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 Zeaxanthin-Independent and Zeaxanthin-Dependent qE Components of Nonphotochemical Quenching Involve Common Conformational Changes within the Photosystem II Antenna in Arabidopsis

The light-harvesting antenna of higher plant photosystem II (LHCII) has the intrinsic capacity to dissipate excess light energy as heat in a process termed nonphotochemical quenching (NPQ). Recent studies suggest that zeaxanthin and lutein both contribute to the rapidly relaxing component of NPQ, qE, possibly acting in the minor monomeric antenna complexes and the major trimeric LHCII, respectively. To distinguish whether zeaxanthin and lutein act independently as quenchers at separate sites, or alternatively whether zeaxanthin fulfills an allosteric role regulating lutein-mediated quenching, the kinetics of qE and the qE-related conformational changes (ΔA₅₃₅) were compared in Arabidopsis (Arabidopsis thaliana) mutant/antisense plants with altered contents of minor antenna (kolhcb6, aslhcb4), trimeric LHCII (aslhcb2), lutein (lut2, lut2npq1, lut2npq2), and zeaxanthin (npq1, npq2). The kinetics of the two components of NPQ induction arising from zeaxanthin-independent and zeaxanthin-dependent qE were both sensitive to changes in the protein composition of the photosystem II antenna. The replacement of lutein by zeaxanthin or violaxanthin in the internal Lhcb protein-binding sites affected the kinetics and relative amplitude of each component as well as the absolute chlorophyll fluorescence lifetime. Both components of qE were characterized by a conformational change leading to nearly identical absorption changes in the Soret region that indicated the involvement of the LHCII lutein 1 domain. Based on these observations, we suggest that both components of qE arise from a common quenching mechanism based upon a conformational change within the photosystem II antenna, optimized by Lhcb subunit-subunit interactions and tuned by the synergistic effects of external and internally bound xanthophylls.The chlorophyll a/b-binding light-harvesting antenna of photosystem II (PSII of higher plants is responsible for the efficient collection and transfer of excitation energy to the reaction center. The PSII antenna comprises the main trimeric light-harvesting complex, LHCII, which is composed of the Lhcb1 to -3 polypeptides, and the minor light-harvesting complexes, CP29, CP26, and CP24, composed of Lhcb4, -5, and -6, respectively. In Arabidopsis (Arabidopsis thaliana), four LHCII trimers associate with two copies each of CP24, CP26, and CP29 and a core dimer of PSII (CP43/D1/D2/CP47) to form the C₂S₂M₂ LHCII-PSII supercomplex (Dekker and Boekema, 2005). In addition, depending upon the growth conditions, two or three extra LHCII trimers per PSII may be present in LHCII-only regions of the grana, providing additional light-harvesting capacity.The PSII antenna is a highly dynamic system that is able to tune the amount of excitation delivered to the PSII reaction center to match physiological need (Horton et al., 1996). The regulation of energy flow occurs by control of the thermal dissipation of excess excitation within the PSII antenna, a process termed nonphotochemical quenching (NPQ). NPQ is heterogeneous, comprising a slowly reversible qI component and a rapidly reversible qE component (Horton et al., 1996). The trigger for qE is the buildup of the transmembrane proton gradient or ΔpH (Briantais et al., 1979). The ΔpH is sensed by the PsbS protein (Li et al., 2004), without which the rapidly reversible behavior of NPQ is lost (Li et al., 2000). Full expression of qE in vivo is associated with the enzymatic deepoxidation of the epoxy-xanthophyll violaxanthin to zeaxanthin, via the action of the xanthophyll cycle (Demmig-Adams, 1990). The majority of the photoconvertible xanthophyll cycle pool is associated with trimeric LHCII, bound at the external V1 binding site (Ruban et al., 1999, 2002a; Caffarri et al., 2001; Liu et al., 2004). Trimeric LHCII binds two other types of xanthophylls internally: two all-trans-luteins at the L1 and L2 sites associated with the central membrane-spanning α-helices; and a 9-cis-neoxanthin at the N1 site associated with the C-helix chlorophyll b domain (Liu et al., 2004). The minor monomeric complexes CP24, CP26, and CP29 all bind lutein at the L1 site. In addition, CP29 binds two xanthophyll cycle carotenoids and one-half to one neoxanthin, CP24 binds two xanthophyll cycle carotenoids, while CP26 binds one xanthophyll cycle carotenoid and one neoxanthin (Peter and Thornber, 1991; Bassi et al., 1993; Ruban et al., 1994, 1999; Morosinotto et al., 2002).Although there is strong evidence that qE occurs in the PSII antenna light-harvesting proteins and that xanthophylls are involved, the mechanism of energy dissipation remains unclear. There is evidence for two distinct quenching mechanisms, one involving zeaxanthin (type I) and the other lutein (type II). In the type I mechanism, it is proposed that qE obligatorily depends upon zeaxanthin acting as a quencher of excited chlorophyll via the formation of a charge transfer state. Evidence for type I is the formation of a carotenoid radical cation absorbing at approximately 1,000 nm that correlates with the extent of qE (Holt et al., 2005). Recently, evidence was obtained that formation of the zeaxanthin radical cation occurs exclusively at the L2 binding site of the minor antenna complexes (Ahn et al., 2008; Avenson et al., 2008), quenching therefore requiring reversible insertion of zeaxanthin into this internal site. Because the effect of this cation on the excited-state lifetime of the minor antenna complexes was found to be very small, it was suggested that in vivo, under the influence of the ΔpH, a large population of complexes would adopt a conformation in which this species could form (Avenson et al., 2008). Evidence was also obtained that a zeaxanthin radical cation may form in trimeric LHCII (Amarie et al., 2007). Again, the effect on the chlorophyll excited-state lifetime was very small, leading these authors to conclude that the type I mechanism could not be responsible for qE (Amarie et al., 2007; Dreuw and Wormit, 2008).In the type II mechanism, qE is an inbuilt property of LHCII proteins; a protein conformational change alters the configuration of bound pigments and results in the xanthophyll bound at the L1 site (normally lutein) becoming an effective quencher of chlorophyll excited states (Ruban et al., 2007; Ilioaia et al., 2008). Evidence for a type II mechanism came from studies of trimeric LHCII aggregates (Ruban et al., 2007). Here, it was concluded that energy dissipation occurs by energy transfer from chlorophyll a to the S₁ state (2Ag¹) of lutein bound at the L1 site. Notably, this quenching mechanism decreases the chlorophyll excited-state lifetime by a magnitude sufficient to fully account for qE in vivo. A change in the conformation of another LHCII-bound xanthophyll (neoxanthin) correlates with the extent of quenching. This conformational change takes place in vivo with an amplitude that correlates with the amount of qE. In the model for type II quenching proposed by Horton and coworkers (1991, 2005), zeaxanthin acts not as a quencher but as an allosteric modulator of the ΔpH sensitivity of this intrinsic LHCII quenching process.Although the type I and type II mechanisms involve different xanthophylls operating at different sites, there are similarities: in particular, both are proposed to involve a ΔpH-triggered, PsbS-mediated conformational change (Ruban et al., 2007; Ahn et al., 2008). Indeed, it is possible that both mechanisms contribute to in vivo qE, since the process occurs in both the presence and absence of zeaxanthin (Adams et al., 1990; Crouchman et al., 2006). The crucial question is whether zeaxanthin-dependent and zeaxanthin-independent qE arise from the same mechanism (type II) or from two different ones (types I and II, respectively). The kinetics of NPQ formation upon the illumination of dark-adapted leaves comprise two components: the first forms rapidly and is zeaxanthin independent; the second, slower component correlates with violaxanthin deepoxidation and therefore is described as zeaxanthin dependent (Adams et al., 1990; Ruban and Horton, 1999). The two components of NPQ formation are of the qE type: both relax rapidly upon darkening (Adams et al., 1990); both are dependent upon PsbS (Li et al., 2000); and both are enhanced by PsbS overexpression (Li et al., 2002; Crouchman et al., 2006). Investigation of these kinetics provides an opportunity to determine whether a single mechanism can account for qE and to give clues to which type of mechanism is involved. Here, we test the hypothesis that the two components arise from different mechanisms: the zeaxanthin-dependent component arising in the minor monomeric antenna by a type I mechanism (Gilmore et al., 1998; Ahn et al., 2008; Avenson et al., 2008), and the zeaxanthin-independent component arising in the major trimeric LHCII by the type II mechanism. An alternative explanation for zeaxanthin-independent qE, at least under low-light conditions, when qE forms transiently, is that it is caused by quenching in the PSII reaction center (Finazzi et al., 2004). Several predictions emerge from this hypothesis. First, the removal of certain Lhcb proteins by mutation would differentially affect the two components of qE. Second, because the two components would be additive and could not compensate for the loss of one another (Niyogi et al., 1998; Pogson et al., 1998), they should each contribute a discrete component to the kinetics of qE formation and relaxation. Third, in mutants lacking lutein, the capacity of the type II mechanism would be reduced, while the zeaxanthin-dependent component would be unaffected. Finally, the two components may be expected to be characterized by different absorption changes in the Soret region, which reflect changes in the absorption spectra of bound pigments brought about by conformational changes within the PSII antenna upon qE formation (Ruban et al., 1993a, 1993b, 2002b; Bilger and Björkman, 1994). We tested this hypothesis by analysis of qE kinetics, fluorescence lifetimes, and qE-related absorption difference spectra. Contrary to the above predictions, the data indicated that both steady-state and transient qE arise from a common mechanism within the PSII antenna, in both the presence and absence of zeaxanthin.     

PHOTOSYSTEM II SUBUNIT R Is Required for Efficient Binding of LIGHT-HARVESTING COMPLEX STRESS-RELATED PROTEIN3 to Photosystem II-Light-Harvesting Supercomplexes in Chlamydomonas reinhardtii

In Chlamydomonas reinhardtii, the LIGHT-HARVESTING COMPLEX STRESS-RELATED PROTEIN3 (LHCSR3) protein is crucial for efficient energy-dependent thermal dissipation of excess absorbed light energy and functionally associates with photosystem II-light-harvesting complex II (PSII-LHCII) supercomplexes. Currently, it is unknown how LHCSR3 binds to the PSII-LHCII supercomplex. In this study, we investigated the role of PHOTOSYSTEM II SUBUNIT R (PSBR) an intrinsic membrane-spanning PSII subunit, in the binding of LHCSR3 to PSII-LHCII supercomplexes. Down-regulation of PSBR expression diminished the efficiency of oxygen evolution and the extent of nonphotochemical quenching and had an impact on the stability of the oxygen-evolving complex as well as on PSII-LHCII-LHCSR3 supercomplex formation. Its down-regulation destabilized the PSII-LHCII supercomplex and strongly reduced the binding of LHCSR3 to PSII-LHCII supercomplexes, as revealed by quantitative proteomics. PHOTOSYSTEM II SUBUNIT P deletion, on the contrary, destabilized PHOTOSYSTEM II SUBUNIT Q binding but did not affect PSBR and LHCSR3 association with PSII-LHCII. In summary, these data provide clear evidence that PSBR is required for the stable binding of LHCSR3 to PSII-LHCII supercomplexes and is essential for efficient energy-dependent quenching and the integrity of the PSII-LHCII-LHCSR3 supercomplex under continuous high light.Plant photosynthetic electron transfer is conducted by a series of reactions at the chloroplast thylakoid membrane, resulting in light-dependent water oxidation, NADP reduction, and ATP formation (Whatley et al., 1963). Two separate photosystems (PSI and PSII) and an ATP synthase catalyze these reactions. PSI and PSII are multiprotein complexes that are mainly embedded in unstacked and stacked regions of the thylakoid membrane, respectively. PSI consists of more than 10 subunits and a number of cofactors such as chlorophyll a, β-carotene, phylloquinone, and three iron-sulfur (4Fe-4S) clusters (Busch and Hippler, 2011). PSI catalyzes light-driven electron transfer from luminal plastocyanin to stromal ferredoxin. The latter reduces the ferredoxin-NADP reductase that, in turn, leads to the formation of NADPH. PSII catalyzes light-induced electron transfer from water to the plastoquinone pool by using chlorophyll a, carotenoids, as well as redox-active cofactors, causing the release of oxygen and protons (Pagliano et al., 2013). The core complex is organized as a dimer. Monomers are composed of the reaction center subunits PSBA (D1) and PSBD (D2), the inner antenna proteins PSBB (CP47) and PSBC (CP43), the α- and β-subunits (PSBE and PSBF) of cytochrome b₅₅₉, as well as a number of intrinsic low-molecular-mass subunits. The core monomer is further associated with an inorganic Mn₄O₅Ca cluster and a few chloride ions (Rivalta et al., 2011; Umena et al., 2011) required for photosynthetic water oxidation. To optimize this process, the oxygen-evolving complex is formed at the luminal side by the extrinsic polypeptides PSBO, PSBP, PSBQ, and PSBR (for review, see Pagliano et al., 2013).To enhance the light-harvesting capacity of PSII, various light-harvesting proteins bind to dimeric PSII core complexes (Dekker and Boekema, 2005). A common structure found for vascular plants and green algae is the C₂S₂ supercomplex, where two copies of monomeric Lhcb4 and Lhcb5 and two LHCII trimers (S-trimer; Boekema et al., 1995) bind to the dimeric PSII core. In vascular plants, larger but less stable PSII supercomplexes, known as C₂S₂M₂, are composed of two extra copies of the monomeric Lhcb6 with two additional LHCII trimers (M-trimer) bound through Lhcb4 and Lhcb5 (Dekker and Boekema, 2005; Caffarri et al., 2009). Even larger complexes containing two additional LHCII trimers (L-trimer), bound via Lhcb6, are found and are known as C₂S₂M₂L_1–2 (Boekema et al., 1999). A recent study in Chlamydomonas reinhardtii identified PSII-LHCII supercomplexes with three LHCII trimers attached to each side of the core (C₂S₂M₂L₂; Tokutsu et al., 2012). Interestingly, such PSII-LHCII supercomplexes associate with LIGHT-HARVESTING COMPLEX STRESS-RELATED PROTEIN3 (LHCSR3; Tokutsu and Minagawa, 2013), an ancient light-harvesting protein required for efficient energy-dependent (qE) quenching in the alga (Peers et al., 2009). The qE component of nonphotochemical quenching (NPQ) is an energy-dependent constituent of NPQ and regulates the thermal dissipation of excess absorbed light energy (Li et al., 2000; Peers et al., 2009). The qE capacity in C. reinhardtii increases proportionally to the light-dependent accumulation of the LHCSR3 protein (Peers et al., 2009). In contrast, in vascular plants, qE is constitutively active and dependent on PSBS, a PSII polypeptide (Li et al., 2000). Mass spectrometric analyses of isolated C₂S₂M₂ PSII supercomplexes revealed the presence of extrinsic subunits PSBP, PSBQ, and PSBR, while PSBS was not identified, suggesting that PSBS does not influence the association of the PSII core with the outer light-harvesting complex system (Pagliano et al., 2014). In line with the proteomic findings, recent data suggest that subunits PSBP, PSBQ, and PSBR contribute to the stability of PSII-LHCII supercomplexes in vascular plants (Caffarri et al., 2009; Ifuku et al., 2011; Allahverdiyeva et al., 2013). A recent quantitative proteomic study performed with C. reinhardtii identified PSBR as the only PSII subunit to be induced upon the shift from photoheterotrophic to photoautotrophic growth conditions similar to LHCSR3 (Höhner et al., 2013).In vascular plants and green algae, PSBR is nucleus encoded and has a mass of about 10 kD. The mature protein has a predicted 70-amino acid luminal N-terminal part and a C-terminal transmembrane span (Ljungberg et al., 1986; Lautner et al., 1988; Webber et al., 1989). An association of PSBR with the oxygen-evolving complex has been suggested, as its presence is required for the stable assembly of PSBP with the PSII core and its absence also impacts the binding of PSBQ to the core (Suorsa et al., 2006; Liu et al., 2009). For stable association with the PSII core complex, PSBR needs the presence of PSBJ (Suorsa et al., 2006). Functionally, the depletion of PSBR protein expression decreased rates of oxygen evolution (Allahverdiyeva et al., 2007, 2013) and quinone reoxidation (Allahverdiyeva et al., 2007). PSBR phosphorylation is known for Arabidopsis (Arabidopsis thaliana; Reiland et al., 2009, 2011; Nakagami et al., 2010) and in the green alga C. reinhardtii (Turkina et al., 2006), although phosphorylation sites are not conserved between the alga and the vascular plant.In this work, we addressed the question of whether down-regulation of PSBR expression would affect LHCSR3 binding to the PSII-LHCII supercomplex in C. reinhardtii. To this end, we took advantage of artificial microRNA (amiRNA) technology to down-regulate PSBR expression and investigated the impact of PSBR down-regulation on photosynthetic performance as well as on PSII-LHCII-LHCSR3 supercomplex formation. Our data provide evidence that PSBR is required for the stable binding of LHCSR3 to PSII-LHCII supercomplexes.     

Comparative Analysis of Light-Harvesting Antennae and State Transition in chlorina and cpSRP Mutants

State transitions in photosynthesis provide for the dynamic allocation of a mobile fraction of light-harvesting complex II (LHCII) to photosystem II (PSII) in state I and to photosystem I (PSI) in state II. In the state I-to-state II transition, LHCII is phosphorylated by STN7 and associates with PSI to favor absorption cross-section of PSI. Here, we used Arabidopsis (Arabidopsis thaliana) mutants with defects in chlorophyll (Chl) b biosynthesis or in the chloroplast signal recognition particle (cpSRP) machinery to study the flexible formation of PS-LHC supercomplexes. Intriguingly, we found that impaired Chl b biosynthesis in chlorina1-2 (ch1-2) led to preferentially stabilized LHCI rather than LHCII, while the contents of both LHCI and LHCII were equally depressed in the cpSRP43-deficient mutant (chaos). In view of recent findings on the modified state transitions in LHCI-deficient mutants (Benson et al., 2015), the ch1-2 and chaos mutants were used to assess the influence of varying LHCI/LHCII antenna size on state transitions. Under state II conditions, LHCII-PSI supercomplexes were not formed in both ch1-2 and chaos plants. LHCII phosphorylation was drastically reduced in ch1-2, and the inactivation of STN7 correlates with the lack of state transitions. In contrast, phosphorylated LHCII in chaos was observed to be exclusively associated with PSII complexes, indicating a lack of mobile LHCII in chaos. Thus, the comparative analysis of ch1-2 and chaos mutants provides new evidence for the flexible organization of LHCs and enhances our understanding of the reversible allocation of LHCII to the two photosystems.In oxygenic photosynthesis, PSII and PSI function in series to convert light energy into the chemical energy that fuels multiple metabolic processes. Most of this light energy is captured by the chlorophyll (Chl) and carotenoid pigments in the light-harvesting antenna complexes (LHCs) that are peripherally associated with the core complexes of both photosystems (Wobbe et al., 2016). However, since the two photosystems exhibit different absorption spectra (Nelson and Yocum, 2006; Nield and Barber, 2006; Qin et al., 2015), PSI or PSII is preferentially excited under naturally fluctuating light intensities and qualities. To optimize photosynthetic electron transfer, the excitation state of the two photosystems must be rebalanced in response to changes in lighting conditions. To achieve this, higher plants and green algae require rapid and precise acclimatory mechanisms to adjust the relative absorption cross-sections of the two photosystems.To date, the phenomenon of state transitions is one of the well-documented short-term acclimatory mechanisms. It allows a mobile portion of the light-harvesting antenna complex II (LHCII) to be allocated to either photosystem, depending on the spectral composition and intensity of the ambient light (Allen and Forsberg, 2001; Rochaix, 2011; Goldschmidt-Clermont and Bassi, 2015; Gollan et al., 2015). State transitions are driven by the redox state of the plastoquinone (PQ) pool (Vener et al., 1997; Zito et al., 1999). When PSI is preferentially excited (by far-red light), the PQ pool is oxidized and all the LHCII is associated with PSII. This allocation of antenna complexes is defined as state I. When light conditions (blue/red light or low light) favor exciton trapping of PSII, the transition from state I to state II occurs. The over-reduced PQ pool triggers the activation of the membrane-localized Ser-Thr kinase STN7, which phosphorylates an N-terminal Thr on each of two major LHCII proteins, LHCB1 and LHCB2 (Allen, 1992; Bellafiore et al., 2005; Shapiguzov et al., 2016). Phosphorylation of LHCII results in the dissociation of LHCII from PSII and triggers its reversible relocation to PSI (Allen, 1992; Rochaix, 2011). Conversely, when the PQ pool is reoxidized, STN7 is inactivated and the constitutively active, thylakoid-associated phosphatase TAP38/PPH1 dephosphorylates LHCII, which then reassociates with PSII (Pribil et al., 2010; Shapiguzov et al., 2010). The physiological significance of state transitions has been demonstrated by the reduction in growth rate seen in the stn7 knock-out mutant under fluctuating light conditions (Bellafiore et al., 2005; Tikkanen et al., 2010).The canonical state transitions model implies spatial and temporal regulation of the allocation of LHC between the two spatially segregated photosystems (Dekker and Boekema, 2005). PSII-LHCII supercomplexes are organized in a tightly packed form in the stacked grana regions of thylakoid membranes, while PSI-LHCI supercomplexes are mainly localized in the nonstacked stromal lamellae and grana margin regions (Dekker and Boekema, 2005; Haferkamp et al., 2010). It has been proposed that, in the grana margin regions, which harbor LHCII and both photosystems, LHCII can migrate rapidly between them (Albertsson et al., 1990; Albertsson, 2001). This idea is supported by the recent discovery of mega complexes containing both photosystems in the grana margin regions (Yokono et al., 2015). Furthermore, phosphorylation of LHCII was found to increase not only the amount of PSI found in the grana margin region of thylakoid membranes (Tikkanen et al., 2008a), but also to modulate the pattern of PSI-PSII megacomplexes under changing light conditions (Suorsa et al., 2015). Nonetheless, open questions remain in relation to the physiological significance of the detection of phosphorylated LHCII in all thylakoid regions, even under the constant light conditions (Grieco et al., 2012; Leoni et al., 2013; Wientjes et al., 2013), although LHCII phosphorylation has been shown to modify the stacking of thylakoid membranes (Chuartzman et al., 2008; Pietrzykowska et al., 2014).State I-to-state II transition is featured by the formation of LHCII-PSI-LHCI supercomplexes, in which LHCII favors the light-harvesting capacity of PSI. Recently, LHCII-PSI-LHCI supercomplexes have been successfully isolated and purified using various detergents (Galka et al., 2012; Drop et al., 2014; Crepin and Caffarri, 2015) or a styrene-maleic acid copolymer (Bell et al., 2015). These findings yielded further insights into the reorganization of supercomplexes associated with state transitions, and it was suggested that phosphorylation of LHCB2 rather than LHCB1 is the essential trigger for the formation of state transition supercomplexes (Leoni et al., 2013; Pietrzykowska et al., 2014; Crepin and Caffarri, 2015; Longoni et al., 2015). Furthermore, characterization of mutants deficient in individual PSI core subunits indicates that PsaH, L, and I are required for docking of LHCII at PSI (Lunde et al., 2000; Zhang and Scheller, 2004; Kouril et al., 2005; Plöchinger et al., 2016).Recently, the state transition capacity has been characterized in the Arabidopsis (Arabidopsis thaliana) mutants with missing LHCI components. Although the Arabidopsis knock-out mutants lacking one of the four LHCI proteins (LHCA1-4) showed enhanced accumulation of LHCII-PSI complexes, the absorption cross-section of PSI under state II conditions was still compromised in the lhca1-4 mutants, and it is suggested that LHCI mediates the detergent-sensitive interaction between ‘extra LHCII’ and PSI (Benson et al., 2015; Grieco et al., 2015). Furthermore, the Arabidopsis mutant ΔLhca lacking all LHCA1-4 proteins was shown to be compensated for the deficiency of LHCI by binding LHCII under state II conditions (Bressan et al., 2016). In spite of this finding, the significant reduction in the absorption cross-section of PSI was still observed in the ΔLhca mutant, suggesting a substantial role of LHCI in light absorption under canopy conditions (Bressan et al., 2016). However, these findings emphasize the acclimatory function of state transitions in balancing light absorption capacity between the two photosystems by modifying their relative antenna size and imply the dynamic and variable organization of PS-LHC supercomplexes.LHC proteins are encoded by the nuclear Lhc superfamily (Jansson, 1994). The biogenesis of LHCs includes the cytoplasmic synthesis of the LHC precursor proteins, their translocation into chloroplasts via the TOC/TIC complex, and their posttranslational targeting and integration into the thylakoid membranes by means of the chloroplast signal recognition particle (cpSRP) machinery (Jarvis and Lopez-Juez, 2013). The posttranslational cpSRP-dependent pathway for the final translocation of LHC proteins into the thylakoid membrane includes interaction of cpSRP43 with LHC apo-proteins and recruitment of cpSRP54 to form a transit complex. Then binding of this tripartite cpSRP transit complex to the SRP receptor cpFtsY follows, which supports docking of the transit complex to thylakoid membranes and its association with the LHC translocase ALB3. Ultimately, ALB3 inserts LHC apo-proteins into the thylakoid membrane (Richter et al., 2010). Importantly, stoichiometric amounts of newly synthesized Chl a and Chl b as well as carotenoid are inserted into the LHC apo-proteins by unknown mechanisms to form the functional LHCs that associate with the core complexes of both photosystems in the thylakoid membranes (Dall’Osto et al., 2015; Wang and Grimm, 2015).The first committed steps in Chl synthesis occur in the Mg branch of the tetrapyrrole biosynthesis pathway. 5-Aminolevulinic acid synthesis provides the precursor for the formation of protoporphyrin IX, which is directed into the Mg branch (Tanaka and Tanaka, 2007; Brzezowski et al., 2015). Chl synthesis ends with the conversion of Chl a to Chl b catalyzed by Chl a oxygenase (CAO; Tanaka et al., 1998; Tomitani et al., 1999). It has been hypothesized that coordination between Chl synthesis and the posttranslational cpSRP pathway is a prerequisite for the efficient integration of Chls into LHC apo-proteins.In this study, we intend to characterize the assembly of LHCs when the availability of Chl molecules or the integration of LHC apo-proteins into thylakoid membranes is limiting. To this end, we compared the assembly of LHCs and the organization of PS-LHC complexes in two different sets of Arabidopsis mutants. Firstly, we used the chlorina1-2 (ch1-2) mutant, which is defective in the CAO gene. The members of the second set of mutants carry knock-out mutations in genes involved in the chloroplast SRP pathway (Richter et al., 2010).Our studies revealed distinct accumulation of PS-LHC supercomplexes between the two sets of mutant relative to wild-type plants. In spite of the defect in synthesis of Chl b, ch1-2 retains predominantly intact PSI-LHCI supercomplexes but has strongly reduced amounts of LHCII. In contrast, the chaos (cpSRP43) mutant exhibits synchronously reduced contents of both LHCI and LHCII, which results in the accumulation of PS core complexes without accompanying LHCs. Thus, the distribution of LHCs in the thylakoid membranes of the two mutants, ch1-2 and chaos, were explored under varying light conditions with the aim of elucidating the influence of modified LHCI/LHCII antenna size on state transitions. Our results contribute to an expanding view on the variety of photosynthetic complexes, which can be observed in Arabidopsis plants with specified mutations in LHC biogenesis.     

STATE TRANSITION7-Dependent Phosphorylation Is Modulated by Changing Environmental Conditions,and Its Absence Triggers Remodeling of Photosynthetic Protein Complexes

In plants and algae, the serine/threonine kinase STN7/STT7, orthologous protein kinases in Chlamydomonas reinhardtii and Arabidopsis (Arabidopsis thaliana), respectively, is an important regulator in acclimation to changing light environments. In this work, we assessed STT7-dependent protein phosphorylation under high light in C. reinhardtii, known to fully induce the expression of LIGHT-HARVESTING COMPLEX STRESS-RELATED PROTEIN3 (LHCSR3) and a nonphotochemical quenching mechanism, in relationship to anoxia where the activity of cyclic electron flow is stimulated. Our quantitative proteomics data revealed numerous unique STT7 protein substrates and STT7-dependent protein phosphorylation variations that were reliant on the environmental condition. These results indicate that STT7-dependent phosphorylation is modulated by the environment and point to an intricate chloroplast phosphorylation network responding in a highly sensitive and dynamic manner to environmental cues and alterations in kinase function. Functionally, the absence of the STT7 kinase triggered changes in protein expression and photoinhibition of photosystem I (PSI) and resulted in the remodeling of photosynthetic complexes. This remodeling initiated a pronounced association of LHCSR3 with PSI-LIGHT HARVESTING COMPLEX I (LHCI)-ferredoxin-NADPH oxidoreductase supercomplexes. Lack of STT7 kinase strongly diminished PSII-LHCII supercomplexes, while PSII core complex phosphorylation and accumulation were significantly enhanced. In conclusion, our study provides strong evidence that the regulation of protein phosphorylation is critical for driving successful acclimation to high light and anoxic growth environments and gives new insights into acclimation strategies to these environmental conditions.Oxygenic photosynthesis converts solar energy into chemical energy. This energy is utilized for carbon dioxide assimilation, allowing the formation of complex organic material. Plant photosynthesis is performed by a series of reactions in and at the thylakoid membrane, resulting in light-dependent water oxidation, NADP reduction, and ATP formation (Whatley et al., 1963). These light reactions are catalyzed by two photosystems (PSI and PSII). A third multiprotein complex, also embedded in the thylakoid membrane, is the cytochrome b₆f (cyt b₆f) complex that links photosynthetic electron transfer processes between the two photosystems and functions in proton translocation. The ATP synthase takes advantage of the proton-motive force that is generated by the light reactions (Mitchell, 1961) to produce ATP. ATP and NADPH, generated through linear electron flow from PSII to PSI, drive the Calvin-Benson-Bassham cycle (Bassham et al., 1950) to fix CO₂. Alternatively, cyclic electron flow (CEF) between PSI and the cyt b₆f complex solely produces ATP (Arnon, 1959).Under normal growth conditions, CEF provides additionally required ATP for CO₂ fixation (Lucker and Kramer, 2013), counteracts overreduction of the PSI acceptor side under stressful environmental cues, and readjusts the ATP poise, leading to increased lumen acidification important for photoprotection (Alric, 2010; Peltier et al., 2010; Leister and Shikanai, 2013; Shikanai, 2014). In microalgae and vascular plants, CEF relies on the NAD(P)H dehydrogenase-dependent and/or PROTON GRADIENT REGULATION5 (PGR5)-related pathways (Munekage et al., 2002, 2004; Petroutsos et al., 2009; Tolleter et al., 2011; Johnson et al., 2014). For both pathways, supercomplexes consisting of PSI-LIGHT HARVESTING COMPLEX I (LHCI) and components of the respective electron transfer routes have been identified. In Arabidopsis (Arabidopsis thaliana), a unique NAD(P)H dehydrogenase-PSI supercomplex with a molecular mass of more than 1,000 kD was discovered (Peng et al., 2008). From Chlamydomonas reinhardtii, Iwai et al. (2010) isolated a protein supercomplex composed of PSI-LHCI, LHCII, the cyt b₆/f complex, ferredoxin-NADPH oxidoreductase (FNR), and PROTON GRADIENT REGULATION-LIKE1 (PGRL1).PGRL1 and PGR5 interact physically in Arabidopsis and associate with PSI to allow the operation of CEF (DalCorso et al., 2008). Functional data suggest that PGRL1 might operate as a ferredoxin-plastoquinone reductase (Hertle et al., 2013). The PGRL1-containing CEF supercomplex isolated from C. reinhardtii is capable of CEF under in vitro conditions in the presence of exogenously added soluble plastocyanin and ferredoxin (Iwai et al., 2010). Terashima et al. (2012) isolated a CEF supercomplex of similar composition from anaerobic growth conditions that was active in vitro and contained proteins such as the chloroplast-localized Ca²⁺ sensor CAS and ANAEROBIC RESPONSE1 (ANR1), which were also shown to be functionally important for efficient CEF in the alga. Notably, it was suggested that the onset of CEF in C. reinhardtii is redox controlled (Takahashi et al., 2013).It has been demonstrated that efficient CEF is crucial for successful acclimation to excess light (Munekage et al., 2004; Dang et al., 2014; Johnson et al., 2014; Kukuczka et al., 2014). The most rapid response to excess light, however, relies on a mechanism called nonphotochemical quenching (NPQ). The fastest constituent of NPQ is energy-dependent (qE) quenching, which operates at a time scale of seconds to minutes and regulates the thermal dissipation of excess absorbed light energy, thereby providing effective photoprotection. In vascular plants, the PSII protein PSII SUBUNIT S is essential for qE (Li et al., 2000), whereas qE induction in the green alga C. reinhardtii is mediated by LIGHT-HARVESTING COMPLEX STRESS-RELATED PROTEIN3 (LHCSR3), an ancient light-harvesting protein that is missing in vascular plants (Peers et al., 2009). CEF and qE are complementary for acclimation to excess light, as double mutants deficient in both mechanisms possess additive phenotypes and are highly sensitive to light (Kukuczka et al., 2014). Another constituent of NPQ is the quenching by state transitions. State transitions are important to balance the excitation energy between PSI and PSII (Bonaventura and Myers, 1969; Murata, 1969). Under light conditions where PSII is preferentially excited, both PSII core and LHCII proteins become phosphorylated (Lemeille and Rochaix, 2010). As a consequence, phosphorylated LHCII proteins detach from PSII and partly connect to PSI (state 2). Under conditions where PSI excitation is predominant, this process is reversed. LHCII proteins are dephosphorylated and associate with PSII (state 1). The extent of state transition between vascular plants such as Arabidopsis and C. reinhardtii differs significantly. The proportion of mobile LHCII antenna is about 80% in the alga, whereas in Arabidopsis, only 15% to 20% of LHCII is transferred to PSI under state 2 conditions (Lemeille and Rochaix, 2010). However, the large increase in PSI antenna size in C. reinhardtii has recently been challenged (Nagy et al., 2014; Ünlü et al., 2014): while 70% to 80% of mobile LHCII detached from PSII in response to transition to state 2 conditions, only a fraction of about 20% functionally attached to PSI.Phosphorylation of LHC proteins requires the function of the STT7 kinase or its ortholog STN7 in C. reinhardtii or Arabidopsis, respectively. In the absence of the STT7/STN7 kinase, the initiation of state transitions is blocked (Depège et al., 2003; Bellafiore et al., 2005). The mobile LHCII fraction of C. reinhardtii includes the two monomeric minor LHCII antenna proteins, CP26 and CP29 (encoded by lhcb5 and lhcb4 genes), and the major chlorophyll a/b binding protein of LHCII, LHCBM5 (Takahashi et al., 2006), but also the LHCSR3 protein was suggested to migrate during state transitions (Allorent et al., 2013). Takahashi et al. (2014) suggested that only CP29 and LHCBM5 directly associate with PSI to form the PSI-LHCI-LHCII supercomplex, while the binding of CP26 could occur indirectly or via the other two proteins. However, it is not yet known whether STT7 directly phosphorylates the LHCII proteins or if this takes place as part of a kinase cascade (Rochaix, 2007). Nevertheless, the direct interaction between STT7 and the LHCII proteins is quite likely, since none of the other chloroplast kinases was found to be specifically required for LHCII phosphorylation (Rochaix, 2014). The activity of the STT7 kinase is mainly determined by the redox status of the plastoquinone pool (Vener et al., 1997; Zito et al., 1999). The identification of a PROTEIN PHOSPHATASE 2C (PP2C)-type phosphatase responsible for the dephosphorylation of the LHCII proteins in Arabidopsis has been described by two studies in parallel pointing to the fact that this enzyme, called PROTEIN PHOSPHATASE1/THYLAKOID-ASSOCIATED PHOSPHATASE38, acts directly on phosphorylated LHCII proteins, in particular when they are associated with the PSI-LHCI supercomplex (Pribil et al., 2010; Shapiguzov et al., 2010). Moreover, it is not known whether these phosphatases are constitutively active or if they are regulated by other means, for example through the redox state of the plastoquinone pool. Nonetheless, both enzymes are conserved in land plants and exhibit orthologous proteins in C. reinhardtii (Rochaix et al., 2012).Another kinase related to STN7/STT7 is encoded in the Arabidopsis and C. reinhardtii genomes and named STN8 and STATE TRANSITION-LIKE1 (STL1), respectively. STN8 is involved in PSII core subunit phosphorylation and influences the repair of PSII after photodamage (Bonardi et al., 2005; Vainonen et al., 2005). Remarkably, the disassembly of the PSII holocomplex is inhibited in STN7/STN8 double mutants (Tikkanen et al., 2008; Fristedt et al., 2009; Dietzel et al., 2011; Nath et al., 2013), suggesting that the phosphorylation of core subunits is required for PSII disassembly. It was further suggested that STN8 controls the transition between linear electron flow and CEF by the phosphorylation of PGRL1 in Arabidopsis (Reiland et al., 2011). As described for STN7, the activity of STN8 is probably regulated via the redox state of the plastoquinone pool (Bennett, 1991; Fristedt et al., 2009). Notably, the action of STN8 is counteracted by a chloroplast PP2C phosphatase (Samol et al., 2012), allowing for the fast reversibility of STN8-mediated acclimation responses. Thus, it appears that an intricate regulatory network of chloroplast protein kinases and phosphatases evolved in vascular plants and algae that drives the acclimation response to various environmental cues, including excess and changing light settings (Rochaix et al., 2012). As STN7/STT7 and STN8/STL1 kinase activities appear to be controlled by the redox poise of the plastoquinone pool, the plastoquinone pool would be a central player in these acclimation responses. On the other hand, the kinases themselves are subjected to phosphorylation (Reiland et al., 2009, 2011; Lemeille et al., 2010; Wang et al., 2013). However, the functional consequences of this phosphorylation are unknown.Recent comparative analyses revealed the presence of at least 15 distinct chloroplast protein kinases, suggesting an intricate kinase phosphorylation network in the chloroplast (Bayer et al., 2012). Generally, the phosphorylation of proteins is one of the most abundant posttranslational modifications. In complex eukaryotic systems, protein phosphorylation occurs most frequently on Ser followed by Thr residues, whereas protein phosphorylation of Tyr residues (1,800:200:1) is comparatively rare (Hunter, 1998; Mann et al., 2002). Protein phosphorylation is a general phenomenon in vivo; it is assumed that about one-third of all proteins are phosphorylated at a given time (Cohen, 2000; Ahn and Resing, 2001; Venter et al., 2001; Manning et al., 2002; Knight et al., 2003). A recent large-scale quantitative evaluation of human proteomic data strengthened the importance of protein phosphorylation for cellular function and human biology (Wilhelm et al., 2014). The C. reinhardtii and Arabidopsis genomes encode large kinase families (Arabidopsis Genome Initiative, 2000; Kerk et al., 2002; Merchant et al., 2007), supporting the view that protein phosphorylation also plays an important role in a plant’s life cycle. It is thus evident that the understanding of protein phosphorylation, including the specificity of residues phosphorylated or dephosphorylated in response to cellular as well as environmental factors, is one key to understanding the complex functional biological networks at the whole-system level. Likewise, it is crucial to design experimental setups allowing the linkage between phosphorylation events and particular physiological consequences to be elucidated.In this regard, we designed experiments to investigate STT7 kinase-dependent phosphorylation dynamics in C. reinhardtii in response to high light and anoxia, employing quantitative proteomics in conjunction with in-depth physiological characterization. These conditions are particularly interesting, as high light conditions are known to fully induce LHCSR3 protein expression and qE, while anoxia promotes CEF activity. Recently, it was demonstrated that qE and CEF are complementary and crucial in acclimation to these environmental cues (Kukuczka et al., 2014). Notably, LHCSR3 phosphorylation was suggested to depend on STT7 function (Bonente et al., 2011), while CEF supercomplex formation was found to be independent of STT7 kinase function (Takahashi et al., 2013), indicating that STT7 function might impact the acclimation to high light and anoxia in different ways. However, our quantitative proteomics and physiological data reveal that STT7-dependent variations in protein phosphorylation profiles have similar dramatic phenotypic consequences in both conditions, strongly suggesting that the regulation of protein phosphorylation is critical for driving successful acclimation to high light and anoxic growth environments.     

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.     

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.     

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.     

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).