The Variegated Mutants Lacking Chloroplastic FtsHs Are Defective in D1 Degradation and Accumulate Reactive Oxygen Species

In the photosynthetic apparatus, a major target of photodamage is the D1 reaction center protein of photosystem II (PSII). Photosynthetic organisms have developed a PSII repair cycle in which photodamaged D1 is selectively degraded. A thylakoid membrane-bound metalloprotease, FtsH, was shown to play a critical role in this process. Here, the effect of FtsHs in D1 degradation was investigated in Arabidopsis (Arabidopsis thaliana) mutants lacking FtsH2 (yellow variegated2 [var2]) or FtsH5 (var1). Because these mutants are characterized by variegated leaves that sometimes complicate biochemical studies, we employed another mutation, fu-gaeri1 (fug1), that suppresses leaf variegation in var1 and var2 to examine D1 degradation. Two-dimensional blue native PAGE showed that var2 has less PSII supercomplex and more PSII intermediate lacking CP43, termed RC47, than the wild type under normal growth light. Moreover, our histochemical and quantitative analyses revealed that chloroplasts in var2 accumulate significant levels of reactive oxygen species, such as superoxide radical and hydrogen peroxide. These results indicate that the lack of FtsH2 leads to impaired D1 degradation at the step of RC47 formation in PSII repair and to photooxidative stress even under nonphotoinhibitory conditions. Our in vivo D1 degradation assays, carried out by nonvariegated var2 fug1 and var1 fug1 leaves, demonstrated that D1 degradation was impaired in different light conditions. Taken together, our results suggest the important role of chloroplastic FtsHs, which was not precisely examined in vivo. Attenuated D1 degradation in the nonvariegated mutants also suggests that leaf variegation seems to be independent of the PSII repair.Excessive light often limits the growth of photosynthetic organisms by irreversibly inactivating the photosynthetic apparatus, a process called photoinhibition (for review, see Barber and Andersson, 1992; Aro et al., 1993). A major target of photodamage is PSII (for review, see Barber and Andersson, 1992; Aro et al., 1993; Murata et al., 2007), a large pigment-protein complex in the thylakoid membrane. In particular, the reaction center D1 protein, which binds cooperatively to D2 and carries cofactors required for electron flow from the manganese cluster of the water-oxidizing complex to the plastoquinone pool (Zouni et al., 2001; Loll et al., 2005), is the primary target of light-induced irreversible oxidative damage (Mattoo et al., 1981; Ohad et al., 1990). Because D1 can be damaged by even low light intensities, photosynthetic organisms cannot avoid photodamage (Tyystjärvi and Aro, 1996; for review, see Barber and Andersson, 1992). To overcome this, photosynthetic organisms have evolved an efficient PSII repair cycle, which involves disassembling PSII, degrading photodamaged D1, and replacing newly synthesized D1 (for review, see Baena-Gonzalez and Aro, 2002). The rate of photodamage is proportional to light energy. When the light intensity exceeds the repair capacity, damaged D1 accumulates, resulting in photoinhibition.In the PSII repair, recent studies in Synechocystis species PCC 6803 and Arabidopsis (Arabidopsis thaliana) suggest important roles of prokaryotic proteases (Lindahl et al., 1996, 2000; Bailey et al., 2002; Sakamoto et al., 2003; Silva et al., 2003; Komenda et al., 2006; Sun et al., 2007; Kapri-Pardes et al., 2007). Among them, FtsH appears to be a major protease. It is a membrane-anchored ATP-dependent zinc metalloprotease that belongs to the ATPases associated with a variety of cellular activities (AAA)+ protein family (for review, see Patel and Latterich, 1998; Ogura and Wilkinson, 2001). The ATPase and protease domain of FtsH was shown to form a hexameric-ring structure (Suno et al., 2006). In Synechocystis species PCC 6803, deletion of one of the thylakoidal FtsHs, slr0228, results in light-sensitive growth, impairment of the PSII repair cycle, and slower D1 degradation under high-light conditions (Silva et al., 2003). In Arabidopsis, 12 FtsH homologues were identified, nine of which are targeted to chloroplasts (Sakamoto et al., 2003). FtsH2 and FtsH5 are the most abundant among all chloroplastic FtsHs and are located in thylakoid membranes (Sakamoto et al., 2003; Yu et al., 2004, 2005). Chloroplastic FtsHs predominantly exist as a heterocomplex consisting of at least two types of isomers, A and B, represented by FtsH1/5 and FtsH2/8, respectively. These two types are functionally distinguishable from each other (Sakamoto et al., 2003; Yu et al., 2004, 2005; Zaltsman et al., 2005b).We have extensively studied Arabidopsis mutants lacking chloroplast FtsHs. A mutant lacking FtsH5 (called yellow variegated1 [var1]) or lacking FtsH2 (var2) is highly vulnerable to PSII photodamage under high light (Chen et al., 2000; Lindahl et al., 2000; Takechi et al., 2000; Sakamoto et al., 2002, 2004). One notable feature in these mutants, in addition to the defective PSII repair, is the leaf-variegated phenotype that displays two sectors in the same leaf (green sectors containing normal chloroplasts and white sectors containing abnormal plastids lacking thylakoid membranes; Supplemental Fig. S1). White sectors are made by living cells and appear to be comparable with green sectors, except for lacking photosynthetic proteins (Kato et al., 2007). These results demonstrated that white sectors in var2 are arrested in chloroplast development. It is thus proposed that FtsH is not only involved in PSII repair but also in the formation of thylakoid membranes. Moreover, a series of genetic studies enabled us to identify trans-acting mutations that suppressed leaf variegation in var2 (Park and Rodermel, 2004; Miura et al., 2007; Yu et al., 2008). Many suppressors appeared to be associated with chloroplast translation, suggesting that the formation of variegated sectors is not simply governed by a specific factor but rather by factors related to chloroplast development (Miura et al., 2007; Yu et al., 2008). Since the variegated phenotype complicates our biochemical study in Arabidopsis, unlike cyanobacteria, a combined usage of var and its suppressors is necessary to further investigate the role of FtsH in the PSII repair cycle.In this study, we show that chloroplasts in green sectors accumulate less PSII supercomplex and more PSII partial complexes than wild-type chloroplasts, likely due to the compromised PSII repair. Interestingly, we found that chloroplasts in green sectors accumulated significantly high levels of reactive oxygen species (ROS), suggesting that var2 indeed suffers from photooxidative stress. Given the essential role of FtsH, we evaluated impaired D1 degradation by the lack of FtsH2 and FtsH5 under different light conditions. Although a defect in degrading PSII reaction center proteins in var2 has been reported to occur under a photoinhibitory light condition (Bailey et al., 2002), D1 degradation examined in this study was very limited. This is because in vivo degradation of D1 is very difficult to measure in variegated leaf tissues (e.g. the presence of green and white leaf sectors interferes with protein normalization). To overcome the difficulty of handling variegated sectors, nonvariegated suppressor lines were subjected to these experiments. Our D1 degradation assays demonstrated that the lack of FtsH2 or FtsH5 significantly impairs D1 degradation. Collectively, our data corroborate important roles of FtsH2 and FtsH5 in avoiding photooxidative stress in chloroplasts.     

Plants Actively Avoid State Transitions upon Changes in Light Intensity: Role of Light-Harvesting Complex II Protein Dephosphorylation in High Light

Photosystem II (PSII) core and light-harvesting complex II (LHCII) proteins in plant chloroplasts undergo reversible phosphorylation upon changes in light intensity (being under control of redox-regulated STN7 and STN8 kinases and TAP38/PPH1 and PSII core phosphatases). Shift of plants from growth light to high light results in an increase of PSII core phosphorylation, whereas LHCII phosphorylation concomitantly decreases. Exactly the opposite takes place when plants are shifted to lower light intensity. Despite distinct changes occurring in thylakoid protein phosphorylation upon light intensity changes, the excitation balance between PSII and photosystem I remains unchanged. This differs drastically from the canonical-state transition model induced by artificial states 1 and 2 lights that concomitantly either dephosphorylate or phosphorylate, respectively, both the PSII core and LHCII phosphoproteins. Analysis of the kinase and phosphatase mutants revealed that TAP38/PPH1 phosphatase is crucial in preventing state transition upon increase in light intensity. Indeed, tap38/pph1 mutant revealed strong concomitant phosphorylation of both the PSII core and LHCII proteins upon transfer to high light, thus resembling the wild type under state 2 light. Coordinated function of thylakoid protein kinases and phosphatases is shown to secure balanced excitation energy for both photosystems by preventing state transitions upon changes in light intensity. Moreover, PROTON GRADIENT REGULATION5 (PGR5) is required for proper regulation of thylakoid protein kinases and phosphatases, and the pgr5 mutant mimics phenotypes of tap38/pph1. This shows that there is a close cooperation between the redox- and proton gradient-dependent regulatory mechanisms for proper function of the photosynthetic machinery.Photosynthetic light reactions take place in the chloroplast thylakoid membrane. Primary energy conversion reactions are performed by synchronized function of the two light energy-driven enzymes PSII and PSI. PSII uses excitation energy to split water into electrons and protons. PSII feeds electrons to the intersystem electron transfer chain (ETC) consisting of plastoquinone, cytochrome b₆f, and plastocyanin. PSI oxidizes the ETC in a light-driven reduction of NADP to NADPH. Light energy is collected by the light-harvesting antenna systems in the thylakoid membrane composed of specific pigment-protein complexes (light-harvesting complex I [LHCI] and LHCII). The majority of the light-absorbing pigments are bound to LHCII trimers that can serve the light harvesting of both photosystems (Galka et al., 2012; Kouřil et al., 2013; Wientjes et al., 2013b). Energy distribution from LHCII is regulated by protein phosphorylation (Bennett, 1979; Bennett et al., 1980; Allen et al., 1981) under control of the STN7 and STN8 kinases (Depège et al., 2003; Bellafiore et al., 2005; Bonardi et al., 2005; Vainonen et al., 2005) and the TAP38/PPH1 and Photosystem II Core Phosphatase (PBCP) phosphatases (Pribil et al., 2010; Shapiguzov et al., 2010; Samol et al., 2012). LHCII trimers are composed of LHCB1, LHCB2, and LHCB3 proteins, and in addition to reversible phosphorylation of LHCB1 and LHCB2, the protein composition of the LHCII trimers also affects the energy distribution from the light-harvesting system to photosystems (Damkjaer et al., 2009; Pietrzykowska et al., 2014). Most of the LHCII trimers are located in the PSII-rich grana membranes and PSII- and PSI-rich grana margins of the thylakoid membrane, and only a minor fraction resides in PSI- and ATP synthase-rich stroma lamellae (Tikkanen et al., 2008b; Suorsa et al., 2014). Both photosystems bind a small amount of LHCII trimers in biochemically isolatable PSII-LHCII and PSI-LHCII complexes (Pesaresi et al., 2009; Järvi et al., 2011; Caffarri et al., 2014). The large portion of the LHCII, however, does not form isolatable complexes with PSII or PSI, and therefore, it separates as free LHCII trimers upon biochemical fractionation of the thylakoid membrane by Suc gradient centrifugation or in native gel analyses (Caffarri et al., 2009; Järvi et al., 2011), the amount being dependent on the thylakoid isolation method. Nonetheless, in vivo, this major LHCII antenna fraction serves the light-harvesting function. This is based on the fact that fluorescence from free LHCII, peaking at 680 nm in 77-K fluorescence emission spectra, can only be detected when the energy transfer properties of the thylakoid membrane are disturbed by detergents (Grieco et al., 2015).Regulation of excitation energy distribution from LHCII to PSII and PSI has, for decades, been linked to LHCII phosphorylation and state transitions (Bennett, 1979; Bennett et al., 1980; Allen et al., 1981). It has been explained that a fraction of LHCII gets phosphorylated and migrates from PSII to PSI, which can be evidenced as increase in PSI cross section and was assigned as transition to state 2 (for review, see Allen, 2003; Rochaix et al., 2012). The LHCII proteins are, however, phosphorylated all over the thylakoid membrane (i.e. in the PSII- and LHCII-rich grana core) in grana margins containing PSII, LHCII, and PSI as well as in PSI-rich stroma lamellae also harboring PSII-LHCII, LHCII, and PSI-LHCII complexes in minor amounts (Tikkanen et al., 2008b; Grieco et al., 2012; Leoni et al., 2013; Wientjes et al., 2013a)—making the canonical-state transition theory inadequate to explain the physiological role of reversible LHCII phosphorylation (Tikkanen and Aro, 2014). Moreover, the traditional-state transition model is based on lateral segregation of PSII-LHCII and PSI-LHCI to different thylakoid domains. It, however, seems likely that PSII and PSI are energetically connected through a shared light-harvesting system composed of LHCII trimers (Grieco et al., 2015), and there is efficient excitation energy transfer between the two photosystems (Yokono et al., 2015). Nevertheless, it is clear that LHCII phosphorylation is a prerequisite to form an isolatable PSI-LHCII complex called the state transition complex (Pesaresi et al., 2009; Järvi et al., 2011). Existence of a minor state transition complex, however, does not explain why LHCII is phosphorylated all over the thylakoid membrane and how the energy transfer is regulated from the majority of LHCII antenna that is shared between PSII and PSI but does not form isolatable complexes with them (Grieco et al., 2015).Plants grown under any steady-state white light condition show the following characteristics of the thylakoid membrane: PSII core and LHCII phosphoproteins are moderately phosphorylated, phosphorylation takes place all over the thylakoid membrane, and the PSI-LHCII state transition complex is present (Järvi et al., 2011; Grieco et al., 2012; Wientjes et al., 2013b). Upon changes in the light intensity, the relative phosphorylation level between PSII core and LHCII phosphoproteins drastically changes (Rintamäki et al., 1997, 2000) in the timescale of 5 to 30 min. When light intensity increases, the PSII core protein phosphorylation increases, whereas the level of LHCII phosphorylation decreases. On the contrary, a decrease in light intensity decreases the phosphorylation level of PSII core proteins but strongly increases the phosphorylation of the LHCII proteins (Rintamäki et al., 1997, 2000). The presence and absence of the PSI-LHCII state transition complex correlate with LHCII phosphorylation (similar to the state transitions; Pesaresi et al., 2009; Wientjes et al., 2013b). Despite all of these changes in thylakoid protein phosphorylation, the relative excitation of PSII and PSI (i.e. the absorption cross section of PSII and PSI measured by 77-K fluorescence) remains nearly unchanged upon changes in white-light intensity (i.e. no state transitions can be observed despite massive differences in LHCII protein phosphorylation; Tikkanen et al., 2010).The existence of the opposing behaviors of PSII core and LHCII protein phosphorylation, as described above, has been known for more than 15 years (Rintamäki et al., 1997, 2000), but the physiological significance of this phenomenon has remained elusive. It is known that PSII core protein phosphorylation in high light (HL) facilitates the unpacking of PSII-LHCII complexes required for proper processing of the damaged PSII centers and thus, prevents oxidative damage of the photosynthetic machinery (Tikkanen et al., 2008a; Fristedt et al., 2009; Goral et al., 2010; Kirchhoff et al., 2011). It is also known that the damaged PSII core protein D1 needs to be dephosphorylated before its proteolytic degradation upon PSII turnover (Koivuniemi et al., 1995). There is, however, no coherent understanding available to explain why LHCII proteins are dephosphorylated upon exposure of plants to HL and PSII core proteins are dephosphorylated upon exposure to low light (LL).The above-described light quantity-dependent control of thylakoid protein phosphorylation drastically differs from the light quality-dependent protein phosphorylation (Tikkanen et al., 2010). State transitions are generally investigated by using different light qualities, preferentially exciting either PSI or PSII. State 1 light favors PSI excitation, leading to oxidation of the ETC and dephosphorylation of both the PSII core and LHCII proteins. State 2 light, in turn, preferentially excites PSII, leading to reduction of ETC and strong concomitant phosphorylation of both the PSII core and LHCII proteins (Haldrup et al., 2001). Shifts between states 1 and 2 lights induce state transitions, mechanisms that change the excitation between PSII and PSI (Murata and Sugahara, 1969; Murata, 2009). Similar to shifts between state lights, the shifts between LL and HL intensity also change the phosphorylation of the PSII core and LHCII proteins (Rintamäki et al., 1997, 2000). Importantly, the white-light intensity-induced changes in thylakoid protein phosphorylation do not change the excitation energy distribution between the two photosystems (Tikkanen et al., 2010). Despite this fundamental difference between the light quantity- and light quality-induced thylakoid protein phosphorylations, a common feature for both mechanisms is a strict requirement of LHCII phosphorylation for formation of the PSI-LHCII complex. However, it is worth noting that LHCII phosphorylation under state 2 light is not enough to induce the state 2 transition but that the P-LHCII docking proteins in the PSI complex are required (Lunde et al., 2000; Jensen et al., 2004; Zhang and Scheller, 2004; Leoni et al., 2013).Thylakoid protein phosphorylation is a dynamic redox-regulated process dependent on the interplay between two kinases (STN7 and STN8; Depège et al., 2003; Bellafiore et al., 2005; Bonardi et al., 2005; Vainonen et al., 2005) and two phosphatases (TAP38/PPH1 and PBCP; Pribil et al., 2010; Shapiguzov et al., 2010; Samol et al., 2012). Concerning the redox regulation mechanisms in vivo, only the LHCII kinase (STN7) has so far been thoroughly studied (Vener et al., 1997; Rintamäki et al., 2000; Lemeille et al., 2009). The STN7 kinase is considered as the LHCII kinase, and indeed, it phosphorylates the LHCB1 and LHCB2 proteins (Bellafiore et al., 2005; Bonardi et al., 2005; Tikkanen et al., 2006). In addition to this, STN7 takes part in the phosphorylation of PSII core proteins (Vainonen et al., 2005), especially in LL (Tikkanen et al., 2008b, 2010). The STN8 kinase is required for phosphorylation of PSII core proteins in HL but does not significantly participate in phosphorylation of LHCII (Bellafiore et al., 2005; Bonardi et al., 2005; Vainonen et al., 2005; Tikkanen et al., 2010). It has been shown that, in traditional state 1 condition, which oxidizes the ETC, the dephosphorylation of LHCII is dependent on TAP38/PPH1 phosphatase (Pribil et al., 2010; Shapiguzov et al., 2010), whereas the PSII core protein dephosphorylation is dependent on the PBCP phosphatase (Samol et al., 2012). However, it remains unresolved whether and how the TAP38/PPH1 and PBCP phosphatases are involved in the light intensity-dependent regulation of thylakoid protein phosphorylation typical for natural environments.Here, we have used the two kinase (stn7 and stn8) and the two phosphatase (tap38/pph1and pbcp) mutants of Arabidopsis (Arabidopsis thaliana) to elucidate the individual roles of these enzymes in reversible thylakoid protein phosphorylation and distribution of excitation energy between PSII and PSI upon changes in light intensity. It is shown that the TAP38/PPH1-dependent, redox-regulated LHCII dephosphorylation is the key component to maintain excitation balance between PSII and PSI upon increase in light intensity, which at the same time, induces strong phosphorylation of the PSII core proteins. Collectively, reversible but opposite phosphorylation and dephosphorylation of the PSII core and LHCII proteins upon increase or decrease in light intensity are shown to be crucial for maintenance of even distribution of excitation energy to both photosystems, thus preventing state transitions. Moreover, evidence is provided indicating that the pH gradient across the thylakoid membrane is yet another important component in regulation of the distribution of excitation energy to PSII and PSI, possibly by affecting the regulation of thylakoid kinases and phosphatases.     

The Stromal Chloroplast Deg7 Protease Participates in the Repair of Photosystem II after Photoinhibition in Arabidopsis

Light is the ultimate source of energy for photosynthesis; however, excessive light leads to photooxidative damage and hence reduced photosynthetic efficiency, especially when combined with other abiotic stresses. Although the photosystem II (PSII) reaction center D1 protein is the primary target of photooxidative damage, other PSII core proteins are also damaged and degraded. However, it is still largely unknown whether degradation of D1 and other PSII proteins involves previously uncharacterized proteases. Here, we show that Deg7 is peripherally associated with the stromal side of the thylakoid membranes and that Deg7 interacts directly with PSII. Our results show that Deg7 is involved in the primary cleavage of photodamaged D1, D2, CP47, and CP43 and that this activity is essential for its function in PSII repair. The double mutants deg5 deg7 and deg8 deg7 showed no obvious phenotypic differences under normal growth conditions, but additive effects were observed under high light. These results suggest that Deg proteases on both the stromal and luminal sides of the thylakoid membranes are important for the efficient PSII repair in Arabidopsis (Arabidopsis thaliana).Chloroplasts of higher plants carry out one of the most important biochemical reactions: the capture of light energy and its conversion into chemical energy. Although light is the ultimate source of energy for photosynthesis, it can also be harmful to plants. Light-induced loss of photosynthetic efficiency, which is generally termed as photoinhibition, limits plant growth and lowers productivity, especially when combined with other abiotic stresses.The main target of photoinhibition is PSII, which catalyzes the light-dependent water oxidation concomitantly with oxygen production (for review, see Prasil et al., 1992; Aro et al., 1993; Adir et al., 2003). In higher plants, PSII consists of more than 20 subunits, including the reaction center D1 and D2 proteins, cytochrome (Cyt) b₅₅₉, the light-harvesting chlorophyll a-binding proteins CP47 and CP43, the oxygen-evolving 33-kD protein (PsbO), and several low molecular mass proteins (Nelson and Yocum, 2006). The PSII reaction center D1 protein has been identified among PSII proteins as the primary target of light-induced damage (Kyle et al., 1984; Mattoo et al., 1984; Ohad et al., 1984; Adir et al., 1990), but several studies have shown that the D2, CP47, and CP43 proteins are degraded under photoinhibitory conditions (Schuster et al., 1988; Yamamoto and Akasaka, 1995; Jansen et al., 1999; Adir et al., 2003). Moreover, several small PSII subunits, such as PsbH, PsbW, and Cyt b₅₅₉, were also found to be frequently replaced within PSII (Hagman et al., 1997; Ortega et al., 1999; Bergantino et al., 2003). Evidence for the involvement of two families of proteases, FtsH and Deg, in the degradation of the D1 protein in thylakoids of higher plants has been recently described (Lindahl et al., 1996, 2000; Bailey et al., 2002; Sakamoto et al., 2003; Silva et al., 2003; Kapri-Pardes et al., 2007; Sun et al., 2007a, 2007b). However, it is still largely unknown whether degradation of D1 and other PSII proteins involves previously uncharacterized proteases.DegP (or HtrA) proteases were initially identified based on the fact that they are required for the survival of Escherichia coli at high temperatures and for the degradation of abnormal periplasmic proteins (Lipinska et al., 1988; Strauch and Beckwith, 1988). DegP is an ATP-independent Ser endopeptidase, and it contains a trypsin-like protease domain at the N terminus, followed by two PDZ domains (Gottesman, 1996; Pallen and Wren, 1997; Clausen et al., 2002). PDZ domains appear to be important for complex assembly and substrate binding through three or four residues in the C terminus of their target proteins (Doyle et al., 1996; Harris and Lim, 2001). DegP switches between chaperone and protease functions in a temperature-dependent manner. The chaperone function dominates at low temperatures, and DegP becomes proteolytically active at elevated temperatures (Spiess et al., 1999). Crystal structures of different members of the DegP protein family (Krojer et al., 2002; Li et al., 2002; Kim et al., 2003; Wilken et al., 2004) have revealed the structure-function relationship of these PDZ-containing proteases. Trimeric DegP is the functional unit, and the hexameric DegP is formed via the staggered association of trimers (Clausen et al., 2002; Kim and Kim, 2005). At normal growth temperatures, the active site of the protease is located within the chamber of hexameric DegP, which is not accessible to the substrates. However, at high temperatures, conformational changes induce the activation of the protease function (Krojer et al., 2002). Recent studies have shed light on the substrate binding-induced formation of larger oligomeric complexes of DegP (Jiang et al., 2008; Krojer et al., 2008).In Arabidopsis (Arabidopsis thaliana), 16 genes coding for DegP-like proteases have been identified, and at least seven gene products are predicted to be located in chloroplasts (Kieselbach and Funk, 2003; Huesgen et al., 2005; Adam et al., 2006; Sakamoto, 2006; Kato and Sakamoto, 2009). Based on proteomic data, four Deg proteases have been shown to be localized to the chloroplast (Peltier et al., 2002; Schubert et al., 2002) and functionally characterized. Deg1, Deg5, and Deg8 are located in thylakoid lumen, and Deg2 is peripherally associated with the stromal side of thylakoid membranes (Itzhaki et al., 1998; Haußühl et al., 2001; Sun et al., 2007a). Recombinant DegP1, now renamed Deg1, has been shown to be proteolytically active toward thylakoid lumen proteins such as plastocyanin and PsbO of PSII in vitro (Chassin et al., 2002). A 5.2-kD C-terminal fragment of the D1 protein was detected in vitro after incubation of recombinant Deg1 with inside-out thylakoid membranes. In transgenic plants with reduced levels of Deg1, fewer of its 16- and 5.2-kD degradation products were observed (Kapri-Pardes et al., 2007). Deg5 and Deg8 form a dodecameric complex in the thylakoid lumen, and recombinant Deg8 is able to degrade the photodamaged D1 protein of PSII in an in vitro assay (Sun et al., 2007a). The 16-kD N-terminal degradation fragment of the D1 protein was detected in wild-type plants but not in a deg5 deg8 double mutant after high-light treatment. The deg5 deg8 double mutant showed increased sensitivity to high light and high temperature in terms of growth and PSII activity compared with the single mutants deg5 and deg8, suggesting that Deg5 and Deg8 have overlapping functions in the primary cleavage of the CD loop of the D1 protein (Sun et al., 2007a, 2007b). In vitro analysis has demonstrated that recombinant stroma-localized Deg2 was also shown to be involved in the primary cleavage of the DE loop of the D1 protein (Haußühl et al., 2001). However, analysis of a mutant lacking Deg2 suggested that Deg2 may not be involved in D1 degradation in vivo (Huesgen et al., 2006).Here, we have expressed and purified a recombinant DegP protease, His-Deg7. In vitro experiments showed that His-Deg7 is proteolytically active toward the PSII proteins D1, D2, CP43, and CP47. In vivo analyses of a deg7 mutant revealed that the mutant is more sensitive to high light stress than the wild-type plants. We demonstrated that Deg7 is a chloroplast stroma protein associated with the thylakoid membranes and that it interacts with PSII, which suggests that it can cleave the stroma-exposed region of substrate proteins. Our results also provide evidence that Deg7 is important for maintaining PSII function.     

TEF30 Interacts with Photosystem II Monomers and Is Involved in the Repair of Photodamaged Photosystem II in Chlamydomonas reinhardtii

The remarkable capability of photosystem II (PSII) to oxidize water comes along with its vulnerability to oxidative damage. Accordingly, organisms harboring PSII have developed strategies to protect PSII from oxidative damage and to repair damaged PSII. Here, we report on the characterization of the THYLAKOID ENRICHED FRACTION30 (TEF30) protein in Chlamydomonas reinhardtii, which is conserved in the green lineage and induced by high light. Fractionation studies revealed that TEF30 is associated with the stromal side of thylakoid membranes. By using blue native/Deriphat-polyacrylamide gel electrophoresis, sucrose density gradients, and isolated PSII particles, we found TEF30 to quantitatively interact with monomeric PSII complexes. Electron microscopy images revealed significantly reduced thylakoid membrane stacking in TEF30-underexpressing cells when compared with control cells. Biophysical and immunological data point to an impaired PSII repair cycle in TEF30-underexpressing cells and a reduced ability to form PSII supercomplexes after high-light exposure. Taken together, our data suggest potential roles for TEF30 in facilitating the incorporation of a new D1 protein and/or the reintegration of CP43 into repaired PSII monomers, protecting repaired PSII monomers from undergoing repeated repair cycles or facilitating the migration of repaired PSII monomers back to stacked regions for supercomplex reassembly.Oxygenic photosynthesis is essential for almost all life on Earth, as it provides the reduced carbon and the oxygen required for respiration. A key enzyme in oxygenic photosynthesis is PSII, which catalyzes the light-driven oxidation of water. The core of PSII in algae and land plants contains D1 (PsbA), D2 (PsbD), CP43 (PsbC), CP47 (PsbB), the α-subunit (PsbE) and β-subunit (PsbF) of cytochrome b₅₅₉, as well as several intrinsic low-molecular-mass subunits. The core monomer is associated with the extrinsic oxygen-evolving complex (OEC) consisting of OEE1 (PSBO), OEE2 (PSBP), and OEE3 (PSBQ), which stabilize the inorganic Mn₄O₅Ca cluster required for water oxidation (for review, see Pagliano et al., 2013). PSII core monomers assemble into dimers to which, at both sides, light-harvesting proteins (LHCII) bind to form PSII supercomplexes. In land plants, each PSII dimer binds two each of the monomeric minor LHCII proteins CP24, CP26, and CP29 in addition to up to four major LHCII trimers (Caffarri et al., 2009; Kouřil et al., 2011). Biochemical evidence suggests that, in the thylakoid membrane, up to eight LHCII trimers can be present per PSII core dimer, presumably because of the existence of a pool of extra LHCII (Kouřil et al., 2013). In Chlamydomonas reinhardtii, lacking CP24, each PSII dimer binds two each of the CP26 and CP29 monomers as well as up to six major LHCII trimers (Tokutsu et al., 2012). The reaction center proteins D1 and D2 bind all the redox-active cofactors required for PSII electron transport (Umena et al., 2011). Light captured by the internal antenna proteins CP43 and CP47 and the outer antenna induces charge separation in PSII, which in turn enables the OEC to oxidize water and provide electrons to the electron transfer chain. In land plants and green algae, PSII supercomplexes are localized to stacked regions of the thylakoid membranes, while the synthesis of PSII cores is considered to take place in stroma lamellae.A particular feature of PSII is its vulnerability to light, with the D1 protein being a target of light-induced damage and the damage being proportional to the photon flux density (PFD) applied (Tyystjärvi and Aro, 1996). To cope with this damage, an elaborate, highly conserved repair mechanism has evolved termed the PSII repair cycle, during which damaged PSII complexes are partially disassembled and the defective D1 protein is replaced by a de novo synthesized copy (for review, see Nixon et al., 2010; Komenda et al., 2012; Mulo et al., 2012; Nath et al., 2013a; Nickelsen and Rengstl, 2013; Tyystjärvi, 2013; Järvi et al., 2015). Photodamage occurs at all light intensities, but when the rate of damage exceeds the capacity for repair, photoinhibition is manifested as a decrease in the proportion of active PSII reaction centers (Aro et al., 1993). While PSII photodamage occurs in the supercomplexes in the stacked membrane regions, the replacement of damaged D1 takes place in stroma lamellae (Aro et al., 2005). Thus, the PSII repair cycle requires the lateral migration of PSII complexes, which is impaired by the macromolecular crowding in stacked thylakoid membranes (Kirchhoff, 2014). Lateral migration of damaged PSII complexes is facilitated by thylakoid membrane unfolding and PSII supercomplex disassembly. Both processes are enhanced by the phosphorylation of the PSII core subunits D1, D2, CP43, and PsbH, which is mainly mediated by the protein kinase STATE TRANSITION8 (STN8; Tikkanen et al., 2008; Fristedt et al., 2009; Herbstová et al., 2012; Nath et al., 2013b; Wunder et al., 2013). Efficient PSII supercomplex disassembly also requires the THYLAKOID FORMATION1 (THF1)/NON-YELLOW COLORING4 (NYC4)/Psb29 protein (Huang et al., 2013; Yamatani et al., 2013). After the migration of PSII monomers to unstacked thylakoid regions, PSII core subunits are dephosphorylated by the PSII core phosphatase PBCP (Samol et al., 2012), which is required for the efficient degradation of D1 (Koivuniemi et al., 1995; Rintamäki et al., 1996; Kato and Sakamoto, 2014). Degradation of D1 is subsequently realized by the membrane-integral FtsH protease (Lindahl et al., 2000; Silva et al., 2003) and by lumenal and stromal Deg proteases (Haussühl et al., 2001; Kapri-Pardes et al., 2007; Sun et al., 2010). Degradation is assisted by the THYLAKOID LUMEN PROTEIN18.3 (TLP18.3), presumably by its phosphatase activity and ability to interact with lumenal Deg1 (Sirpiö et al., 2007; Wu et al., 2011; Zienkiewicz et al., 2012). D1 proteolysis follows the partial disassembly of the PSII complex, during which CP43 and low-molecular-mass subunits are released to generate a CP43-free PSII monomer (Aro et al., 2005). Thereafter, a newly synthesized D1 copy is cotranslationally inserted from a plastidial 70S ribosome into the thylakoid membrane and processed by the CARBOXYL TERMINAL PEPTIDASE A (CTPA; Zhang et al., 1999, 2000; Che et al., 2013). In Arabidopsis (Arabidopsis thaliana), the D1 synthesis rate appears to be negatively regulated by the PROTEIN DISULFIDE ISOMERASE6 (PDI6; Wittenberg et al., 2014). Moreover, yet unknown steps during PSII repair require the stromal cyclophilin ROTAMASE CYP4 and stromal HEAT SHOCK PROTEIN70 (Schroda et al., 1999; Yokthongwattana et al., 2001; Cai et al., 2008). The PSII repair cycle is completed by the reassembly of the CP43 protein, ligation of the OEC, back migration of PSII to stacked membrane regions, and supercomplex formation. Except for CtpA, all mentioned factors appear to be specific for PSII repair, while many more auxiliary factors play roles in PSII de novo synthesis and repair (for review, see Järvi et al., 2015).In this study, we report on the functional characterization of the THYLAKOID ENRICHED FRACTION30 (TEF30) protein in C. reinhardtii. In this organism, TEF30 was first identified in a proteomics study on isolated thylakoid membranes (Allmer et al., 2006). TEF30 attracted our attention because its abundance increased 1.7-fold in membrane-enriched fractions of C. reinhardtii cells that had been shifted from 41 to 145 µmol photons m⁻² s⁻¹ for 8 h (Mettler et al., 2014; Supplemental Fig. S1). The TEF30 ortholog in Arabidopsis M-ENRICHED THYLAKOID PROTEIN1 (MET1; where M stands for mesophyll cells) was functionally characterized only recently (Bhuiyan et al., 2015). Both MET1 and TEF30 interact quantitatively with monomeric PSII core particles at the stroma side of the thylakoid membranes and play a role in the assembly of PSII monomers and/or their migration to stacked membrane regions for supercomplex assembly. While MET1 appears to exert this function during PSII de novo biogenesis and during the PSII repair cycle in Arabidopsis, TEF30 appears to function exclusively during PSII repair in C. reinhardtii.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

Photosystem II and Pigment Dynamics among Ecotypes of the Green Alga Ostreococcus

We investigated the photophysiological responses of three ecotypes of the picophytoplankter Ostreococcus and a larger prasinophyte Pyramimonas obovata to a sudden increase in light irradiance. The deepwater Ostreococcus sp. RCC809 showed very high susceptibility to primary photoinactivation, likely a consequence of high oxidative stress, which may relate to the recently noted plastid terminal oxidase activity in this strain. The three Ostreococcus ecotypes were all capable of deploying modulation of the photosystem II repair cycle in order to cope with the light increase, but the effective clearance of photoinactivated D1 protein appeared to be slower in the deepwater Ostreococcus sp. RCC809, suggesting that this step is rate limiting in the photosystem II repair cycle in this strain. Moreover, the deepwater Ostreococcus accumulated lutein and showed substantial use of the xanthophyll cycle under light stress, demonstrating its high sensitivity to light fluctuations. The sustained component of the nonphotochemical quenching of fluorescence correlated well with the xanthophyll deepoxidation activity. Comparisons with the larger prasinophyte P. obovata suggest that the photophysiology of Ostreococcus ecotypes requires high photosystem II repair rates to counter a high susceptibility to photoinactivation, consistent with low pigment package effects in their minute-sized cells.The prasinophytes are marine planktonic green algae with a phylogenetic position branching near the base of the green lineage (Baldauf, 2003; Turmel et al., 2009). They are widespread in temperate (Diez et al., 2001; Zhu et al., 2005) and polar (Lovejoy et al., 2007) marine habitats, in which they are often significant contributors to primary production (Not et al., 2004). The prasinophytes include the smallest known eukaryotic photoautotroph, Ostreococcus tauri (Courties et al., 1994; Chrétiennot-Dinet et al., 1995), whose particularly simple structure makes it an attractive model minimal chlorophyte, and indeed, minimal eukaryote (Derelle et al., 2006). Recently, genomic sequences for three Ostreococcus strains, isolated from different ecological niches, have become available (Derelle et al., 2006; Palenik et al., 2007), thus increasing the interest of these models for understanding acclimation processes in this deep-branching group of chlorophytes.Photoacclimation strategies differ in two Ostreococcus strains (Cardol et al., 2008; Six et al., 2008), which, although belonging to different phylogenetic clades, are nonetheless morphologically indistinguishable (Rodríguez et al., 2005). O. tauri, a eutrophilic lagoon species, modulates PSII content to enable acclimation and growth over a wide range of irradiances. In marked contrast, Ostreococcus sp. (O. sp.) RCC809, isolated at 105 m depth in the tropical Atlantic Ocean, modulates the size of its large PSII antenna in a strategy that accommodates a narrower range of light levels but that incurs lower nutrient costs compared with photoacclimation in O. tauri (Six et al., 2008). The evidence for different light acclimation strategies between these two Ostreococcus ecotypes raises the question of the underlying physiological processes for niche adaptation in these closely related organisms. Cardol et al. (2008) recently analyzed the coastal O. tauri and the deepwater O. sp. RCC809 grown under low to moderate light and found exciting evidence for a plastid terminal oxidase electron flow path in O. sp. RCC809 from PSII back to oxygen, short-circuiting the usual Z scheme in a mechanism to generate a transthylakoidal pH gradient without net generation of reductant. Like all oxygenic photoautotrophs, the prasinophytes suffer photoinactivation of PSII (Aro et al., 1993; Tyystjarvi, 2008; Guskov et al., 2009) at a rate approximately proportional to the incident irradiance (Nagy et al., 1995; Hakala et al., 2005). To counter this photoinactivation, a PSII repair cycle proteolytically removes the photoinactivated D1 protein (Silva et al., 2003) and replaces it through de novo synthesis and reassembly with the remaining subunits (Aro et al., 1993). If photoinactivation outruns the rate of repair, the PSII pool suffers net photoinhibition (Aro et al., 2005; Nishiyama et al., 2005, 2006; Murata et al., 2007), leading to a decrease in photosynthetic capacity and potentially to a decrease in growth. To limit photoinhibition, photosynthetic cells use physiological processes that dissipate excess light energy into heat, thereby preempting the generation of toxic reactive oxygen species (Baroli et al., 2004; Holt et al., 2004) that can inhibit metabolism, notably including the PSII repair processes (Nishiyama et al., 2006; Murata et al., 2007). These excitation dissipation mechanisms manifest as a drop in PSII fluorescence yield termed nonphotochemical quenching of fluorescence (NPQ). In land plants and characterized chlorophytes, NPQ is notably associated with changes in light-harvesting complex conformation along with pigmentation changes (Demmig-Adams and Adams, 1992; Baroli et al., 2004; Holt et al., 2004; Li et al., 2004).The specialization of Ostreococcus ecotypes to contrasting environments suggests that they may have evolved distinct capacities to cope with rapid fluctuations in light. Here, we investigate this question by subjecting three different Ostreococcus ecotypes to short-term increases in light irradiance to uncover their capacities for PSII repair and susceptibilities to photoinactivation. We use a target theory approach (Nagy et al., 1995; Sinclair et al., 1996) to parameterize their susceptibility to primary photoinactivation in a form useful for predicting and modeling responses to changes in irradiance. We moreover compare the Ostreococcus strains to a much larger prasinophyte derived from temperate surface waters, Pyramimonas obovata, to explore how cell size can influence photophysiology in the prasinophytes.     

Kinetic and Spectral Resolution of Multiple Nonphotochemical Quenching Components in Arabidopsis Leaves

Using novel specially designed instrumentation, fluorescence emission spectra were recorded from Arabidopsis (Arabidopsis thaliana) leaves during the induction period of dark to high-light adaptation in order to follow the spectral changes associated with the formation of nonphotochemical quenching. In addition to an overall decrease of photosystem II fluorescence (quenching) across the entire spectrum, high light induced two specific relative changes in the spectra: (1) a decrease of the main emission band at 682 nm relative to the far-red (750–760 nm) part of the spectrum (Δ F₆₈₂); and (2) an increase at 720 to 730 nm (Δ F₇₂₀) relative to 750 to 760 nm. The kinetics of the two relative spectral changes and their dependence on various mutants revealed that they do not originate from the same process but rather from at least two independent processes. The Δ F₇₂₀ change is specifically associated with the rapidly reversible energy-dependent quenching. Comparison of the wild-type Arabidopsis with mutants unable to produce or overexpressing the PsbS subunit of photosystem II showed that PsbS was a necessary component for Δ F₇₂₀. The spectral change Δ F₆₈₂ is induced both by energy-dependent quenching and by PsbS-independent mechanism(s). A third novel quenching process, independent from both PsbS and zeaxanthin, is activated by a high turnover rate of photosystem II. Its induction and relaxation occur on a time scale of a few minutes. Analysis of the spectral inhomogeneity of nonphotochemical quenching allows extraction of mechanistically valuable information from the fluorescence induction kinetics when registered in a spectrally resolved fashion.One of the most important photoprotective mechanisms against high-light (HL) stress in photosynthetic organisms is the nonphotochemical quenching (NPQ) of excitation energy, which is mostly due to thermal deactivation of pigment excited states in the antenna of PSII. There exist a number of literature reviews on the subject (Demmig-Adams and Adams, 1992; Horton et al., 1996; Horton and Ruban, 1999, 2005; Niyogi, 1999, 2000; Müller et al., 2001; Golan et al., 2004; Krause and Jahns, 2004). Chlorophyll (Chl) fluorescence, and in particular pulse amplitude-modulated (PAM) fluorometry as introduced by Schreiber et al. (1986), has become by far the dominant technique to measure NPQ in leaves, chloroplasts, and intact microorganisms (Krause and Weis, 1991; Govindjee, 1995; Maxwell and Johnson, 2000; Krause and Jahns, 2003; Schreiber, 2004), more recently often combined with specific NPQ mutant studies (Golan et al., 2004; Kalituho et al., 2006, 2007; Dall''Osto et al., 2007). In this technique, periodic saturating light pulses are applied, superimposed on the continuous actinic irradiation applied to induce NPQ, in order to transiently close the PSII reaction centers (RCs). Since the photochemistry contribution (photochemical quenching) is thus brought to zero, the method allows us to follow the dynamics of the NPQ development and relaxation by fluorescence in a relatively simple manner (Krause and Jahns, 2003, 2004).Mostly based on its relaxation kinetics, NPQ has been divided technically into the three kinetic components qE, qT, and qI, for the rapid, middle, and slow phases of relaxation (Horton and Hague, 1988), initially attributed to energy-dependent quenching, state transitions, and photoinhibitory quenching (Quick and Stitt, 1989). The rapidly forming and reversible part of NPQ, qE, is the most thoroughly studied. It is well established that this type of quenching is a finely regulated process in which the main governing factors are the proton gradient across the chloroplast thylakoid membrane, Δ pH (Wraight and Crofts, 1970; Briantais et al., 1979), the xanthophyll cycle (i.e. conversion of violaxanthin to antheraxanthin and zeaxanthin [Zx]; Demmig et al., 1987; Demmig-Adams, 1990; Demmig-Adams and Adams, 1992), and the action of the PsbS protein (Funk et al., 1995; Li et al., 2000, 2004; Niyogi et al., 2005). The actual molecular mechanism is still unknown, although there is no shortage of hypotheses and proposed quencher candidates: energy transfer from Chl to Zx in the major light-harvesting complex (LHCII; Frank et al., 2000); electron transfer from a carotenoid to Chl forming a Zx-Chl or lutein-Chl charge-transfer state (Holt et al., 2005; Avenson et al., 2009); direct or indirect quenching by the PsbS protein (Li et al., 2000; Niyogi et al., 2005); energy transfer from Chl to lutein in LHCII (Horton et al., 1991; Ruban et al., 2007) linked to the aggregation of or a conformational change in LHCII; and last but not least, a far-red (FR) light-emitting quenched Chl-Chl charge-transfer state formed by the aggregation of LHCII (Miloslavina et al., 2008). Quenching in the PSII RC has also been proposed (Weis and Berry, 1987; Finazzi et al., 2004; Huner et al., 2005; Ivanov et al., 2008) as an additional type of Zx-independent quenching. Alternatively, it has been suggested that quenching by lutein can complement the Zx-dependent quenching (Niyogi et al., 2001; Li et al. 2009). Johnson et al. (2009) have recently given support to the notion that both Zx-dependent and Zx-independent quenching originate from the same PsbS-dependent mechanism, which is modulated by Zx (Crouchman et al., 2006).While the rapidly relaxing phase qE is now well characterized in its dependence on the various factors, the much slower qT and qI phases are still controversial, and each of them may have contributions from more than one mechanism. The qI component has been traditionally attributed to photoinhibition of PSII (Somersalo and Krause, 1988), associated with coordinated degradation and repair of the photosystem (Powles and Björkman, 1982; Kyle, 1987; Krause, 1988; Aro et al., 1993; Long et al., 1994; Murata et al., 2007). Lately, though, it is more widely accepted that under most conditions the photoinhibition is low and qI, like qE, is a result of thermal deactivation of excited states. Different hypotheses have been put forward to account for its seeming irreversibility: persistent transmembrane Δ pH (Gilmore and Yamamoto, 1992), stable protonation of proteins (Horton et al., 1994), accumulation of inactive PSII reaction centers (Briantais et al., 1992; Schansker and van Rensen, 1999), or stable binding of Zx to CP29 (Färber et al., 1997). The connection of the qT phase with state transitions has been doubted as well, and in fact it is now thought that the fraction of energy redistributed from PSI to PSII under high-light conditions is negligible (Walters and Horton, 1991, 1993) and that the qT must have a different origin or that it has erroneously been ascribed as NPQ (Schansker et al., 2006).Along with the large amount of contradictory evidence on the nature and location of the NPQ quenching site(s), the question of whether the light-induced reversible NPQ represents one single mechanism of deexcitation located in a single site brought about by the combined action of PsbS and Zx (Johnson et al., 2009) or whether it comprises several parallel and largely independent mechanisms acting on different parts of the PSII antenna has not been finally answered. One way to answer this question might be to carefully examine the spectral properties of NPQ-related fluorescence changes. Quenching in different locations of the PSII antenna or with different mechanisms might give rise to a differential quenching in various parts of the PSII antenna that might affect the PSII fluorescence spectra in different ways. This appears possible, since the various pigment-protein complexes of the photosynthetic apparatus have slightly different absorption and emission spectra (Holzwarth, 1991; Holzwarth and Roelofs, 1992). However, in the vast majority of modulated Chl fluorescence instrumentation, including the most widely used PAM fluorometer (Schreiber et al., 1986), the signal is integrated over a broad wavelength range, usually covering the whole range of 710 nm or greater. This integration over the long-wave part of the spectrum has several undesirable consequences and is associated with the unnecessary loss of available information. For example, the fluorescence of PSII peaks in the region of 680 to 685 nm, whereas beyond 700 nm, the PSII fluorescence intensity drops to less than 20% of its peak intensity. In contrast, the fluorescence of intact PSI complexes is dominant in the region above 710 nm (Haehnel et al., 1982; Karukstis and Sauer, 1983; Holzwarth et al., 1985; Holzwarth, 1986; Slavov et al., 2008). Thus, the widely used instrumentation measures the NPQ parameters in a region with reduced PSII contribution and relatively high PSI contribution to total fluorescence, despite the fact that NPQ is generally considered to be primarily a PSII phenomenon. Only in a few studies has the fluorescence in the red and the FR region been separated in order to evaluate the contribution of PSI and its influence on the NPQ parameters (Genty et al., 1990; Peterson et al., 2001). NPQ might also shift the fluorescence properties of the PSII antenna complexes or give rise to entirely new fluorescing components (Miloslavina et al., 2008). This would remain undetected if the NPQ fluorescence changes are not resolved in the spectral domain. It follows from these considerations that a great deal of insight into the NPQ mechanisms and locations may be gained if the spectral dimension is added to the NPQ fluorescence characterization. Among the many advantages of such an approach, one would then be able to distinguish whether NPQ simply leads to a uniform decrease of PSII fluorescence across the emission range or whether this decrease is nonuniform, localized in specific pigment protein complexes, and/or whether new fluorescing species are actually being produced in the NPQ process.The HL-induced NPQ effects on the leaf fluorescence spectra have often been studied also at low temperature, where the differentiation between pigment sites is better (Krause et al., 1983; Demmig and Björkman, 1987; Ruban and Horton, 1994). However, the possibility to resolve the kinetics of NPQ development and relaxation is largely lost when performing the measurements at low temperatures. The 77 K spectra of leaves and thylakoid membranes are characterized by three main peaks, F₆₈₅, F₆₉₅, and F₇₃₀, believed to originate predominantly from Chl a in CP47 of PSII, a specific Chl in CP43 of PSII, and PSI, respectively (Satoh and Butler, 1978; van Dorssen et al., 1987; Andrizhiyevskaya et al., 2005; Komura et al., 2007). Fluorescence from the major LHCII peaks at 680 nm (Rijgersberg and Amesz, 1978) and from the PSII reaction center Chls at 683 nm (Roelofs et al., 1993; Andrizhiyevskaya et al., 2005). Low-temperature studies on the effects of HL irradiation are confined to the changes in the FR-to-red fluorescence ratio, which are the result of the quenching of PSII fluorescence or energy redistribution between the photosystems (state transitions). Ruban and Horton (1994) have shown that photochemical quenching in Guzmania is maximal at 688 nm, whereas nonphotochemical processes quench preferentially at 683 and 698 nm.In this study, we undertook a detailed investigation of the NPQ-associated spectral changes in the fluorescence spectra of Arabidopsis (Arabidopsis thaliana) measured at room temperature (RT) and at 77 K. It follows from the above discussion that deeper insight into the mechanisms of NPQ processes may be gained by combining the kinetic and the spectral information of the fluorescence changes occurring in NPQ. For this purpose, we developed a multiwavelength spectrometer with parallel detection, allowing us to follow the entire time-dependent fluorescence spectra of leaves during the induction and relaxation phases of NPQ with high sensitivity.Specific questions to be addressed in this study are the following. Are there more than one NPQ processes and NPQ sites? Are these processes occurring in a linked fashion or are they independent? How do they depend on the various cofactors known to affect NPQ, in particular regarding the roles of PsbS and Zx? Using this novel approach of adding the spectral information to the NPQ fluorescence changes, we discovered specific spectral changes associated with different NPQ components. By comparing the effects measured on various NPQ mutants of Arabidopsis, it is possible to assign these NPQ components to specific quenching processes. The results provide evidence that the total NPQ is a combination of several parallel and largely independent processes, likely occurring at different locations in the photosynthetic apparatus.     

Surveying Rubisco Diversity and Temperature Response to Improve Crop Photosynthetic Efficiency

The threat to global food security of stagnating yields and population growth makes increasing crop productivity a critical goal over the coming decades. One key target for improving crop productivity and yields is increasing the efficiency of photosynthesis. Central to photosynthesis is Rubisco, which is a critical but often rate-limiting component. Here, we present full Rubisco catalytic properties measured at three temperatures for 75 plants species representing both crops and undomesticated plants from diverse climates. Some newly characterized Rubiscos were naturally “better” compared to crop enzymes and have the potential to improve crop photosynthetic efficiency. The temperature response of the various catalytic parameters was largely consistent across the diverse range of species, though absolute values showed significant variation in Rubisco catalysis, even between closely related species. An analysis of residue differences among the species characterized identified a number of candidate amino acid substitutions that will aid in advancing engineering of improved Rubisco in crop systems. This study provides new insights on the range of Rubisco catalysis and temperature response present in nature, and provides new information to include in models from leaf to canopy and ecosystem scale.In a changing climate and under pressure from a population set to hit nine billion by 2050, global food security will require massive changes to the way food is produced, distributed, and consumed (Ort et al., 2015). To match rising demand, agricultural production must increase by 50 to 70% in the next 35 years, and yet the gains in crop yields initiated by the green revolution are slowing, and in some cases, stagnating (Long and Ort, 2010; Ray et al., 2012). Among a number of areas being pursued to increase crop productivity and food production, improving photosynthetic efficiency is a clear target, offering great promise (Parry et al., 2007; von Caemmerer et al., 2012; Price et al., 2013; Ort et al., 2015). As the gatekeeper of carbon entry into the biosphere and often acting as the rate-limiting step of photosynthesis, Rubisco, the most abundant enzyme on the planet (Ellis, 1979), is an obvious and important target for improving crop photosynthetic efficiency.Rubisco is considered to exhibit comparatively poor catalysis, in terms of catalytic rate, specificity, and CO₂ affinity (Tcherkez et al., 2006; Andersson, 2008), leading to the suggestion that even small increases in catalytic efficiency may result in substantial improvements to carbon assimilation across a growing season (Zhu et al., 2004; Parry et al., 2013; Galmés et al., 2014a; Carmo-Silva et al., 2015). If combined with complimentary changes such as optimizing other components of the Calvin Benson or photorespiratory cycles (Raines, 2011; Peterhansel et al., 2013; Simkin et al., 2015), optimized canopy architecture (Drewry et al., 2014), or introducing elements of a carbon concentrating mechanism (Furbank et al., 2009; Lin et al., 2014a; Hanson et al., 2016; Long et al., 2016), Rubisco improvement presents an opportunity to dramatically increase the photosynthetic efficiency of crop plants (McGrath and Long, 2014; Long et al., 2015; Betti et al., 2016). A combination of the available strategies is essential for devising tailored solutions to meet the varied requirements of different crops and the diverse conditions under which they are typically grown around the world.Efforts to engineer an improved Rubisco have not yet produced a “super Rubisco” (Parry et al., 2007; Ort et al., 2015). However, advances in engineering precise changes in model systems continue to provide important developments that are increasing our understanding of Rubisco catalysis (Spreitzer et al., 2005; Whitney et al., 2011a, 2011b; Morita et al., 2014; Wilson et al., 2016), regulation (Andralojc et al., 2012; Carmo-Silva and Salvucci, 2013; Bracher et al., 2015), and biogenesis (Saschenbrecker et al., 2007; Whitney and Sharwood, 2008; Lin et al., 2014b; Hauser et al., 2015; Whitney et al., 2015).A complementary approach is to understand and exploit Rubisco natural diversity. Previous characterization of Rubisco from a limited number of species has not only demonstrated significant differences in the underlying catalytic parameters, but also suggests that further undiscovered diversity exists in nature and that the properties of some of these enzymes could be beneficial if present in crop plants (Carmo-Silva et al., 2015). Recent studies clearly illustrate the variation possible among even closely related species (Galmés et al., 2005, 2014b, 2014c; Kubien et al., 2008; Andralojc et al., 2014; Prins et al., 2016).Until recently, there have been relatively few attempts to characterize the consistency, or lack thereof, of temperature effects on in vitro Rubisco catalysis (Sharwood and Whitney, 2014), and often studies only consider a subset of Rubisco catalytic properties. This type of characterization is particularly important for future engineering efforts, enabling specific temperature effects to be factored into any attempts to modify crops for a future climate. In addition, the ability to coanalyze catalytic properties and DNA or amino acid sequence provides the opportunity to correlate sequence and biochemistry to inform engineering studies (Christin et al., 2008; Kapralov et al., 2011; Rosnow et al., 2015). While the amount of gene sequence information available grows rapidly with improving technology, knowledge of the corresponding biochemical variation resulting has yet to be determined (Cousins et al., 2010; Carmo-Silva et al., 2015; Sharwood and Whitney, 2014; Nunes-Nesi et al., 2016).This study aimed to characterize the catalytic properties of Rubisco from diverse species, comprising a broad range of monocots and dicots from diverse environments. The temperature dependence of Rubisco catalysis was evaluated to tailor Rubisco engineering for crop improvement in specific environments. Catalytic diversity was analyzed alongside the sequence of the Rubisco large subunit gene, rbcL, to identify potential catalytic switches for improving photosynthesis and productivity. In vitro results were compared to the average temperature of the warmest quarter in the regions where each species grows to investigate the role of temperature in modulating Rubisco catalysis.     

Patterns of Metabolite Changes Identified from Large-Scale Gene Perturbations in Arabidopsis Using a Genome-Scale Metabolic Network

Metabolomics enables quantitative evaluation of metabolic changes caused by genetic or environmental perturbations. However, little is known about how perturbing a single gene changes the metabolic system as a whole and which network and functional properties are involved in this response. To answer this question, we investigated the metabolite profiles from 136 mutants with single gene perturbations of functionally diverse Arabidopsis (Arabidopsis thaliana) genes. Fewer than 10 metabolites were changed significantly relative to the wild type in most of the mutants, indicating that the metabolic network was robust to perturbations of single metabolic genes. These changed metabolites were closer to each other in a genome-scale metabolic network than expected by chance, supporting the notion that the genetic perturbations changed the network more locally than globally. Surprisingly, the changed metabolites were close to the perturbed reactions in only 30% of the mutants of the well-characterized genes. To determine the factors that contributed to the distance between the observed metabolic changes and the perturbation site in the network, we examined nine network and functional properties of the perturbed genes. Only the isozyme number affected the distance between the perturbed reactions and changed metabolites. This study revealed patterns of metabolic changes from large-scale gene perturbations and relationships between characteristics of the perturbed genes and metabolic changes.Rational and quantitative assessment of metabolic changes in response to genetic modification (GM) is an open question and in need of innovative solutions. Nontargeted metabolite profiling can detect thousands of compounds, but it is not easy to understand the significance of the changed metabolites in the biochemical and biological context of the organism. To better assess the changes in metabolites from nontargeted metabolomics studies, it is important to examine the changed metabolites in the context of the genome-scale metabolic network of the organism.Metabolomics is a technique that aims to quantify all the metabolites in a biological system (Nikolau and Wurtele, 2007; Nicholson and Lindon, 2008; Roessner and Bowne, 2009). It has been used widely in studies ranging from disease diagnosis (Holmes et al., 2008; DeBerardinis and Thompson, 2012) and drug discovery (Cascante et al., 2002; Kell, 2006) to metabolic reconstruction (Feist et al., 2009; Kim et al., 2012) and metabolic engineering (Keasling, 2010; Lee et al., 2011). Metabolomic studies have demonstrated the possibility of identifying gene functions from changes in the relative concentrations of metabolites (metabotypes or metabolic signatures; Ebbels et al., 2004) in various species including yeast (Saccharomyces cerevisiae; Raamsdonk et al., 2001; Allen et al., 2003), Arabidopsis (Arabidopsis thaliana; Brotman et al., 2011), tomato (Solanum lycopersicum; Schauer et al., 2006), and maize (Zea mays; Riedelsheimer et al., 2012). Metabolomics has also been used to better understand how plants interact with their environments (Field and Lake, 2011), including their responses to biotic and abiotic stresses (Dixon et al., 2006; Arbona et al., 2013), and to predict important agronomic traits (Riedelsheimer et al., 2012). Metabolite profiling has been performed on many plant species, including angiosperms such as Arabidopsis, poplar (Populus trichocarpa), and Catharanthus roseus (Sumner et al., 2003; Rischer et al., 2006), basal land plants such as Selaginella moellendorffii and Physcomitrella patens (Erxleben et al., 2012; Yobi et al., 2012), and Chlamydomonas reinhardtii (Fernie et al., 2012; Davis et al., 2013). With the availability of whole genome sequences of various species, metabolomics has the potential to become a useful tool for elucidating the functions of genes using large-scale systematic analyses (Fiehn et al., 2000; Saito and Matsuda, 2010; Hur et al., 2013).Although metabolomics data have the potential for identifying the roles of genes that are associated with metabolic phenotypes, the biochemical mechanisms that link functions of genes with metabolic phenotypes are still poorly characterized. For example, we do not yet know the principles behind how perturbing the expression of a single gene changes the metabolic system as a whole. Large-scale metabolomics data have provided useful resources for linking phenotypes to genotypes (Fiehn et al., 2000; Roessner et al., 2001; Tikunov et al., 2005; Schauer et al., 2006; Lu et al., 2011; Fukushima et al., 2014). For example, Lu et al. (2011) compared morphological and metabolic phenotypes from more than 5,000 Arabidopsis chloroplast mutants using gas chromatography (GC)- and liquid chromatography (LC)-mass spectrometry (MS). Fukushima et al. (2014) generated metabolite profiles from various characterized and uncharacterized mutant plants and clustered the mutants with similar metabolic phenotypes by conducting multidimensional scaling with quantified metabolic phenotypes. Nonetheless, representation and analysis of such a large amount of data remains a challenge for scientific discovery (Lu et al., 2011). In addition, these studies do not examine the topological and functional characteristics of metabolic changes in the context of a genome-scale metabolic network. To understand the relationship between genotype and metabolic phenotype, we need to investigate the metabolic changes caused by perturbing the expression of a gene in a genome-scale metabolic network perspective, because metabolic pathways are not independent biochemical factories but are components of a complex network (Berg et al., 2002; Merico et al., 2009).Much progress has been made in the last 2 decades to represent metabolism at a genome scale (Terzer et al., 2009). The advances in genome sequencing and emerging fields such as biocuration and bioinformatics enabled the representation of genome-scale metabolic network reconstructions for model organisms (Bassel et al., 2012). Genome-scale metabolic models have been built and applied broadly from microbes to plants. The first step toward modeling a genome-scale metabolism in a plant species started with developing a genome-scale metabolic pathway database for Arabidopsis (AraCyc; Mueller et al., 2003) from reference pathway databases (Kanehisa and Goto, 2000; Karp et al., 2002; Zhang et al., 2010). Genome-scale metabolic pathway databases have been built for several plant species (Mueller et al., 2005; Zhang et al., 2005, 2010; Urbanczyk-Wochniak and Sumner, 2007; May et al., 2009; Dharmawardhana et al., 2013; Monaco et al., 2013, 2014; Van Moerkercke et al., 2013; Chae et al., 2014; Jung et al., 2014). Efforts have been made to develop predictive genome-scale metabolic models using enzyme kinetics and stoichiometric flux-balance approaches (Sweetlove et al., 2008). de Oliveira Dal’Molin et al. (2010) developed a genome-scale metabolic model for Arabidopsis and successfully validated the model by predicting the classical photorespiratory cycle as well as known key differences between redox metabolism in photosynthetic and nonphotosynthetic plant cells. Other genome-scale models have been developed for Arabidopsis (Poolman et al., 2009; Radrich et al., 2010; Mintz-Oron et al., 2012), C. reinhardtii (Chang et al., 2011; Dal’Molin et al., 2011), maize (Dal’Molin et al., 2010; Saha et al., 2011), sorghum (Sorghum bicolor; Dal’Molin et al., 2010), and sugarcane (Saccharum officinarum; Dal’Molin et al., 2010). These predictive models have the potential to be applied broadly in fields such as metabolic engineering, drug target discovery, identification of gene function, study of evolutionary processes, risk assessment of genetically modified crops, and interpretations of mutant phenotypes (Feist and Palsson, 2008; Ricroch et al., 2011).Here, we interrogate the metabotypes caused by 136 single gene perturbations of Arabidopsis by analyzing the relative concentration changes of 1,348 chemically identified metabolites using a reconstructed genome-scale metabolic network. We examine the characteristics of the changed metabolites (the metabolites whose relative concentrations were significantly different in mutants relative to the wild type) in the metabolic network to uncover biological and topological consequences of the perturbed genes.     

Focus on Ethylene: Ethylene and the Regulation of Physiological and Morphological Responses to Nutrient Deficiencies

To cope with nutrient deficiencies, plants develop both morphological and physiological responses. The regulation of these responses is not totally understood, but some hormones and signaling substances have been implicated. It was suggested several years ago that ethylene participates in the regulation of responses to iron and phosphorous deficiency. More recently, its role has been extended to other deficiencies, such as potassium, sulfur, and others. The role of ethylene in so many deficiencies suggests that, to confer specificity to the different responses, it should act through different transduction pathways and/or in conjunction with other signals. In this update, the data supporting a role for ethylene in the regulation of responses to different nutrient deficiencies will be reviewed. In addition, the results suggesting the action of ethylene through different transduction pathways and its interaction with other hormones and signaling substances will be discussed.When plants suffer from a mineral nutrient deficiency, they develop morphological and physiological responses (mainly in their roots) aimed to facilitate the uptake and mobilization of the limiting nutrient. After the nutrient has been acquired in enough quantity, these responses need to be switched off to avoid toxicity and conserve energy. In recent years, different plant hormones (e.g. ethylene, auxin, cytokinins, jasmonic acid, abscisic acid, brassinosteroids, GAs, and strigolactones) have been implicated in the regulation of these responses (Romera et al., 2007, 2011, 2015; Liu et al., 2009; Rubio et al., 2009; Kapulnik et al., 2011; Kiba et al., 2011; Iqbal et al., 2013; Zhang et al., 2014).Before the 1990s, there were several publications relating ethylene and nutrient deficiencies (cited in Lynch and Brown [1997] and Romera et al. [1999]) without establishing a direct implication of ethylene in the regulation of nutrient deficiency responses. In 1994, Romera and Alcántara (1994) published an article in Plant Physiology suggesting a role for ethylene in the regulation of Fe deficiency responses. In 1999, Borch et al. (1999) showed the participation of ethylene in the regulation of P deficiency responses. Since then, evidence has been accumulating in support of a role for ethylene in the regulation of both Fe (Romera et al., 1999, 2015; Waters and Blevins, 2000; Lucena et al., 2006; Waters et al., 2007; García et al., 2010, 2011, 2013, 2014; Yang et al., 2014) and P deficiency responses (Kim et al., 2008; Lei et al., 2011; Li et al., 2011; Nagarajan and Smith, 2012; Wang et al., 2012, 2014c). Both Fe and P may be poorly available in most soils, and plants develop similar responses under their deficiencies (Romera and Alcántara, 2004; Zhang et al., 2014). More recently, a role for ethylene has been extended to other deficiencies, such as K (Shin and Schachtman, 2004; Jung et al., 2009; Kim et al., 2012), S (Maruyama-Nakashita et al., 2006; Wawrzyńska et al., 2010; Moniuszko et al., 2013), and B (Martín-Rejano et al., 2011). Ethylene has also been implicated in both N deficiency and excess (Tian et al., 2009; Mohd-Radzman et al., 2013; Zheng et al., 2013), and its participation in Mg deficiency has been suggested (Hermans et al., 2010).In this update, we will review the information supporting a role for ethylene in the regulation of different nutrient deficiency responses. For information relating ethylene to other aspects of plant mineral nutrition, such as N₂ fixation and responses to excess of nitrate or essential heavy metals, the reader is referred to other reviews (for review, see Maksymiec, 2007; Mohd-Radzman et al., 2013; Steffens, 2014).