear to interact with different domains of the phytochrome. Thus, these proteins do not share a common mechanism of interaction with phytochromes. A number of processes are known where interaction between phytochromes and cryptochromes is involved [6,67,163–165]. The interaction of photoreceptors upon quick shrinking of protoplasts isolated from leaves of mutant Arabidopsis, which was preliminary induced by a blue light pulse, was studied in detail [128]. In protoplasts of the mutant hy4, scarce on cryptochrome 1, and in double mutants of Arabidopsis with mutations in the genes phyA and phyB, deficient in phyA and phyB simultaneously, the shrinking was rather weak, whereas in mutants, deficient in one of the phytochromes, the protoplast shrinking was partial. It was concluded that a response to the blue light was strictly controlled by the phytochromes of both types. Studies regarding the after-effects induced by the interaction between RL and UV-B radiation in plants, involving the phytochrome and, hypothetically, a UV-B photoreceptor, were also reported. In some experiments it has been shown [166] that the phytochrome system modifies the growth inhibition of hypocotyl and cotyledonary leaves of lettuce due to UV-B application. It was supposed that the phytochrome system could adapt the plant to the damaging effect of UV-B. The interaction between a blue-light photoreceptor and a separate UV-B photoreceptor was investigated in stomatal guard cells [167]. In early experiments Lingakumar and Kulandaivelu [165] have studied the effects of low intensity red light and far-red light as well as UV-B on the activity of PS II and indicated that additional red light protected the photosynthetic apparatus from UV-B damage. They supposed that red light acting through the phytochrome is involved in photoprotection of PS II against UV-B in light-grown seedlings Vigna sinensis L. The influence of preirradiation in the region of

75

550–730 nm on UV-B induced damage of the photosynthetic apparatus in primary leaves of 20-d-old spinach plants (Spinacia overacea L. sv. “Giant”) and completely developed leaves of three-year-old field-ash plants exposed to UV-B was investigated in another study [168]. Leaf exposure to UV-B led to loss of chlorophyll (Chl), decline in activity of the photosynthetic apparatus indicated by Chl fluorescence and delayed light emission as well as accumulation of tiobarbituric acid reactive substances (TBARS). Pre-exposure to low intensity red radiation (I = 2–6 W m−2 , m = 660 and 623 nm) of 2–4 h duration eliminated partly the damage of the photosynthetic apparatus and reduced the content of TBARS in detached leaves held in darkness. Pre-irradiation of leaves with other spectral regions (m = 730, 690, 600 and 550 nm) had no influence on damage of the photosynthetic apparatus. It is suggested that the phytochrome or predecessors of Chl having absorption bands in the region of 620–660 nm may be involved in the protection of the photosynthetic apparatus against UV-B. Indeed, an ever-growing body of evidence suggests that at least one of the plastid components of light signaling is a tetrapyrrole [169]. This is in line with the observations of Kropat et al. [170,171], who demonstrated that exogenic porphyrins (Mg-Proto and Mg-ProtoMe) in Chlamydomonas cells can activate a heatand light-responsive HSP70A promoter fragment, but they do not affect an HSP70A promoter fragment that is induced only by heat. That suggests the tetrapyrrols induced transcription via a lightresponsive cis-element. The possible mechanism governing the protectory effect of RL from UV-B damage can be an increased activity of the antioxidant system resulting from irradiation with RL. Such effect was indicated in primary wheat leaves after short-term irradiation of etiolated seedlings with low intensity red light. In this case, the effect of RL (m = 660 nm) was partly reversible by FRL (m = 730 nm) that suggested the involvement of phytochrome (Dr. Lubimov, personal communication). In these experiments, the enhanced activity of peroxidase was indicated in wheat leaf tissue irradiated with RL. Thus, H2 O2 might constitute another link of the phytochrome transduction pathway (Fig. 1). However, the components of the red light signal pathways and the involvement of a UV-B photoreceptor in transduction of UV-B and RL signals are still unclear. Remarkably, little is known about the underlying mechanisms of UV-B perception and signal transduction. Brown et al. [62] used a genetic approach in Arabidopsis to identify the components involved specifically in UV-B perception or signaling. These authors reported that the Arabidopsis protein UV resistance locus 8 (UVR8) is a UV-B-specific signaling component that orchestrates the expression of a range of genes with vital UV-protective functions. According to this study, UVR8 regulates the expression of the transcription factor HY5 specifically when the plant is exposed to UV-B and protein HY5 is a key effector of the UVR8 signaling pathway that is required for survival under UV-B radiation. UVR8 is located principally in the nucleus and associates with chromatin via histones. Chromatin immunoprecipitation showed that UVR8 associates with chromatin in the HY5 promoter region, providing a mechanistic basis for its involvement in regulating transcription. It was concluded that UVR8 defines a UV-B-specific signaling pathway in plants that orchestrates the protective gene expression responses to UV-B required for plant survival in sunlight. The HY5 is a key element common for phytochrome and UV-A/blue light photoreceptor transduction chain [13] (Fig. 2). Therefore, the interaction between UV-B and phytochrome signalings may involve this regulatory protein. 8. Nucleus, chloroplast and mitochondrial signaling We viewed the early steps of photoreceptor signaling dissecting a cascade of signaling events occurring mainly in the cytosol and the

76

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

nucleus. However, the role of mitochondria and chloroplasts in cell light signal transduction is also dissected. Although nuclear genome encodes more than 90% of the proteins necessary for chloroplast maintenance and function, the functional state of the chloroplast has an influence on the state of the nucleus via retrograde signaling and synthesis of proteins in the nucleus. The research of the photoregulation of gene expression has focused on the nuclear genes that encode messages for chloroplast proteins: the small subunit (SSU) of ribulose-1,6-bisphosphate carboxylase/oxygenase (rubisco) and the major light-harvesting chlorophyll a/b-binding proteins associated with the light-harvesting complex of photosystem II (LHCIIb proteins). These proteins play important roles in chloroplast development and greening; hence their regulation by phytochrome has been studied in detail. This is agreed with early observations, which have shown that application of herbicide norflurazon, impairing the primary photosynthetic processes, led to reduced expression of nuclear-encoded genes such as LHCB (chlorophyll a/b binding protein) and RBCS (small subunit of Rubisco) [172]. It is suggested that chlorophyll precursors, such as Mgprotoporphyrin IX and protoporphyrin IX can be implicated in the signaling from the chloroplast to the nucleus [173]. Besides, active holophytochrome depends on phytochromobibilin, generated in the chloroplast. However, it is becoming increasingly clear that the chloroplast plays an important role, not only in regulating nuclear gene expression via retrograde signals, but also in controlling photomorphogenic responses [173,174]. How are the retrograde signals transduced from the chloroplast to the nucleus and the role of chloroplast in photomorphogenesis is so far under question. There are many observations concerning components of light signal transduction chain in animal cells, which evidences an important role of hem-containing enzymes such as cytochrome c oxidase in mitochondrial respiration chain [175,176]. Fig. 6 shows the main signaling pathways suggested in an animal cell. Opposite to plant organisms, animals do not contain phytochrome-like struc-

tures but porphyrins and hem-containing enzymes, for example, cytochrome c oxidase, are viewed in the literature as light photoreceptors [176]. Note that, a novel mitochondrial-signaling pathway in mammalian cells activated by red and near-IR radiation was discovered in 2004 [see 177]. It was shown recently [178] that IR-A radiation (760–1440 nm), elicits a retrograde signaling response in normal human skin fibroblasts. Broadband radiation (760–1440 nm) induced the formation of mitochondrial-derived ROS ([ROS]m) in cultured human dermal fibroblasts. Increase in [ROS]m caused an increase in intracellular redox potential, Eh (Fig. 6). Mitochondrial ROS generation (Fig. 6) has been measured during investigating the mechanisms of low power laser therapy [179]. It is important to note that both stimulation and inhibition of the respiratory chain can result in enhanced ROS generation [180]. Mitochondria have the capacity to communicate with the nucleus also by the structural changes in the organelle itself, e.g. by changes in fission–fusion homeostasis in a dynamic mitochondrial network [for review see 181]. Involvement of the redox-sensitive transcription factor NF-␬B and another transcription factor AP-1 in cellular signaling in irradiated cells has been suggested theoretically (Fig. 6). Together with photosynthesis, mitochondrial respiration is an important source of energy in plant cells. However, there is a little information concerning the role of mitochondrial components in light signal transduction in plant cells. The regulation of expression and activity of enzymes of mitochondrial respiration chain can be one of key elements of link between photosynthesis and respiration. There are observations that activity of rotenoneinsensitive NADH dehydrogenase from plant mitochondria is under phytochrome control (NADH-ase in Fig. 6) and is activated with red light (m = 660 nm) (Fomenko, personal communication). It might be suggested that plant mitochondria plays an important role in light signal transduction in a plant cell and a cascade of signaling events between the mitochondria and the nucleus will be detected next time.

Fig. 6. A schematic explaining putative mitochondrial retrograde signaling pathways after absorption visible and IR-A radiation (marked hv) by the photoacceptor, cytochrome c oxidase. Arrows ↑ and ↓ mark increase or decrease in the values, brackets [ ] mark concentration. FFH = changes in mitochondrial fusion–fission homeostasis; AP-1 = activator protein-1 (transcription factor); NF-␬B = nuclear factor kappa B (redox-sensitive transcription factor). Experimentally proved (→) and theoretically suggested ( ) pathways are shown. Em , plasma membrane electrical potential;  m , mitochondrial membrane potential; NADH-asa, NADH-dehydrogenase; Cytcox, Cytochrome c oxidase; Eh , cellular redox potential; intracellular pH (pHi ). Adopted from [176] with some modifications (gene activation marked in Fig. 6). For more details [see 176].

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

9. Conclusion Besides the above-listed photoreceptors, messengers and effector proteins, other compounds and cycles may be involved in light signal transduction. For example, in Euglena gracilis cells activated by blue light, ADC controls phototaxis and it can be considered as a new photoreceptor [182]. It is possible that choline-containing phospholipids like phosphatidylcholine and choline (choline chloride) itself, found in the cells in a free state, can play some definite role in the regulation of the chain of light signal transduction. This proposal agrees well with the fact that exogenic choline-containing compounds, in particular choline itself, can modify phytochrome effects [68]. It is known that growth retardant, chlorocholine chloride, the analogue of choline chloride, used exogenously, affects the permeability of the cuts from pea leaves for betacyanine and ␤alanine [183]. These observations indicate an influence of choline and chlorocholine chloride on cell membrane permeability that may be a result of changes in membrane lipid composition. As an example, choline chloride application to radish plants modified the phospholipid composition of the plasma membranes [184]. Choline-containing compounds may affect the activity of ion channels, primarily, calcium channels, either by changing the lipid composition of the membrane or affecting protein kinase C, the activity of which being modulated simultaneously by Ca2+ and phospholipids. In turn, modification of the phospholipid composition in the membranes and the activity of ion systems are connected with each other [185–192]. In conclusion, it should be stressed that many components of the receptor signal transduction chains and their regulation pathways in animal cells were also found in the light signal transduction chains of plant cells. However, there are some differences in the composition and function of these elements; hence, in many cases it was difficult to make unambiguous conclusions. The many systems of ion exchange and, first of all, Ca2+ channels as well as Ca2+ are likely to be a common link in all cases of signal transduction [10,98,176]. Water channels, ions channels and cell volume are likely to be important as well [187–192]. An important role among the mechanisms affecting the primary processes of light signal transduction in plant cells is likely to belong to proteolysis and phosphorylation of proteins, which control the activity of the system of ion exchange, transcription factors and other transduction elements. Phytochrome also controls several antioxidative enzymes, which take part in stress signal transduction during development of environmental stress such as UV-B radiation [189,193]. The chain of transduction is a consequence of events, in which several light signal transduction pathways that include the Ca2+ CaM system and cyclic nucleotides are implicated. Cytoplasmic and nuclear proteins interacting with photoreceptors can also be involved. The ideas considered are given in schematic diagrams (Figs. 1 and 2). They represent our global view of the mechanisms of light signaling that can, in many aspects, constitute an universal scheme illustrating general pathways of transduction of various cell signals, in particular the hormone ones. Acknowledgements We are grateful to Professors D.A. Los and V.P. Zinchenko for reading this manuscript and for valuable/critical comments and discussion. This work was financially supported by grants from Russian Foundation for Basic Research and by the program PCB RAS. References [1] G. Neuhaus, C. Bowler, R. Kern, N.-H. Chua, Calcium/calmodulin-dependent and -independent phytochrome signal transduction pathways, Cell 73 (1993) 937–952.

77

[2] M.M. Neff, C. Fankhauser, J. Chory, Light: an indicator of time and place, Genes Dev. 14 (2000) 257–271. [3] E. Schäfer, C. Bowler, Phytochrome-mediated photoreception and signal transduction in higher plants, EMBO Reports 3 (2002) 1042–1048. [4] H. Wang, X.W. Deng, Phytochrome Signaling Mechanism. The Arabidopsis Book, American Society of Plant Biologists, 2002, p. 238. [5] C. Lin, Blue light receptors and signal transduction, Plant Cell 14 (2002) 207–225. [6] H. Wang, X.W. Deng, Dissecting the phytochrome A-dependent signaling network in higher plants, Trends Plant Sci. 4 (2003) 172–178. [7] H. Nayyar, Calcium as environmental sensor in plants, Curr. Sci. 84 (2003) 893–902. [8] M. Chen, J. Chory, C. Fankhauser, Light signal transduction in higher plants, Ann. Rev. Gen. 38 (2004) 87–117. [9] E.P. Spalding, K.M. Folta, Illuminating topics in plant photobiology, Plant Cell Environ. 28 (2005) 39–53. [10] S.S. Medvedev, Plant calcium signaling system, Russ. J. Plant Physiol. 52 (2005) 282–305. [11] V.D. Kreslavski, S.I. Allakhverdiev, Transduction mechanisms of photoreceptor signal in plant cell, Russ. J. Biologicheskie Membrany 23 (2006) 275–295. [12] E. Schäfer, F. Nagy, Photomorphogenesis in Plants and Bacteria. Function and Signal Transduction Mechanisms, Springer, Dordrecht, The Netherlands, 2006. [13] Y. Jiao, O.S. Lau, X.W. Deng, Light-regulated transcriptional networks in higher plants, Nat. Rev. Genet. 8 (2007) 217–230. [14] M. Beato, S. Chavez, M. Truss, Transcriptional regulation by steroid hormones, Steroids 61 (1996) 240–251. [15] G.J. Allen, S.R. Muir, D. Sanders, Release of Ca2+ from individual plant vacuoles by both InsP3 and cyclic ADP-ribose, Science 268 (1995) 735–737. [16] S.K. Sopory, M.R. Chandok, Light induced signal transduction pathway involving inositol phosphates, Sub. Cell Biochem. 26 (1996) 345–370. [17] D. Sanders, B. Colin, J.F. Harper, Communicating with calcium, Plant Cell 11 (1999) 691–706. [18] D. Sanders, J. Pelloux, C. Brownlee, J.F. Harper, Calcium at the crossroads of signaling, Plant Cell 14 (2002) 401–417. [19] A.J. Trewavas, C. Rodrigues, C. Rato, R. Malho, Cyclic nucleotides: the current dilemma, Curr. Opin. Plant Biol. 5 (2002) 425–429. [20] W. Yang, S. Xia, Mechanisms of regulation and function of G-protein-coupled receptor kinases, World J. Gastroenterol. 12 (2006) 7753–7757. [21] C. Bowler, G. Neuhaus, H. Yamagata, N.-H. Chua, Cyclic GMP and calcium mediate phytochrome phototransduction, Cell 77 (1994) 73–81. [22] I.D. Volotovski, Ca2+ and intracellular signaling in plant cells: a role in phytochrome transduction, Membr. Cell Biol. 12 (1998) 721–742. [23] I.D. Volotovski, S.G. Sokolovsky, O.V. Molchan, M.R. Knight, Second messengers mediate increases in cytosolic calcium in tobacco protoplasts, Plant Physiol. 117 (1998) 1023–1030. [24] H. Okamoto, M. Matsui, X.W. Deng, Overexpression of the heterotrimeric Gprotein ␣-subunit enhances phytochrome-mediated inhibition of hypocotyl elongation in Arabidopsis, Plant Cell 13 (2001) 1639–1652. [25] S.M. Assmann, Heterotrimeric and unconventional GTP binding proteins in plant cell signaling, Plant Cell 14 (2002) 355–373. [26] J.H. Hurvey, The adenylyl and guanylyl cyclase superfamily, Curr. Opin. Struct. Biol. 8 (1998) 770–777. [27] S.C. Maheshwari, J.P. Khurana, S.K. Sopory, Novel light-activated protein kinases as key regulators of plant growth and development, Biosciences 24 (1999) 499–514. [28] S.J. Roux, Signal transduction in phytochrome responses, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Higher Plants, 2nd ed., Kluwer Academic Publishers, Dordrecht, 1994, pp. 187–209. [29] M. Ni, J.M. Tepperman, P.H. Quail, PIF3, a phytochrome interacting factor necessary for normal photoinduced signal transduction, is a novel basic helix–loop–helix protein, Cell 95 (1998) 657–667. [30] M. Ni, J.M. Tepperman, P.H. Quail, Binding of phytochrome B to its nuclear signaling partner PIF3 is reversibly induced by light, Nature 400 (1999) 781–784. [31] J.M. Christie, W.R. Briggs, Blue light sensing in higher plants, J. Biol. Chem. 276 (2001) 11457–11460. [32] B. Al-Sady, W. Ni, S. Kircher, E. Schäfer, P.H. Quail, Photoactivated phytochrome induces rapid PIF3 phosphorylation prior to proteasome-mediated degradation, Mol. Cell 23 (2006) 439–446. [33] F. Nagy, E. Schäfer, Nuclear and cytosolic events of light-induced, phytochrome-regulated signaling in higher plants, EMBO J. 19 (2000) 157–163. [34] E. Huq, P.H. Quail, PIF4, a phytochrome-interacting bHLH factor, functions as a negative regulator of phytochrome B signaling in Arabidopsis, EMBO J. 21 (2002) 2441–2450. [35] E. Oh, J. Kim, E. Park, J.-I. Kim, C. Kang, G. Choi, PIL5, phytochrome-interacting basic helix–loop–helix protein, is a key negative regulator of seed germination in Arabidopsis thaliana, Plant Cell 16 (2004) 3045–3058. [36] P.H. Quail, Phytochrome signal transduction network, in: E. Schäfer, F. Nagy (Eds.), Photomorphogenesis in Plants and Bacteria, Springer, Dordrecht, The Netherlands, 2006, pp. 335–356. [37] M.T. Osterlund, C.S. Hardtke, N. Wei, X.W. Deng, Targeted destabilization of HY5 during light-regulated development of Arabidopsis, Nature 405 (2000) 462–466. [38] C. Bowler, N.-H. Chua, Emerging themes of plant signal transduction, Plant Cell 6 (1994) 1529–1541.

78

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

[39] R.E. Kendrick, G. Kronenberg, in: R.E. Kendric, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994. [40] P.H. Quail, Phytochrome photosensory signaling networks, Nat. Rev. Mol. Cell Biol. 3 (2002) 85–93. [41] W.R. Briggs, M.A. Olney, Photoreceptors in plant photomorphogenesis to date: five phytochromes, two cryptochromes, one phototropin, and one superchrome, Plant Physiol. 125 (1) (2001) 85–88. [42] K.M. Folta, Green light stimulates early stem elongation, antagonizing lightmediated growth inhibition, Plant Physiol. 135 (2004) 1407–1416. [43] S. Tokutomi, D. Matsuoka, K. Zikihara, Molecular structure and regulation of phototropin kinase by blue light, Biochim. Biophys. Acta 1784 (2008) 133–142. [44] N.C. Rockwell, Y.-S. Su, J.C. Lagarias, Phytochrome structure and signaling mechanisms, Ann. Rev. Plant Biol. 57 (2006) 837–858. [45] L. Krall, J.W. Reed, The histidine kinase-related domain participates in phytochrome B function but is dispensable, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 8169–8174. [46] R.A. Sharrock, T. Clack, Patterns of expression and normalized levels of the five Arabidopsis phytochromes, Plant Physiol. 130 (2002) 442–456. [47] S.L. Tu, J.C. Lagarias, The phytochromes, in: W.R. Briggs, J. Spudich (Eds.), Handbook of Photosensory Receptors, Wiley-VCH, Weinheim, Germany, 2005, pp. 121–145. [48] S.G. Møller, P.J. Ingles, G.C. Whitelam, The cell biology of phytochrome signaling, New Physiol. 154 (2002) 553–590. [49] A.L. Mancinelli, The physiology of phytochrome action, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994, pp. 211–269. [50] H. Smith, Phytochromes and light signal perception by plants—an emerging synthesis, Nature 407 (2000) 585–691. [51] L.-G. Ma, J.-M. Li, L.-J. Qu, Z.-L. Chen, H.-Y. Zhao, X.W. Deng, Light control of Arabidopsis development entails co-ordinated regulation of genome expression and cellular pathways, Plant Cell 13 (2001) 2589–2607. [52] J. Hughes, T. Lamparter, F. Mittmann, E. Hartmann, W. Gärtner, A. Wilde, T. Börner, A prokaryotic phytochrome, Nature 386 (1997) 663. [53] S.-H. Bhoo, S.J. Davis, J. Walker, B. Karniol, R.D. Vierstra, Bacteriophytochromes are photochromic histidine kinase using a biliverdin chromophore, Nature 414 (2001) 776–777. [54] T. Lamparter, J. Hughes, Phytochromes and phytochrome-like proteins in cyanobacteria, in: A. Batschauer (Ed.), Photoreceptors and Light Signalling, Cambridge, RSC, UK, 2003, pp. 203–227. [55] M. Mutsuda, K.P. Michel, X.F. Zhang, B.L. Montgomery, S.S. Golden, Biochemical properties of CikA, an unusual phytochrome-like histidine protein kinase that resets the circadian clock in Synechococcus elongatus PCC7942, J. Biol. Chem. 278 (2003) 19102–19110. [56] K. Evans, A.P. Fordham-Skelton, H. Mistry, C.D. Reynolds, A.M. Lawless, M.Z. Papiz, A bacteriophytochrome regulates the synthesis of LH4 complexes in Rhodopseudomonas palustris, Photosynth. Res. 85 (2005) 169–180. [57] C. Fankhauser, K.C. Yeh, J.C. Lagarias, H. Zhang, T.D. Elich, J. Chory, PKS1, a substrate phosphorylated by phytochromes that modulates light signaling in Arabidopsis, Science 284 (1999) 1539–1541. [58] C. Lin, D. Shalitin, Cryptochrome structure and signal transduction, Annu. Rev. Plant Biol. 54 (2003) 469–496. [59] W.R. Briggs, J.M. Christie, Phototropins 1 and 2: versatile plant blue-light receptors, Trends Plant Sci. 7 (2002) 204–210. [60] J.M. Christie, Phototropin blue-light receptors, Annu. Rev. Plant Biol. 58 (2007) 21–45. [61] T. Imaizumi, H.G. Tran, T.E. Swartz, W.R. Briggs, S.A. Kay, FKF1 is essential for photoperiodic-specific light signaling in Arabidopsis, Nature 426 (2003) 302–306. [62] B.A. Brown, C. Cloix, G.H. Jiang, E. Kaiserli, P. Herzyk, D.J. Kliebenstein, G.I. Jenkins, A UV-B-specific signaling component orchestrates plant UV protection, Proc. Natl. Acad. Sci. U.S.A. 102 (50) (2005) 18225–18230. [63] K. Nozue, T. Kanegae, T. Imaizumi, S. Fukuda, H. Okamoto, K.C. Yeh, J.C. Lagarias, M. Wada, A phytochrome from the fern Adiantum with features of the putative photoreceptor NPH1, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 15826–15830. [64] H. Kawai, T. Kanegae, S. Christensen, T. Kiyosue, Y. Sato, T. Imaizumi, A. Kadota, M. Wada, Responses of ferns to red light are mediated by an unconventional photoreceptor, Nature 421 (2003) 287–290. [65] N. Suetsugu, F. Mittmann, G. Wagner, J. Hughes, M. Wada, A chimeric photoreceptor gene, NEOCHROME, has arisen twice during plant evolution, Proc. Natl. Acad. Sci. U.S.A. 102 (2005) 13705–13709. [66] H.A. Borthwick, S.B. Hendricks, M.W. Parker, E.H. Toole, V.K. Toole, A reversible photoreaction controlling seed germination, Proc. Natl. Acad. Sci. U.S.A. 38 (1952) 662–666. [67] G. Wagner, Cryptochrome, phototropin and phytochrome: molecules, evolution and interaction, in: I.D. Volotovski (Ed.), Plant Photobiology and Photosynthesis, Natl. Acad. Sci. Belarus, Minsk, 2000, pp. 55–123. [68] E.F. Kobzar’, V.D. Kreslavski, E.N. Muzafarov, Red radiation and choline compounds influence growth and greening of wheat seedlings, Photosynthetica 36 (1999) 333–340. [69] V.D. Kreslavski, E.F. Kobzar’, E.N. Muzafarov, Effect of red light and linuron on growth and ethylene production in Chlorella, Biol. Plant 39 (1997) 427–430. [70] M. Koornneef, J.W. Cone, R.G. Dekens, E.G. O’Herne-Roberts, C.J.P. Spruit, R.E. Kendrick, Photomorphogenic responses of long hypocotyl mutants of tomato, J. Plant Physiol. 120 (1985) 153–165.

[71] W. Haupt, D.P. Häder, Photomovement, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Higher Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994, pp. 707–732. [72] M. Kraml, Light direction and polarisation, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Higher Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994, pp. 186–191. [73] L.H. Pratt, Distribution and localization of phytochrome within the plant, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Higher Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994, pp. 163–185. [74] S. Kircher, P. Gil, L. Kozma-Bognar, E. Fejes, V. Speth, T. Husselstein-Muller, D. Bauer, E. Adam, E. Schäfer, F. Nagy, Nucleocytoplasmic partitioning of the plant photoreceptors phytochrome A, B, C, D, and E is regulated differentially by light and exhibits a diurnal rhythm, Plant Cell 14 (2002) 1541–1555. [75] W. Haupt, Light-mediated movement of chloroplasts, Annu. Rev. Plant Physiol. 33 (1982) 205–233. [76] A. Winands, G. Wagner, Phytochrome of green alga Mougeotia: cDNA sequence, autoregulation and phylogenetic position, Plant Mol. Biol. 32 (1996) 589–597. [77] F. Mittmann, G. Brücker, M. Zeidler, A. Repp, T. Abts, E. Hartmann, J. Hughes, Targeted knockout in Physcomitrella reveals direct actions of phytochrome in the cytoplasm, Proc. Natl. Acad. Sci. U.S.A. 101 (2004) 13939–13944. [78] S. Kircher, L. Kozma-Bognar, L. Kim, E. Adam, K. Harter, E. Schäfer, F. Nagy, Light quality-dependent nuclear import of the plant photoreceptors phytochrome A and B, Plant Cell 11 (1999) 1445–1456. [79] L. Kim, S. Kircher, R. Toth, E. Adam, E. Schäfer, F. Nagy, Light induced nuclear import of phytochrome-A:GFP fusion proteins is differentially regulated in transgenic tobacco and Arabidopsis, Plant J. 22 (2000) 125–133. [80] R. Yamaguchi, M. Nakamura, N. Mochizuki, S.A. Kay, A. Nagatani, Lightdependent translocation of a phytochrome B-GFP fusion protein to the nucleus in transgenic Arabidopsis, J. Cell Biol. 145 (1999) 437–445. [81] J. Kim, H. Yi, G. Choi, B. Shin, P.-S. Song, G. Choi, Functional characterization of phytochrome interacting factor 3 in phytochrome-mediated light signal transduction, Plant Cell 15 (2003) 2399–2407. [82] G.C. Whitelam, P.F. Devlin, Distribution and localization of phytochrome within the plant, Plant Cell Environ. 20 (1997) 752–758. [83] H. Smith, G.C. Whitelam, The shade avoidance syndrome: multiple responses mediated by multiple phytochromes, Plant Cell Environ. 20 (1997) 840–844. [84] S. Kircher, F. Wellmer, P. Nick, A. Rügner, E. Schäfer, K. Harter, Nuclear import of the parsley bZIP transcription factor CPRF2 is regulated by phytochrome photoreceptors, J. Cell Biol. 144 (1999) 201–211. [85] F. Wellmer, S. Kircher, A. Rügner, H. Frohnmeyer, E. Schäfer, K. Harter, Phosphorylation of the parsley bZIP transcription factor CPRF2 is regulated by light, J. Biol. Chem. 274 (1999), pp. 29476–19482. [86] C.H. Chung, S.-V. Lim, P.-S. Song, J.D. Berlin, J.R. Gooding, Swelling of etiolated oat protoplasts induced by phytochrome, cAMP, and gibberellic acid: a kinetic study, Plant Cell Physiol. 29 (1988) 855–860. [87] M.E. Bossen, R.E. Kendrick, W.J. Vredenberg, The involvement of a G-protein in phytochrome-regulated, Ca2+ -dependent swelling of etiolated wheat protoplasts, Physiol. Plant 80 (1990) 55–62. [88] P.S. Shacklock, N.D. Read, A.J. Trewavas, Cytosolic free calcium mediates red light-induced photomorphogenesis, Nature 358 (1992) 753–755. [89] B.S. Serlin, R.R. Krasnoshtein, J. Krol, K.D. Sumida, Phytochrome activation of K+ channels and chloroplast rotation in Mougeotia: the escape times, Plant Cell Physiol. 37 (1996) 175–179. [90] E. Johannes, E. Ermolayeva, D. Sanders, Red light-induced membrane potential transients in the moss Physcomitrella patens: ion channel interaction in phytochrome signaling, J. Exp. Bot. 48 (1997) 599–608. [91] M.R.G. Roelfsema, R. Hedrich, Studying guard cells in the intact plant: modulation of stomatal movement by apoplastic factors, New Physiol. 153 (2002) 425–431. [92] S. Takagi, S.G. Kong, Y. Mineyuki, M. Furuya, Regulation of actin-dependent cytoplasmic motility by type II phytochrome occurs within seconds in Vallisneria gigantea epidermal cells, Plant Cell 15 (2003) 331–345. [93] A. Harada, T. Sakai, K. Okada, Phot1 and phot2 mediate blue light-induced transient increase in cytosolic Ca2+ through different manners in Arabidopsis leaves, Proc. Natl. Acad. Sci. U.S.A. 14 (2003) 8583–8588. [94] R. Racusen, R.L. Satter, Rhythmic and phytochrome-regulated changes in transmembrane potential in Samanea pulvini, Nature 255 (1975) 408–410. [95] E.P. Spalding, Ion channels and the transduction of light signals, Plant Cell Environ. 23 (2000) 665–674. [96] D.J. Aidley, P.R. Stanfield, Ion Channels: Molecules in Action, Cambridge University Press, Cambridge, New York, 1996, pp. 204. [97] S. Zimmermann, T. Ehrhardt, G. Plesch, B. Müller-Röber, Ion channels in plant signaling, Cell. Mol. Life Sci. 55 (1999) 183–203. [98] P.J. White, Calcium channels in higher plants, Biochim. Biophys. Acta 1465 (2000) 171–189. [99] T. Nagata, S. Iizumi, K. Satoh, H. Ooka, J. Kaway, P. Carninci, Y. Hayashizaki, Y. Otomo, K. Murakami, K. Matsubara, S. Kikuchi, Comparative analysis of plant and animal calcium signal transduction element using plant full-length cDNA data, Mol. Biol. Evol. 21 (2004) 1855–1870. [100] E.P. Spalding, D.J. Cosgrove, Large plasma-membrane depolarization precedes rapid blue-light-induced growth inhibition in cucumber, Planta 178 (1989) 407–410. [101] M.H. Cho, E.P. Spalding, An anion channel in Arabidopsis hypocotyls activated by blue light, Proc. Natl. Acad. Sci. U.S.A. 93 (1996) 8134–8138.

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80 [102] E.D. Ermolayeva, S. Sanders, E. Johannes, Ionic mechanism and role of phytochrome-mediated membrane depolarization in caulonemal side branch initial formation in the moss Physcomitrella patens, Planta 201 (1997) 109–118. [103] J. Alexandre, J.P. Lassalles, R.T. Kado, Opening of Ca2+ channels in isolated red beet root vacuole membrane by inositol 1,4,5-trisphosphate, Nature 343 (1990) 567–570. [104] F. Lemtiri-Chlieh, G.A. Berkowitz, Cyclic adenosine monophosphate regulates calcium channels in the plasma membrane of Arabidopsis leaf guard and mesophyll cells, J. Biol. Chem. 279 (34) (2004) 35306–35312. [105] O.M. Zherelova, Ca2+ -channels in Nitelopsis optusa, Characeae cell plasmalemma and regulation of their activity, Comp. Biochem. Physiol. 94A (1) (1989) 141–145. [106] V. Demidchik, R.J. Davenport, M. Tester, Nonselective cation channels in plants, Annu. Rev. Plant Biol. 53 (2002) 67–107. [107] S.J. Roux, J. Yguerabide, Photoreversible conductance changes induced by phytochrome in model lipid membranes, Proc. Natl. Acad. Sci. U.S.A. 70 (3) (1973) 762–764. [108] M.N. Weisenseel, H.K. Ruppert, Phytochrome and calcium iones are involved in light-induced membrane depolarization in Nitella, Planta 137 (1977) 225–229. [109] G. Böse, P. Schwille, T. Lamparter, The mobility of phytochrome within protonemal tip cells of the moss Ceratodon purpureus, monitored by fluorescence correlation spectroscopy, Biophys. J. 87 (2004) 2013–2021. [110] K.C. Yeh, J.C. Lagarias, Eukaryotic phytochromes: light-regulated serine/threonine protein kinases with histidine kinase ancestry, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 13976–13981. [111] C. Fankhauser, Phytochromes as light-modulated protein kinases, Cell Dev. Biol. 11 (2001) 467–473. [112] P. Mäser, S. Thomine, J.I. Schroeder, J.M. Ward, K. Hirschi, et al., Phylogenetic relationships within cation transporter families of Arabidopsis, Plant Physiol. 126 (2001) 1646–1667. [113] R.R. Lew, B.S. Serlin, C.L. Schaut, M.E. Stockton, Red light regulates calciumactivated potassium channels in Mougeotia plasma membrane, Plant Physiol. 92 (1990) 822–830. [114] B. Förster, Injected inositol 1,4,5-trisphosphate activates Ca2+ -sensitive K+ channels in the plasmalemma of Eremosphaera viridis, FEBS Lett. 269 (1990) 197–201. [115] S.R. Muir, M.A. Bewell, D. Sanders, G.J. Allen, Ligand-gated Ca2+ channels and signaling in higher plants, J. Exp. Bot. 48 (1997) 589–597. [116] C. Kohler, T. Merkle, G. Neuhaus, Characterization of a novel gene family of putative cyclic nucleotide and calmodulin-regulated ion channels in Arabidopsis thaliana, Plant J. 18 (1999) 97–104. [117] C. Kohler, G. Neuhaus, Characterization of calmodulin binding to cyclic nucleotide-gated ion channels from Arabidopsis thaliana, FEBS Lett. 471 (2000) 133–136. [118] T. Aoyama, N.-H. Chua, A glucocorticoid-mediated transcriptional induction system in transgenic plants, Plant J. 11 (1997) 605–612. [119] G.S. Aharon, A. Gelli, W.A. Snedden, E. Blumwald, Activation of a plant plasma membrane Ca2+ -channel by TG ␣1, a heterotrimeric G-protein ␣-subunit homologue, FEBS Lett. 424 (1998) 17–21. [120] A.M. Jones, J.R. Ecker, J.-G. Chen, A reevaluation of the role of the heterotrimeric G protein in coupling light responses in Arabidopsis, Plant Physiol. 131 (2003) 1623–1627. [121] Y. Nishizuka, Protein kinase signal transduction, Trends Biochem. Sci. 9 (4) (1984) 163–166. [122] P.G. Kostyuk, M.A. Ostrovskaya, Intracellular Signalling, Nauka, Moscow, 1988 (in Russian). [123] L. Thion, C. Mazars, P. Thuleau, A. Graziana, M. Rossignol, M. Moreau, R. Ranjeva, Activation of plasma membrane voltage-dependent calcium-permeable channels by disruption of microtubules in carrot cells, FEBS Lett. 340 (1996) 45–50. [124] A. Tretyn, R.E. Kendrick, M.E. Bossen, The effect of a calcium-channel antagonist, nifedipine and agonist Bay K-8644, on the phytochrome-controlled swelling of etiolated wheat protoplasts, Physiol. Plant 78 (1990) 230–235. [125] G. Neuhaus, C. Bowler, K. Hiratsuka, H. Yamagata, N.-H. Chua, Phytochromeregulated repression of gene expression requires calcium and cGMP, EMBO J. 16 (1997) 2554–2564. [126] B.S. Serlin, S.J. Roux, Modulation of chloroplast movement in the green alga Mougeotia by the Ca2+ -ionophore A23187 and by calmodulin antagonists, Proc. Natl. Acad. Sci. U.S.A. 81 (1984) 6368–6372. [127] J. Elzenga, M. Staal, H. Prins, Calcium-calmodulin signaling is involved in lightinduced acidification by epidermal leaf cells of pea, Pisum sativum L., J. Exp. Bot. 48 (1997) 2055–2060. [128] X. Wang, M. Iino, Interaction of cryptochrome 1, phytochrome, and ion fluxes in blue- light-induced shrinking of Arabidopsis hypocotyl protoplasts, Plant Physiol. 117 (1998) 1265–1279. [129] U. Russ, F. Grolig, G. Wagner, Changes of cytoplasmic free Ca2+ in the green alga Mougeotia scalaris as monitored with indo-1, and their effect on the velocity of chloroplast movement, Planta 184 (1991) 105–112. [130] M.R. Knight, H. Knight, N.J. Watkins, Calcium and the generation of plant form, Philos. Trans. R. Soc. Lond. (B) 350 (1995) 83–86. [131] I.D. Volotovski, Transduction mechanisms of plant phytochrome effects, Russ. J. Biophys. 40 (4) (1995) 1291–1307. [132] M.J. Berridge, M.D. Bootman, H.L. Roderick, Calcium signalling: dynamics, homeostasis and remodeling, Nat. Rev. Mol. Cell Biol. 4 (2003) 517–529.

79

[133] M. Thaler, W. Steigner, B. Förster, K. Köhler, W. Simonis, W. Urbach, Calcium activation of potassium channels in the plasmalemma of Eremosphaera viridis, J. Exp. Bot. 40 (1989) 1195–1203. [134] R.E. Zielinski, Calmodulin and calmodulin binding protein in plants, Annu. Rev. Plant Physiol. Plant Mol. Biol. 49 (3) (1998) 697–725. [135] V.S. Reddy, G.S. Ali, A.S. Reddy, Genes encoding calmodulin-binding proteins in the Arabidopsis genome, J. Biol. Chem. 277 (2002) 9840–9852. [136] C. Zörb, K.D. Brunner, M. Perbandt, Ch. Betzel, G. Wagner, Cloning, recombinant expression and characterization of wild type-105-trp-calmodulin of the green alga Mougeotia scalaris, Bot. Acta 111 (1998) 346–353. [137] E.E. Corcoran, A.R. Means, Defining Ca2+ /calmodulin-dependent protein kinase cascades in transcriptional regulation, J. Biol. Chem. 276 (5) (2001) 2975–2978. [138] B. Mueller-Roeber, C. Pical, Inositol phospholipid metabolism in Arabidopsis. Characterized and putative isoforms of inositol phospholipid kinase and phosphoinositide-specific phospholipase C, Plant Physiol. 130 (2002) 22–46. [139] R.P. Newton, L. Roef, E. Witters, H. Van Onckelen, Cyclic nucleotides in higher plants: the enduring paradox, New Phytol. 143 (1999) 427–455. [140] H. Richards, S. Das, C.H. Smith, L. Pereira, A. Geisbrecht, N.J. Devitt, D.E. Games, J. van Geyschem, A.G. Brenton, R.P. Newton, Cyclic nucleotide content of tobacco BY-2 cells, Phytochemistry 61 (5) (2002) 531–537. [141] S.R. Muir, D. Sanders, Inositol 1,4,5-trisphosphate-sensitive Ca2+ release across nonvacuolar membranes in cauliflower, Plant Physiol. 114 (1997) 1511–1521. [142] L.V. Dubovskaya, O.V. Molchan, I.D. Volotovski, Photoregulation of endogenic level of cAMP in oat seedlings, Russ. J. Plant Physiol. 48 (1) (2001) 26–29. [143] I.D. Volotovski, L.V. Dubovskaya, O.V. Molchan, Photoreceptor phytochrome regulates the cyclic guanosine 3,5-monophosphate synthesis in Avena sativa L. cells, Bulg. J. Plant Physiol. 29 (1–2) (2003) 3–12. [144] S.K. Sopory, M. Munshi, Protein kinases and phosphatases and their role in signalling in plants, Crit. Rev. Plant Sci. 17 (1998) 245–318. [145] L. Zhang, Y.-T. Lu, Calmodulin-binding protein kinases in plants, Trends Plant Sci. 8 (2003) 123–127. [146] S. Pandey, S.K. Sopory, Zea mays CCaMK: autophosphorylation-dependent substrate phosphorylation and down-regulation by red light, J. Exp. Bot. 52 (357) (2001) 691–700. [147] D.C. Elliott, J.D. Skinner, Calcium-dependent phospholipids-activated protein kinase in plants, Photochemistry 25 (1) (1986) 39–45. [148] D.C. Elliott, V.S. Kokke, Partial purification and properties of a protein kinase C type enzyme from plants, Photochemistry 26 (1987) 2929–2935. [149] P.A. Janmey, The cytoskeleton and cell signaling: component localization and mechanical coupling, Physiol. Rev. 78 (1998) 763–781. [150] G. Wagner, Intracellular movement, Prog. Bot. 57 (1996) 68–80. [151] B. Al-Rawass, F. Grolig, G. Wagner, High irradiance blue light affects cortical microtubules in the green alga Mougeotia scalaris, Plant Cell Physiol. 38 (1997) 882–886. [152] N.L. Klyachko, Actin cytosceleton and shape of plant cell, Russ. J. Plant Physiol. 51 (6) (2004) 918–925. [153] P.H. Quail, Phytochrome interacting factors, in: G. Whitelam, K. Halliday (Eds.), Light and Plant Development, Blackwell Publishing, Oxford, UK, 2007, pp. 81–105. [154] G. Choi, H. Yi, J. Lee, Y.-K. Kwon, M.S. Soh, B. Shin, Z. Luka, T.R. Hahn, P.-S. Song, Phytochrome signaling is mediated through nucleoside diphosphate kinase 2, Nature 401 (1999) 610–613. [155] N. Tanaka, T. Ogura, T. Noguchi, H. Hirano, N. Yabe, K. Hasunuma, Phytochromemediated light signals are transduced to nucleoside diphosphate kinase in Pisum sative L. cv. Alaska, J. Photochem. Photobiol. B 45 (1998) 113–121. [156] J.F. Martinez-Garcia, E. Huq, P.H. Quail, Direct targeting of light signals to a promoter element-bound transcription factor, Science 288 (2000) 859–863. [157] N.A. Eckardt, Light signaling revisited, Plant Cell 16 (2004) 1355–1357. [158] H. Shen, J. Moon, E. Huq, PIF1 is regulated by light-mediated degradation through the ubiquitin-26S proteasome pathway to optimize photomorphogenesis of seedlings in Arabidopsis, Plant J. 44 (2005) 1023–1035. [159] Y. Shen, R. Khanna, M. Christine, C.M. Carle, P.H. Quail, Phytochrome induces rapid PIF5 phosphorylation and degradation in response to red-light activation, Plant Physiol. 145 (2007) 1043–1051. [160] Y. Shen, J.-I. Kim, P.-S. Song, NDPK2 as a signal transducer in the phytochromemediated light signaling, J. Biol. Chem. 280 (7) (2005) 5740–5749. [161] J. Kim, Y. Shen, Y.J. Han, J.E. Park, D. Kirchenbauer, M.S. Soh, F. Nagy, E. Schafer, P.S. Song, Phytochrome phosphorylation modulates light signaling by influencing the protein–protein interaction, Plant Cell 16 (2004) 2629–2640. [162] J. Chory, D. Wu, Weaving the complex web of signal transduction, Plant Physiol. 125 (2001) 77–80. [163] H. Mohr, Coaction between pigment systems, in: R.E. Kendrick, G.H.M. Kronenberg (Eds.), Photomorphogenesis in Plants, Kluwer Academic Publishers, Dordrecht, The Netherlands, 1994, pp. 353–376. [164] J.J. Casal, Phytochromes, cyptochromes, phototrop, photoreceptor interactions in plants, Photochem. Photobiol. 71 (2000) 1–11. [165] K. Lingakumar, G. Kulandaivelu, Regulatory role of phytochrome on ultraviolet-B (280–315 nm) induced changes in growth and photosynthetic activity of Vigna sinensis L., Photosynthetica 29 (3) (1993) 341–351. [166] E.F. Kobzar’, V.D. Kreslavski, E.N. Muzafarov, Photomorphogenetic responses to UV radiation and short-term red light in lettuce seedlings, Plant Growth Regul. 26 (1998) 73–76. [167] W. Eisinger, R.A. Bogomolni, L. Taiz, Interactions between a blue–green reversible photoreceptor and a separate UV-B receptor in stomatal guard cells, Am. J. Bot. 90 (2003) 1560–1566.

80

V.D. Kreslavski et al. / Journal of Photochemistry and Photobiology C: Photochemistry Reviews 10 (2009) 63–80

[168] V.D. Kreslavski, A.A. Ivanov, A.A. Kosobrukhov, Low energy light in the 620–660 nm range reduces the UV-B-induced damage to photosystem II in spinach leaves, Biophysics 49 (5) (2004) 767–771. [169] M. Surpin, R.M. Larkin, J. Chory, Signal transduction between the chloroplast and the nucleus, Plant Cell (Supplement) (2002) S327–338. [170] J. Kropat, U. Oster, W. Rudiger, C.F. Beck, Chlorophyll precursors are signals of chloroplast origin involved in light induction of nuclear heat-shock genes, Proc. Natl. Acad. Sci. U.S.A. 94 (1997) 14168–14172. [171] J. Kropat, U. Oster, W. Rudiger, C.F. Beck, Chloroplast signaling in the light induction of nuclear HSP70 genes requires the accumulation of chlorophyll precursors and their accessibility to the cytoplasm/nucleus, Plant J. 24 (2000) 523–531. [172] R.E. Susek, F.M. Ausubel, J. Chory, Signal transduction mutants of Arabidopsis uncouple nuclear CAB and RBCS gene expression from chloroplast development, Cell 74 (1993) 787–799. [173] S.G. Møller, T. Kunkel, N.-H. Chua, A plastidic ABC protein involved in intercompartmental communication of light signaling, Genes Dev. 15 (2001) 90–103. [174] N. Mochizuki, J.A. Brusslan, R. Larkin, A. Nagatani, J. Chory, Arabidopsis genomes uncoupled 5 (GUN5) mutant reveals the involvement of Mgchelatase H subunit in plastid-to-nucleus signal transduction, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 2053–2058. [175] T. Karu, Mechanisms of low-power laser light action on cellular level, in: Z. Simunovic (Ed.), Lasers in Medicine and Dentistry, Vitgraf, Rijeka (Croatia), 2000, pp. 97–125. [176] T.I. Karu, Mitochondrial signaling in mammalian cells activated by red and near-IR radiation, Photochem. Photobiol. 84 (2008) 1091–1099. [177] T.I. Karu, L.V. Pyatibrat, N.I. Afanasyeva, A novel mitochondrial signaling pathway activated by visible-to-near infrared radiation, Photochem. Photobiol. 80 (2004) 366–372. [178] P. Schroeder, C. Pohl, C. Calles, C. Marks, S. Wild, J. Krutmann, Cellular response to infrared radiation involves retrograde mitochondrial signaling, Free Radic. Biol. Med. 43 (2007) 128–135. [179] T. Karu, Primary and secondary mechanisms of action of visible-to-near IR radiation on cells, J. Photochem. Photobiol. B. Biol. 49 (1999) 1–17. [180] P.S. Brookes, Y. Yoon, J.L. Robotham, M.W. Anders, S.S. Sheu, Calcium, ATP, and ROS: a mitochondrial love–hate triangle, Am. J. Physiol. Cell Physiol. 287 (2004) C817–C833. [181] M.T. Ryan, N.J. Hoogenraad, Mitochondrial–nuclear communications, Annu. Rev. Biochem. 76 (2007) 701–722. [182] M. Ntefidou, M. Iseki, M. Watanabe, M. Lebert, D.-P. Häder, Photoactivated adenylyl cyclase controls phototaxis in the flagellate Euglena gracilis, Plant Physiol. 133 (2003) 1517–1521. [183] D.M. Fabijan, P.P. Dhindsa, D.M. Reid, Effects of growth retardants on tissue permeability in Pisum sativum and Beta vulgaris, Planta 152 (1981) 481–486.

[184] G.V. Novitskaya, T.K. Kocheshkova, T.V. Feofitilactova, Yu.I. Novotski, Effect of choline chloride on lipid content and composition in leaves of major magnetooriented redish species, Russ. J. Plant Physiol. 46 (4) (2004) 618–625. [185] S.I. Allakhverdiev, Y. Nishiyama, I. Suzuki, Y. Tassaka, N. Murata, Genetic engineering of the unsaturation of fatty acids in membrane lipids alters the tolerance of Synechocystis to salt stress, Proc. Natl. Acad. Sci. U.S.A. 96 (1999) 5862–5867. [186] S.I. Allakhverdiev, M. Kinoshita, M. Inaba, I. Suzuki, N. Murata, Unsaturated fatty acids in membrane lipids protect the photosynthestic machniary against salt-induced damage in Synechococcus, Plant Physiol. 125 (2001) 1842–1853. [187] S.I. Allakhverdiev, A. Sakamoto, Y. Nishiyama, N. Murata, Inactivation of photosystems I and II in response to osmotic stress in Synechococcus. Contribution of water channels, Plant Physiol. 122 (2000) 1201–1208. [188] S.I. Allakhverdiev, A. Sakamoto, Y. Nishiyama, M. Inaba, N. Murata, Ionic and osmotic effects of NaCl-induced inactivation of photosystem I and II in Synechococcus, Plant Physiol. 123 (2000) 1047–1056. [189] S.I. Allakhverdiev, N. Murata, Environmental stress inhibits the synthesis de novo of proteins involved in the photodamage-repair cycle of photosystem II in Synechocystis sp. PCC 6803, Biochim. Biophys. Acta 1657 (2004) 23–32. [190] S.I. Allakhverdiev, N. Murata, Salt stress inhibits photosystems II and 1 in cyanobacteria, Photosynth. Res. 98 (2008) 529–539. [191] Y. Kanesaki, I. Suzuki, S.I. Allakhverdiev, K. Mikami, N. Murata, Salt stress and hyperosmotic stress regulate the expression of different sets of genes in II in Synechocystis sp. PCC 6803, Biochem. Biophys. Res. Commun. 290 (2002) 339–348. [192] A. Shapiguzov, A.A. Lyukevich, S.I. Allakhverdiev, T.V. Sergeyenko, I. Suzuki, N. Murata, D.A. Los’, Osmotic shrinkage of cells of Synechocystis sp. PCC 6803 by water efflux via aquaporins regulates the osmostress-inducible gene expression, Microbiology 151 (2005) 447–455. [193] Z. Qi, M. Yue, R. Han, X.L. Wang, The damage repair role of He-Ne laser on plants exposed to different intensities of ultraviolet-B radiation, Photochem. Photobiol. 75 (2002) 680–686. [194] P. Leivar, E. Monte, B. Al-Sady, C. Carle, A. Storer, J.M. Alonso, J.R. Ecker, P.H. Quail, The Arabidopsis phytochrome-interacting factor PIF7, together with PIF3 and PIF4, regulates responses to prolonged red light by modulating phyB levels, Plant Cell 20 (2008) 337–352. [195] J.J. Casal, M.J. Yanovsky, Regulation of gene expression by light, Int. J. Dev. Biol. 49 (2005) 501–511. [196] P.H. Quail, Phytochrome-regulated gene expression, J. Integ. Plant Biol. 49 (1) (2007) 11–20. [197] J.C. Litts, J.M. Kelly, J.C. Lagarias, Structure–function studies on phytochrome, J. Biol. Chem. 258 (18) (1983) 11025–11031.