Sulfur Assimilation in Photosynthetic Organisms

PP62CH07-Takahashi

ARI

ANNUAL REVIEWS

4 April 2011

14:47

Further

Annu. Rev. Plant Biol. 2011.62:157-184. Downloaded from www.annualreviews.org by John Innes Centre on 05/18/11. For personal use only.

Click here for quick links to Annual Reviews content online, including: • Other articles in this volume • Top cited articles • Top downloaded articles • Our comprehensive search

Sulfur Assimilation in Photosynthetic Organisms: Molecular Functions and Regulations of Transporters and Assimilatory Enzymes Hideki Takahashi,1,2 Stanislav Kopriva,3 Mario Giordano,4 Kazuki Saito,1,5 and Rudiger Hell6 ¨ 1

RIKEN Plant Science Center, Yokohama 230-0045, Japan

2

Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, Michigan 48824; email: [email protected] 3 John Innes Centre, Norwich NR4 7UH, United Kingdom; email: [email protected] 4

Department of Marine Sciences, Università Politecnica delle Marche, 60131 Ancona, Italy; email: [email protected]

5 Graduate School of Pharmaceutical Sciences, Chiba University, Chiba 263-8522, Japan; email: [email protected] 6 Heidelberg Institute for Plant Sciences, University of Heidelberg, 69120 Heidelberg, Germany; email: [email protected]

Annu. Rev. Plant Biol. 2011. 62:157–84

Keywords

First published online as a Review in Advance on March 3, 2011

sulfur, sulfate, transport, metabolism, regulation, signaling

The Annual Review of Plant Biology is online at plant.annualreviews.org

Abstract

This article’s doi: 10.1146/annurev-arplant-042110-103921 c 2011 by Annual Reviews. Copyright All rights reserved 1543-5008/11/0602-0157$20.00

Sulfur is required for growth of all organisms and is present in a wide variety of metabolites having distinctive biological functions. Sulfur is cycled in ecosystems in nature where conversion of sulfate to organic sulfur compounds is primarily dependent on sulfate uptake and reduction pathways in photosynthetic organisms and microorganisms. In vascular plant species, transport proteins and enzymes in this pathway are functionally diversified to have distinct biochemical properties in specific cellular and subcellular compartments. Recent findings indicate regulatory processes of sulfate transport and metabolism are tightly connected through several modes of transcriptional and posttranscriptional mechanisms. This review provides up-to-date knowledge in functions and regulations of sulfur assimilation in plants and algae, focusing on sulfate transport systems and metabolic pathways for sulfate reduction and synthesis of downstream metabolites with diverse biological functions.

157

PP62CH07-Takahashi

ARI

4 April 2011

14:47

Contents


INTRODUCTION . . . . . . . . . . . . . . . . . . SULFATE TRANSPORT SYSTEMS . . . . . . . . . . . . . . . . . . . . . . . . Sulfate Transport Mechanisms . . . . . Functions of Sulfate Transporters in Plants . . . . . . . . . . . . . . . . . . . . . . . . SULFATE REDUCTION AND METABOLISM . . . . . . . . . . . . . . . . . . . Pathways for Sulfate Reduction . . . . . Cysteine Biosynthesis . . . . . . . . . . . . . . Glutathione Biosynthesis and Cellular Activities . . . . . . . . . . . . . . . Methionine Biosynthesis and Its Control . . . . . . . . . . . . . . . . . . . . . Synthesis of Sulfated Compounds . . . REGULATORY MECHANISMS . . . . Environmental Factors and Intrinsic Signals Modulating Sulfate Uptake and Reduction . . . . . . . . . . Regulatory Components of Sulfate Transport and Assimilation . . . . . . Cysteine Synthase Complex . . . . . . . . Posttranscriptional Regulation of Methionine Biosynthesis . . . . . . . . Systems Regulation . . . . . . . . . . . . . . . . Sulfur Acclimation Signals in Chlamydomonas . . . . . . . . . . . . . . . . . . IMPACT OF SULFUR AVAILABILITY ON THE EVOLUTION OF PHOTOSYNTHETIC ORGANISMS . . . . . . . . . . . . . . . . . . . . . CONCLUSIONS . . . . . . . . . . . . . . . . . . . .

158 159 159 160 162 162 163 164 165 166 166

166 167 169 170 170 171

172 173

INTRODUCTION

DMSP: dimethylsulfoniopropionate DMS: dimethylsulfide

158

Sulfur is an essential element cycling through the global ecosystem (Figure 1). Sulfur is present in nature in both inorganic and organic forms. Sulfate (SO4 2− ) contains sulfur in the +VI redox state and is the most oxidized and most common form of sulfur in nature. Plants and microorganisms reduce sulfate to sulfide and incorporate it into organic metabolites.

Takahashi et al.

Reduction of sulfate to sulfide is an energydependent assimilative process. Herbivores rely on reduced sulfur compounds produced by plants. Organic sulfur in the waste and remains of plants and animals can be degraded by microorganisms and then regenerated to inorganic sulfate to close the cycle. Under anaerobic environment, sulfate- and sulfur-reducing bacteria may obtain energy from dissimilative sulfate reduction. Sulfur is also cycled in the atmosphere. Algae synthesize dimethylsulfoniopropionate (DMSP), which becomes volatile as dimethylsulfide (DMS), a portion of which is then released to the atmosphere. Sulfide and sulfur dioxide gasses erupt from volcanoes, and hot springs and heavy industries are additional sources of volatile sulfur. These volatile compounds are oxidized to sulfate in the atmosphere and recycled to biosphere as rain falls. Sulfur is therefore cycled in soil, water and atmospheric environments globally as minerals and volatiles, and they are metabolized in the assimilative and dissimilative pathways of living organisms (reviewed in 49, 121, 145, 173). Photosynthetic organisms synthesize a wide variety of sulfur compounds, using sulfate as a primary sulfur source (121, 173). Sulfur is present in major cellular components essential for the maintenance of cell viability. Besides the S-amino acids, cysteine and methionine, sulfur is contained in membrane sulfolipids (10) and cell walls (156a). Thiols of proteins and peptides and the tripeptide glutathione serve for redox control (40). Sulfur is also present in vitamins and cofactors such as thiamine, biotin, and coenzyme A. Plant hormones, brassinosteroid and jasmonate, are deactivated by sulfation (44, 127). By contrast, sulfation is essential for the function of peptide hormones stimulating plant cell growth (2, 136). Glucosinolates and alliins are secondary metabolites characteristic of Brassicaceae and Alliaceae plant species (60, 83). Their breakdown products are pungent and odorous, and they may work as repellants against predators (9). Besides being important defense chemicals in plants, glucosinolates and alliins are beneficial for humans, as they

PP62CH07-Takahashi

ARI

4 April 2011

14:47

Atmosphere SO2 SO2 SO42–

SO42– SO2

Ocean

Algae

SO42–

Plants Assimilation

DMS

Assimilation


S2–

Fossil resources

DMSP

R-SH Mineralization

Soil microorganisms

Sediments

S2–

Mineralization

Land SO42–

Dissimilative reduction S0

Dissimilative oxidation

Figure 1 Biogeochemical cycle of sulfur in nature. Sulfate is assimilated by plants ( green arrow) and algae (blue arrow). Soil microorganisms use sulfur in dissimilative reactions and mineralize organic sulfur compounds to sulfate. Sulfur can be released to the atmosphere as volatile compounds from oceans and volcanoes and through anthropogenic activities. The volatile sulfur compounds are oxidized to sulfate in the atmosphere.

induce detoxifying enzymes that prevent tumor formation (12, 138, 191). Choline-O-sulfate is another sulfated secondary metabolite identified as a potent osmoprotectant in some plant species (63). Given the importance of sulfur compounds for the life cycle of photosynthetic organisms, functions of transport proteins and enzymes involved in sulfate metabolism have been intensively investigated over the past few decades. Since 2000, functional genomics of the model plant species, Arabidopsis thaliana, has provided us with a more precise understanding of its physiological functions and regulations. In this review, we focus on the molecular functions of components of sulfur transport and metabolism (Figures 2 and 3), referring to biochemical and genetic backgrounds. We also describe how these metabolic pathways and upstream regulatory processes are integrated to balance the systems in response to changes in environmental sulfur conditions and intrinsic signals. A brief overview of how sulfur may have influenced the evolution and radiation of photosynthetic taxa is also given.

SULFATE TRANSPORT SYSTEMS Sulfate Transport Mechanisms Influx of sulfate through plasma membranebound transport proteins occurs against the inside-negative gradient of membrane potential, requiring a driving force for transport. Plants primarily use proton/sulfate cotransport systems to mediate sulfate influx (118, 185). This system utilizes proton gradients across the membranes as motive force, and the kinetic phase with a low Km value becomes active under sulfur-limited conditions (24, 118). Sulfate transporters are structurally related to the family of membrane-bound solute transporters predicted to have 12 membrane-spanning domains (185). In addition, they contain STAS (sulfate transporters and antisigma factor antagonists) domains in C termini, which may have regulatory functions in controlling activity and localization of transporters to membranes (170, 180). A completely different mechanism facilitates acquisition of sulfate in bacteria. The bacterial sulfate transporting complex consists of a

www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

159

PP62CH07-Takahashi

ARI

4 April 2011

14:47

Shoot

Mesophyll cells

Xylem parenchyma cells and Xylem phloem parenchyma cells

Vacuoles SO42–

SO42– storage

SO42–

Arabidopsis thaliana

SULTR2;1


Chloroplasts SO42– reduction

Epidermis

Cortex

SO42–

SO42–

SO42–

Endodermis

SO42–

Pericycle and xylem parenchyma cells Xylem SO42– SULTR4;1 SULTR4;2

SO42–

SO42–

SO42–

SO42–

SO42–

SULTR1;1 SULTR1;2

SO42–

SULTR2;1 SULTR3;5 Apoplastic SO42–

Apoplastic SO42–

Root Figure 2 Sulfate transport systems in Arabidopsis. Red circles on arrowed lines indicate sulfate transporters (SULTR) mediating influx of sulfate at the plasma membrane or release of sulfate from the vacuoles. Dashed lines indicate yet unknown transport pathways that may function for efflux of sulfate.

sulfate binding protein, two membrane-bound proteins and an ATP-binding protein that hydrolyzes ATP to drive the import of sulfate (119, 184). Interestingly, a similar complex exists in chloroplasts of green algae but not in higher plants (123), suggesting evolutionary diversification of sulfate acquisition systems among photosynthetic organisms. The identification of a plastidic sulfate transporter in higher plants and other phosynthetic taxa remains one of the most important tasks for sulfur research. Sulfate efflux is important for distribution of sulfur throughout plant organs. Sulfate 160

Takahashi et al.

probably can be released to the outside of the cell through a passive mechanism driven by outside-positive gradients of membrane potentials at the plasma membranes (188). A similar mechanism is postulated for import of sulfate to vacuoles (188) driven by the positive electrochemical gradients generated by tonoplast H+ -ATPase or H+ -pyrophosphatase (129).

Functions of Sulfate Transporters in Plants Genes encoding sulfate transporters were first identified by functional complementation of

PP62CH07-Takahashi

ARI

4 April 2011

14:47

Cytosol Chloroplasts/plastids

TS

Thr

OPH SO42–

ATPS

APS

APR

SO32–

SiR

OAS-TL S2– Cys OAS

APK

R-OH


SO42–

SULTR

SO42–

ATPS

APK

SOT

Sulfated compounds

APS

Vacuoles

SULTR SO42–

GSH

Cys

X GST GS-X

Ser SAT OAS

X-CysGly Glu

SAM SAM

GSHS γ-GluCys

OAS OAS-TL

MS

SAH Met SAM

SAH

GSH

Ser SAT S2–

Hcy

Met

CBL

CGS γ-ECS

CLT PAPS

MS

γ-GluCys GSHS

SAT Ser

PAPS

Cyst

Hcy

S2–

GGT

OAS-TL

Cys

Mitochondria

MRP GS-X

Figure 3 Cellular organization of sulfur metabolism. Enzymes and transporters are indicated in red characters. Dashed lines indicate putative pathways for metabolite transport. Abbreviations of metabolites: APS, adenosine 5 -phosphosulfate; Cys, cysteine; Cyst, cystathionine; Glu, glutamate; γ-GluCys, γ-glutamylcysteine; GSH, glutathione; GS-X, glutathione conjugate; Hcy, homocysteine; Met, methionine; OAS, O-acetylserine; OPH, O-phosphohomoserine; PAPS, 3 -phosphoadenosine 5 -phosphosulfate; R-OH, hydroxylated precursor; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine; Ser, serine; Thr, threonine; X-CysGly, cysteinylglycine conjugate. Abbreviations of enzymes and transporters: APK, APS kinase; APR, APS reductase; ATPS, ATP sulfurylase; CBL, cystathionine β-lyase; CGS, cystathionine γ-synthase; CLT, thiol transporter (chloroquine resistance transporter-like transporter); γ -ECS, γ-glutamylcysteine synthetase; GGT, γ-glutamyltransferase; GSHS, glutathione synthetase; GST, glutathione-S-transferase; MRP, multidrug resistance-associated protein; MS, methionine synthase; OAS-TL, OAS(thiol)lyase; SAM, S-adenosylmethionine synthetase; SAT, serine acetyltransferase; SiR, sulfite reductase; SOT, sulfotransferase; SULTR, sulfate transporter; TS, threonine synthase.

yeast strains that contain mutations in sulfate transporters (185, 187). A number of sulfate transporters have been cloned from various plant species. The family of plant sulfate transporter is comprised of 12–16 genes from each plant species and can be classified into four distinct functional groups (188). For Arabidopsis, the analysis of knockout mutants confirmed that two high-affinity sulfate transporters, SULTR1;1 and SULTR1;2, facilitate uptake of sulfate into roots especially under

sulfur-limited conditions (Figure 2) (5, 208). Accordingly, they are predominantly expressed in root hairs, root epidermal, and cortical cells (182, 189, 209). The phase I transport system designated by earlier physiological measurements of sulfate influx activity in plant roots has low Km for sulfate and is activated by sulfur limitation (24, 118, 120). Although certain degrees of redundancy of functionality and spatial expression patterns may exist, SULTR1;2 is considered to be the major component. By


SULTR: sulfate transporter

161

ARI

4 April 2011

14:47

contrast, SULTR1;1 may represent a more specialized component for the uptake of trace sulfate as it has a lower Km value and is more strongly induced by sulfur limitation (189, 209). Once sulfate is absorbed into epidermis cells, it can be transferred to the central cylinder through cell-to-cell plasmodesmal connections (Figure 2). This seems to be the way sulfate crosses the barrier of the Casparian strip at the endodermal cell layers (188). Although sulfate is transferred horizontally from the epidermis to the central cylinder and within the cells in the central cylinder, it can leak from the symplast (cells) to the apoplast (cell wall space), probably through a yet unidentified passive-transporting mechanism. SULTR2;1 is a low-affinity sulfate transporter expressed in the central cylinder, and its induction by sulfur limitation suggests its role in facilitating sulfate translocation from the roots to the shoots (88, 189, 190). SULTR2;1 has been postulated to mediate the influx of sulfate in the xylem parenchyma cells where the concentration of sulfate in the symplast needs to be elevated for an unknown passive mechanism to be functional in facilitating the efflux of sulfate to the xylem (188). Interestingly, SULTR3;5 cofacilitates the influx of sulfate through SULTR2;1, however, it is not functional as the sulfate transporter by itself (88). In barley, a high-affinity sulfate transporter, HVST1 (186), is expressed in pericycle and xylem parenchyma cells and is likely involved in the retrieval process in the central cylinder (159). Besides these mechanisms, tonoplast-localized sulfate transporters, SULTR4;1 and SULTR4;2, release sulfate from the vacuoles and increase symplastic fluxes of sulfate before entering the xylem (89). SULTR2;1 was also suggested to control the transfer of sulfate to developing seeds (4). Another phloem-localized sulfate transporter, the high-affinity SULTR1;3, is present in phloem companion cells and controls transfer of sulfur from source to sink organs (207). SULTR2;2 is also expressed in the phloem but shows low substrate affinity (189). These transport systems absorb sulfate to the companion cells and may control the amounts of sulfur transported


PP62CH07-Takahashi

162

Takahashi et al.

in the phloem sap. In addition to sulfate, glutathione and S-methylmethionine are forms of sulfur transported in the phloem (15, 69, 117). Inside the cells, sulfate is either stored in vacuoles or metabolized in chloroplasts and plastids. As mentioned in the previous section, the influx of sulfate to the vacuoles likely occurs following inside-positive electrochemical gradients. In contrast, transporters responsible for the vacuolar efflux of sulfate have been identified from Arabidopsis as belonging to the SULTR4 group (89). Accumulation of transcripts for SULTR4;1 and SULTR4;2 under sulfur limitation indicates that the release of the vacuolar sulfate is promoted when demands for sulfur increase (89). For sulfate to enter the chloroplasts, a bacterial-type sulfate transporter complex is essential in green alga Chlamydomonas reinhardtii (123). However, a homologous system is absent in either nuclear or chloroplast genomes of flowering plants, suggesting the system may be completely different or nonspecific. Altogether, the members of the plant sulfate transporter gene family are classified into four distinct groups according to their sequences, biochemical characteristics, and physiological functions (Figure 2) (188). Among the 12 members in Arabidopsis, those in Group 3 are least characterized and their activity as sulfate transporters has not been confirmed yet. However, in Lotus japonicus the activity of a SULTR3 homolog is important for functional symbiosis with Rhizobia (114). Previously, sulfate transporter–like proteins having no N- and C-terminal hydrophilic extensions including STAS domains were designated as Group 5 sulfate transporters (188). However, SULTR5;2 functions as a molybdate transporter, MOT1 (8, 192).

SULFATE REDUCTION AND METABOLISM Pathways for Sulfate Reduction The long-standing controversies about free and bound pathways of sulfate reduction and the


PP62CH07-Takahashi

ARI

4 April 2011

14:47

substrates of sulfate-reducing enzymes in plants seem to have been resolved both by biochemical and genetic approaches and by the increased availability of genomic sequences (103). Before reduction, sulfate has to be activated by adenylation catalyzed by ATP sulfurylase (ATPS) (EC: 2.7.7.4). The resulting adenosine 5 -phosphosulfate (APS) forms a branching point in the pathway (Figure 3). In the primary sulfate assimilation, APS is first reduced by APS reductase (APR) (EC: 1.8.99.2) to sulfite, which is further reduced to sulfide by ferredoxin dependent sulfite reductase (SiR) (EC: 1.8.7.1). Sulfide is then incorporated into the amino acid skeleton of O-acetylserine to form cysteine. APS can, however, also be phosphorylated by APS kinase (APK) (EC: 2.7.1.25) to form 3 phosphoadenosine 5 -phosphosulfate (PAPS), a donor of activated sulfate for many sulfation reactions. In all photosynthetic organisms, sulfate reduction occurs in the plastids, with the notable exception of Euglena gracilis, which locates the sulfate-reducing enzymes in mitochondria (16, 154). Whereas the reductive steps are localized exclusively in the organelles, ATP sulfurylase is also present in the cytosol (126, 166, 169). Accordingly, plants and algae possess multiple isoforms of ATP sulfurylase (105, 154). Very surprisingly, plant ATP sulfurylases have a different evolutionary origin than those from green algae, and they are more similar to their animal counterparts (154). Eukaryotic marine microalgae, such as diatoms and haptophytes, possess both types of ATP sulfurylase, the plant-like isoform forming a fusion with APS kinase and inorganic pyrophosphatase that most probably increases catalytic efficiency of the otherwise very inefficient enzyme. APR is also encoded by small multigene families in most photosynthetic organisms, and similar to ATP sulfurylase, two distinct isoforms exist (102, 154). APR of flowering plants and green algae is a multidomain protein consisting of an N-terminal reductase domain and a C-terminal thioredoxin/glutaredoxin-like part (13, 56, 103, 179). The reductase domain binds an [Fe4 S4 ] cluster as a cofactor (94, 100, 101).

Although the properties of the cluster are well described, its exact function in the reaction mechanism of APR remains unknown (94, 101). Another isoform of APR has initially been identified in the moss Physcomitrella patens (106). This enzyme, APR-B, does not require the FeS cluster for activity, resulting in lower catalytic efficiency but greater stability. APR-B was found in several basal plant species and, interestingly, is the only isoform present in the marine microalgae sequenced so far (105, 154). Phylogenetic analyses suggest that the FeSbinding APR is the ancestral isoform. APR-B may thus represent an adaptation to environments with low iron availability (154). Sulfite reductase catalyzes the six-electron reduction of sulfite to sulfide. The enzyme is dependent on siroheme and FeS centers as prosthetic groups (112). In contrast to the upstream enzymes, generally lower number of genes and the same type of sulfite reductase exist in all photosynthetic organisms (93, 105, 154). An interesting addition to the scheme of sulfate assimilation was the identification of plant sulfite oxidase (33). This peroxisome-localized enzyme possesses molybdenum cofactor and oxidizes sulfite to sulfate, transferring the electrons to molecular oxygen to form hydrogen peroxide (61). The contribution of sulfite oxidase to sulfur flux in the cell is not clear yet, nor is its biological function, apart from conferring resistance to high levels of sulfur dioxide (17).

APS: adenosine 5 -phosphosulfate APR: adenosine 5 -phosphosulfate reductase PAPS: 3 -phosphoadenosine 5 -phosphosulfate SAT: serine acetyltransferase OAS: O-acetylserine OAS-TL: O-acetylserine (thiol) lyase

Cysteine Biosynthesis Sulfide generated by sulfite reductase is the substrate for cysteine biosynthesis (Figure 3) (68, 173). To integrate free sulfide into the carbon skeleton, serine is first activated by serine acetyltransferase (SAT) (Serat; EC 2.1.3.30) that uses acetyl coenzyme A to form O-acetylserine (OAS). In the second step the acetyl moiety is exchanged by sulfide in a β-replacement reaction catalyzed by O-acetylserine (thiol) lyase (OAS-TL) [β-substituted alanine synthase (Bsas); EC 2.5.1.47]. Bacteria use the same two-step pathway to synthesize cysteine, whereas most


163

ARI

4 April 2011

14:47

fungi use methionine as the first organic sulfur compounds and animals require methionine in their diet as a source of reduced sulfur. SAT and OAS-TL in bacteria and plants are associated in a hetero-oligomeric cysteine synthase complex (68, 110, 173, 200). So far all SAT activity in protein extracts has been found in association with OAS-TL, whereas free and active OAS-TL homodimers are present in excess. OAS-TL is inactive in the cysteine synthase complex, but the excess of free OAS-TL seems to be required to achieve full SAT activity and subsequent conversion of OAS to cysteine. Consequently, no substrate channeling takes place (26, 31). OAS has a dissociating effect on the cysteine synthase complex that is overruled by the stabilizing action of sulfide, pointing to a metabolic mechanism of regulation (68, 173, 200). The mechanism of inhibition of OAS-TL by interaction with SAT was elegantly resolved by binding and structural studies. The C terminus of SAT is highly flexible and binds to the catalytic grove of OAS-TL. The block of enzymatic activity can be released by high concentrations of OAS that compete for binding with the C terminus of SAT (41, 80). The evidence from three-dimensional structures, modeling and biophysical analyses of bacterial and Arabidopsis OAS-TLs and SATs indicates that free homohexameric SAT is a dimer of two trimers (157) that presumably also forms the core of the complex with one OAS-TL dimer attached at opposing ends of the SAT hexamer (37, 199a). However, a recent contradictory report describes the quaternary structure of a cysteine synthase complex from soybean cytosol as one SAT trimer with up to three OAS-TL dimers bound to the three C termini of SATs (115), suggesting isoform- and speciesspecific differences in arrangements of the complex structures. SAT and OAS-TL are ubiquitously expressed in plant cells and are encoded by several nuclear genes (68). Cysteine can be synthesized in the cytosol, plastids, and mitochondria in nearly all plant species (Figure 3). Null mutants of each of the major OAS-TLs A, B, and C of


PP62CH07-Takahashi

164

Takahashi et al.

the three compartments are also well able to develop (66, 197). This suggests efficient exchange of sulfide, OAS, and cysteine across organelle membranes. In Arabidopsis the three major SAT isoforms (Serat1;1, Serat2;1, and Serat2;2) are targeted to these three compartments (144) and interact with OAS-TLs (66). Two minor SAT isoforms (Serat3;1 and Serat3;2) in the cytosol have long C-terminal extensions and are believed to be unable to interact with OASTLs. Studies of various SAT mutants showed that OAS is predominantly synthesized in mitochondria and that cytosol, but not plastids, is the major compartment for cysteine synthesis (57, 198). These observations fit well with the distribution of 125-μM sulfide and 9-μM cysteine in chloroplasts and 125-μM sulfide and 332-μM cysteine in the cytosol of Arabidopsis (113). Physiological significance of OAS production by mitochondrial SAT (Serat2;2/SAT3) was demonstrated by severe growth phenotypes and reduced OAS levels of Arabidopsis plants in which expression of SERAT2;2/SAT3 was silenced (57, 198). In contrast to the high production rate in mitochondria, OAS is a limiting factor for cysteine biosynthesis in chloroplasts (175). Oxidative stress appears to be a trigger for enhanced expression or activity of chloroplastic SAT (Serat2;1) (30, 158). Taken together, these current studies suggest a scenario in a leaf where chloroplasts generate sulfide via reductive sulfate assimilation, the mitochondria provide the bulk of OAS, and the cytosol produces most of the cysteine (57, 198). Besides cysteine synthesis, other enzymatic functions are suggested for OAS-TL-like isoforms, including cyanoalanine synthase in mitochondria (65), S-sulfocysteine synthase (CS26) in plastids (11), and L-cysteine desulfhydrase in cytosol (1).

Glutathione Biosynthesis and Cellular Activities The manifold functions of the tripeptide γ-glutamylcysteinylglycine (glutathione) include scavenging and detoxification of reactive oxygen species (ROS), heavy metals and xenobiotics, catalytic sulfur donation, transport and


PP62CH07-Takahashi

ARI

4 April 2011

14:47

intermediary storage of sulfur, redox signaling, and others (40). The most relevant enzymes utilizing glutathione are glutathione reductase, γ-glutamyltransferase (GGT), glutathioneS-transferase (GST), and glutaredoxins. The glutathione redox function is such an important task that cytosol and mitochondria of Arabidopsis host a back-up system in case of insufficient glutathione reduction (130). Recent examples demonstrating the importance of glutathione redox homeostasis include its role in the transmission of cotranslational proteolytic processing defects (43) and in enabling auxin transport and signaling (7, 107). Glutathione is synthesized in two ATPdependent steps that are catalyzed by γglutamylcysteine synthetase (γ-ECS) and glutathione synthetase (GSHS) (Figure 3). γECS catalyzes the regulatory step, and is subject to transcriptional control in response to environmental perturbations (204). The structure and activity of γ-ECS are redox sensitive: Fully reduced γ-ECS has only little activity, whereas the fully oxidized form is highly active and has a higher apparent molecular weight (67). The three-dimensional structure of γ-ECS reveals the redox control of its conformation by two intramolecular disulfide bridges (53, 71, 79, 81). In many plants, γ-ECS is localized only to plastids,while in pea and spinach leaves, subcellular fractionation experiments showed that γ-ECS activity is present in both chloroplasts and cytosol (67, 95). GSHS activity in Arabidopsis is distributed between plastids and the cytosol as a result of differential splicing of the same GSHS transcript (53, 195). Arabidopsis T-DNA mutants of GSHS are seedling lethal, but fully complemented with cytosol-targeted GSHS, showing that glutathione can enter plastids (153). An additional recent study indicates Arabidopsis membrane proteins CLTs similar to chloroquine transporters of the malaria parasite Plasmodium falciparum are responsible for transporting γ-glutamylcysteine and glutathione from chloroplast to cytosol and contribute to glutathione homeostasis (137) (Figure 3). Turnover of glutathione is tightly connected to GGT activities. Of the three

active GGTs in Arabidopsis, GGT4 cleaves glutathione-S-conjugates at the internal side of the tonoplast (54; named GGT3 in 150) after transport of conjugates to the vacuole by MRP-type ABC transporters as part of the phase II detoxification of xenobiotics (165). GGT1 and GGT2 are associated with the outer surface of the plasma membrane and seem to be involved in cellular glutathione uptake and long-distance transport, respectively. GGT1 responds to oxidative stress, similar to an undefined GGT activity that contributes to degradation of GSSG (oxidized glutathione) in the apoplast of barley roots (38, 149). GGT2 may be involved in GSH breakdown and transport into siliques (128, 149). Intracellular degradation of free glutathione in cytoplasm is probably initiated in a GGT-independent way by γ-glutamylcyclotransferase that releases 5-oxoproline and cysteinylglycine as evidenced from analysis of 5-oxoprolinase knockout mutants (148). The molecular identity of γ-glutamylcyclotransferase is currently the missing link to establish a γ-glutamyl cycle similar to the one in animals but with plant-specific functions.

Methionine Biosynthesis and Its Control Cystathionine γ-synthase (CGS) and cystathionine β-lyase (CBL) catalyze the two consecutive steps of methionine biosynthesis in chloroplasts (164) (Figure 3). Cysteine and O-phosphohomoserine (OPH) are the substrates for synthesis of cystathionine by CGS. Cleavage of cystathionine by CBL generates homocysteine. Homocysteine is subsequently methylated to form methionine by methionine synthase (MS) using methyltetrahydrofolate as a methyl donor. This final step is present in both chloroplasts and cytosol (32, 162). Methionine is further converted to S-adenosylmethionine (SAM) in cytosol. SAM serves as a methyl donor but recycles back to S-adenosylhomocysteine (SAH) and then to homocysteine (162). Transport of methonine, SAM, and SAH across the chloroplast envelope is essential to fulfill the


165

ARI

4 April 2011

14:47

requirement of SAM in chloroplasts and to drive the metabolic cycle (162) (Figure 3). Control of methionine biosynthesis occurs at the junction of pathways synthesizing methionine and threonine. Although OPH is a common substrate for both threonine synthase (TS) and CGS, TS has an extremely higher affinity for it than does CGS (27, 163). In addition, SAM, the product of methionine biosynthetic pathway, enhances the activity of TS and its affinity to OPH (27). The major flux of carbon flows to the threonine biosynthetic pathway when methionine and SAM are sufficiently accumulated. Pathways for methionine biosynthesis will receive a supply of carbon skeletons when SAM levels decline and TS becomes less active (3, 70).


PP62CH07-Takahashi

Synthesis of Sulfated Compounds Sulfation is a common modification of proteins and metabolites in plant cells. The best known examples of sulfated compounds are the glucosinolates in Brassicaceae (60) and the peptide hormones phytosulfokines (136). Glucosinolates are an important form of defense against herbivores (60): Their breakdown products are both toxic and deterrent to the attacker (9), but they are also components of general pathogen defense and have anticarcinogenic activity in humans (9, 138, 191). Phytosulfokines are sulfated pentapeptides that stimulate cell proliferation (135, 136). Disruption of the phytosulfokine receptor leads to premature senescence of rosette leaves and a loss of the ability to form calli (135). Another sulfated oligopeptide, PSY1, was recently isolated from Arabidopsis and shown to be important for normal development (2). Both peptides require tyrosine sulfation for biological activity (2, 136). Sulfotransferases catalyzing sulfation of hydroxylated precursors and sulfation of protein tyrosine residues use PAPS as the source of the sulfo group (96, 99). The sulfotransferase gene family in Arabidopsis contains 18 members (96), although the substrate specificity is known in only seven isoforms including those for sulfation of glucosinolates (73, 155), flavonoids (45), 166

Takahashi et al.

jasmonate (44), and brassinosteroid (127). The great importance of tyrosine sulfation of peptide hormones has been revealed by a strong dwarf phenotype of plants lacking tyrosine sulfotransferase (99). Similarly, Arabidopsis mutants lacking two isoforms of APS kinase and thus limiting synthesis of PAPS displayed a semidwarf phenotype and accumulated only 10–15% of glucosinolates compared with wildtype plants (139).

REGULATORY MECHANISMS Environmental Factors and Intrinsic Signals Modulating Sulfate Uptake and Reduction Sulfate uptake and assimilation are tightly regulated according to the plant demands for reduced sulfur. In the past decade, molecular studies, particularly gene expression profiling, brought about a significant increase in our understanding of the physiological and molecular responses of plant sulfur metabolism to environmental factors and to disturbances in levels of pathway intermediates and products. The interconnection of sulfate assimilation with nitrogen and carbon metabolism as well as the role of the pathway for stress response have been firmly established (14, 104, 108, 109, 125). Control of sulfur assimilation occurs primarily at the steps of sulfate uptake and APS reduction, where the transcript levels of sulfate transporters and APR are strictly regulated by sulfate availability and well correlated with the fluxes of sulfate import and reduction (178, 193, 208). Besides transcriptional regulation of the sulfate transporter and APR, sulfite reductase and mitochondrial SAT strongly participate in controlling sulfur metabolism, as their inadequate activity results in severe growth defects (57, 93, 198). Although there is usually a good correlation between regulation of mRNA levels and enzyme/transporter activities, many components of sulfur metabolism are additionally controlled by complex interplays of transcriptional and posttranscriptional regulations (108, 208).


PP62CH07-Takahashi

ARI

4 April 2011

14:47

Several groups of metabolites are assigned important roles in the regulation of sulfate assimilation. The pathway is under biochemical control by the reaction products and intermediates. Feedback inhibition of sulfate uptake, APR, and γ-ECS by GSH is an integral part of the demand-driven control of the pathway (79, 117, 193). By contrast, OAS, the intermediate of cysteine biosynthesis, is an important positive regulator of the pathway (108, 140, 186). OAS accumulates during sulfur deficiency and affects expression of a similar set of genes (72). Although accumulation of OAS triggers a response similar to that of sulfate starvation (147) and regulation of a subset of sulfate assimilation genes by this condition correlates with OAS levels (73), other reports showed an uncoupling of the sulfate starvation response from OAS (77). The function of OAS as a general signal of internal S status remains questionable, though its apparent function in decomposing the cysteine synthase complex suggests a significant role of this intermediary metabolite in regulation. Among other compounds affecting sulfate assimilation, the phytohormones play the most prominent roles. This is not surprising for the stress-related hormones, jasmonate, abscisic acid, and salicylate, as sulfur-containing compounds have an important function in plant stress defense (161). Indeed, jasmonate coordinately induces multiple genes of sulfate assimilation (64, 86, 177). Conversely, genes involved in jasmonate synthesis are upregulated in sulfur-starved plants (72, 131, 141) and jasmonate levels are affected in plants with disturbed PAPS metabolism (139, 168). Abscisic acid specifically induces cytosolic OAS-TL (6) but reduces activity of APR by an unknown posttranscriptional mechanism (108). Treatment with salicylate results in increased GSH levels (39). High salicylate levels have also been implicated in tolerance to nickel, via induction of SAT activity and increase in GSH (42). Both jasmonate and salicylate induce mRNA accumulation and activity of APR (108). Another group of phytohormones directly involved in regulation of sulfate uptake and assimilation are the cytokinins. Signals under cytokinin and

its receptor repressed the expression of highaffinity sulfate transporters and sulfate uptake capacity of Arabidopsis roots (134). In contrast, treatment with zeatin induced accumulation of transcripts for SULTR2;1 and APR (108, 146).

Regulatory Components of Sulfate Transport and Assimilation As described in the previous section, sulfate transport and assimilation are regulated by complex mechanisms that involve signals specific for sulfur or general intrinsic signals. However, actual regulatory components have been identified only very recently. New approaches in genetic screens, multi-omics, and functional genomics deciphering cellular responses to sulfur limitation allowed us discoveries of key regulators in the pathway (Figure 4). SULFUR LIMITATION1. SLIM1 is a key transcriptional regulator of sulfate uptake identified from a genetic screen for Arabidopsis mutants disrupted in the sulfur-limitation response, using GFP controlled by a sulfur starvation–inducible SULTR1;2 promoter as a reporter (132). The slim1 mutant was impaired in inducing GFP fluorescence and native SULTR1;2 transcripts, and it showed reduced sulfate uptake and growth under sulfur-limited conditions. Transcriptome analysis of a slim1 mutant indicated that sulfate transporters and enzymes for glucosinolate biosynthesis are regulated oppositely by SLIM1 in response to sulfur limitation (132). SLIM1 globally controls the balance of sulfur utilization as an upstream coordinator (Figure 4). Among the genes regulated by sulfur limitation, APR seems to be an exception as its transcript levels are not regulated by SLIM1. Independent regulatory mechanisms thus have to be postulated for APR. SLIM1 belongs to a group of ethyleneinsensitive3-like (EIL)-family transcription factors (132). However, its function seems to be specific to the sulfur response (132) and can be distinguished from those for EIN3 and its homologues, EIL1 and EIL2, that predominantly mediate the ethylene response (55).


167

PP62CH07-Takahashi

ARI

4 April 2011

14:47

a

SO42– uptake

SO42–

Glucosinolates

Internal

SO42–

SO42–

Arabidopsis thaliana

transport

miR395

APS

SLIM1 Nitrogen assimilation

?

Photosynthesis

OAS CSC

Cys Trp

Met

–S Auxin

GSH


MYBs

Jasmonate

Glucosinolate biosynthesis

b

SAC1

Chlamydomonas reinhardtii

SNRK2.2 Sulfate uptake Sulfur assimilation

SNRK2.1

Photosynthesis ROS

Figure 4 Regulatory pathways and components in the sulfur-limitation (−S) response of Arabidopsis thaliana (a) and Chlamydomonas reinhardtii (b). Red lines indicate pathways that specifically control sulfate transport and metabolism. Gray lines indicate other general pathways affected by sulfur deficiency.

microRNA-395. miR395 is involved in the regulation of sulfur metabolism. It accumulates under sulfur starvation and targets three ATP sulfurylase isoforms, ATPS1, ATPS3, ATPS4, and the sulfate transporter SULTR2;1, leading to posttranscriptional degradation of target gene transcripts (84, 90). An important aspect of miR395-mediated regulation is its involvement in the SLIM1 regulatory circuit (90) (Figure 4). When the supply of sulfur is limited, SLIM1 induces miR395 accumulation in both shoots and roots. This explains the suppression of SULTR2;1 in shoots, which may contribute to limiting the distribution of sulfur from older to younger leaves (122). miR395 is expressed in phloem companion cells, suggesting the actual sites for the degradation of targets are relatively restricted (90). Thus, it should be noted that regulation of SULTR2;1 cannot always be explained by miR395 accumulation. Unexpected 168

Takahashi et al.

as being the degradation target of miR395, SULTR2;1 mRNA accumulates in roots under sulfur-limited conditions (88, 189). Thus it seems that more sulfate enters xylem under sulfur-deficient conditions as a result of the nonoverlapping patterns of expression between miR395 and SULTR2;1 (90), although an additional regulatory pathway for SULTR2;1 induction needs to be considered. In addition to these mechanisms, ectopic overexpression of miR395 results in the modulation of the plant sulfur status to induce the expression of SULTR1 and SULTR4 sulfate transporters in roots and the accumulation of sulfate in the old leaves (122). Auxin response factor. The 5 -promoter region of SULTR1;1 shows the presence of an auxin response factor (ARF) binding site for the sulfur response (133). This sulfur-responsive element (SURE) confers transcriptional


PP62CH07-Takahashi

ARI

4 April 2011

14:47

regulation specific for sulfur limitation but not for auxin (133). However, during long-term sulfur starvation, degradation of indole glucosinolates by nitrilase provides a precursor for auxin biosynthesis (116). Transcriptome analysis suggests that sulfur starvation may induce auxin signals (141). However, overexpression of sulfur-responsive auxin signaling components affected numerous metabolic pathways apart from sulfur metabolism (34). This leaves actual linkages between the sulfur response and auxin signals currently unspecified. SURE was present in SULTR1;1 but was absent in SULTR1;2, suggesting diversification of the regulation of these transporters (133). In fact, SULTR1;1 is controlled more specifically by sulfur limitation, whereas SULTR1;2 seems to be regulated by general metabolic demand and cellular status (171), again pointing to a specific role for SURE in the sulfur-limitation response. MYB. Several MYB transcription factors have been identified as positive regulators of glucosinolate biosynthesis (21, 46, 47, 74) (Figure 4). They were discovered by a genetic screen for Arabidopsis dominant mutants resistant to a tryptophan analogue (21), by in silico analysis of transcriptome data of the sulfur response (74), or by a screening of Arabidopsis lines with altered levels of glucosinolate concentrations (46, 47). They belong to the R2R3-type MYB family and were classified into two groups. MYB28, MYB29, and MYB76 induced the expression of biosynthetic enzymes for methionine-derived aliphatic glucosinolates (47, 74), whereas MYB34, MYB51, and MYB122 activated the synthesis of indolic glucosinolates (21, 46). MYB34 and MYB122 alternatively functioned by stimulating auxin biosynthesis, whereas MYB51 was specific to the activation of the glucosinolate pathway (46). MYB34 and its downstream enzymes were negatively controlled by SLIM1 in response to sulfur limitation (132), whereas the effect of SLIM1 on MYB28 and MYB29 regulation was unclear (74), suggesting the presence of both SLIM1-dependent and -independent pathways

for control of glucosinolate biosynthesis (Figure 4). Interestingly, these MYBs can also stimulate expression of primary sulfur assimilation enzymes, enhancing substrate supply for glucosinolate biosynthesis. Although all six MYB factors regulate APR and APS kinase, the trans-activation of ATPS was isoform specific in relation to the aliphatic and indolic group (205).

CSC: cysteine synthase complex

Cysteine Synthase Complex Plant and bacterial cysteine synthase complexes (CSC) share many structural and kinetic properties, but they appear to have quite different functions with respect to regulation. In E. coli tight and efficient control of the cysteine regulon is based on the constitutive expression of the cysE gene that encodes SAT and feedback inhibition of CysE by cysteine (110). Spontaneous conversion of OAS results in Nacetylserine that acts as an inducer of the CysB DNA binding protein to control the other genes of the regulon including cysK (encoding OAS-TL) (110). In plants SAT and OAS-TL are considered as components of metabolic control systems in three aspects. CSC operates cysteine biosynthesis in cytosol, plastids, and mitochondria, where the stability of the complex is maintained by sulfide to have the SATs in the CSC remain active as long as sulfate availability is not limiting (31, 200). During sulfur deprivation the level of sulfide declines and OAS accumulates, and the latter triggers dissociation of the CSC in conjunction with a reduction of SAT activity. Besides control of cysteine synthesis by protein-protein interaction in the CSC, the pathway intermediate, OAS, contributes to transcriptional regulations of sulfate uptake and reduction (72, 109, 186) (Figure 4). Furthermore, a cytosolic OAS-TL interacts with a cytosolic STAS domain of the sulfate transporter when heterologously expressed in yeast (181), although it is unclear how this could have a regulatory function in view of the high OAS-TL abundance. The plant CSC functions as a determinant of cysteine biosynthesis, sensing the availability of sulfide. This model is based on the reversible


169

ARI

4 April 2011

14:47

protein-protein interaction of SAT and OASTL described above. It also explains in vivo observations: (a) Overexpression of enzymatically inactive SAT in cytosol enhances the total rate of cysteine synthesis (201) more than the overexpression of active SAT does (183, 201); (b) The oastlAB double mutants survive with only the mitochondrial OAS-TL capable of interacting with SAT (66). In both cases the increased availability of sulfide can stabilize the association of CSCs in the remaining subcellular compartments and make them more active for synthesis of OAS. The active state of SAT in the complex refers not so much to an increased reaction velocity but to a reduced feedback sensitivity to cysteine (68, 144). A SAT from soybean was shown to become much less feedback sensitive upon phosphorylation by a calciumdependent protein kinase (124). Although they lack the phosphorylation sites, another soybean SAT as well as Arabidopsis cytosolic and mitochondrial SAT became less feedback sensitive to cysteine inside the CSC (115). The unchanged feedback sensitivity of E. coli SAT inside or outside the CSC may be the decisive difference compared with the plant CSC.


PP62CH07-Takahashi

Posttranscriptional Regulation of Methionine Biosynthesis Methionine biosynthesis is tightly controlled by posttranscriptional regulation of its key enzyme, cystathionine γ-synthase (CGS) (22). The mechanism requires SAM for its control and involves destabilization of CGS1 mRNA at the MTO1 region where mutations for methionine overaccumulating phenotypes have been frequently found in Arabidopsis mto1 mutants (23). In the presence of SAM, translation of CGS1 can be arrested at the downstream vicinity of the MTO1 region, which allows CGS1 mRNA to be rapidly degraded to control methionine biosynthesis (152). In addition, alternative CGS1 transcripts having 90 or 87 nucleotide deletions downstream of the site of translation arrest were stable for overproduction of methionine (59). These lines of evidence indicate the significance of posttranscriptional 170

Takahashi et al.

regulation of CGS1 in methionine biosynthesis. However, generalizing the mechanism remains controversial as a CGS1 mRNA of potato has been shown to be stable even in the presence of methionine (111).

Systems Regulation The systems regulation is best elucidated by consolidating multi-omics data sets and computing mathematical models (176). In the past several years, a number of efforts have been made to integrate multi-omics directed at a holistic understanding of sulfur metabolism in Arabidopsis plants. The subsequent findings unraveled the systems networks of molecular components—genes, transcripts, proteins, and metabolites—that respond to environmental stimuli including changes in sulfur availability. Transcriptomic studies indicated that, apart from sulfate uptake and assimilation, a variety of metabolic pathways are modulated by sulfate depletion (72, 131, 141). Secondary metabolism (sulfur and flavonoid), nitrogen metabolism, oxidative stress, and hormone synthesis (auxin and jasmonate) were responsive to a depletion of the sulfate supply, and those responses were generally mimicked by adding OAS (72) or knocking out a sulfate transporter gene, SULTR1;2 (131). Integration of metabolomics and transcriptomics in sulfur-starved Arabidopsis plants further indicated a general trend of systematic reconfiguration of certain biological processes, such as enhanced photorespiration, nitrogen imbalance, glucosinolate synthesis and catabolism, flavonoid biosynthesis, lipid breakdown, as well as auxin and jasmonate metabolism (73, 74, 75, 142, 143). Predictions of the functions of sulfotransferases (AtSOT16, 17 and 18) (73) involved in the last step of glucosinolate synthesis and of the two R2R3 MYB transcription factors (MYB28 and MYB29) (74) that control the synthesis of Met-derived glucosinolates was successfully made through this integrated multi-omics approach. More broadly, part of the systems responses to sulfur depletion included those relevant to


PP62CH07-Takahashi

ARI

4 April 2011

14:47

senescence or stress that may occur under general nutrient deficiency (196). These responses were suggested by a close similarity of the transcriptomic and metabolomic responses between the serat (serine acetyltransferase, SAT) quadruple mutants displaying sulfur deficiency and plants grown under nitrogen-, phosphorus-, potassium-, and sulfur-depleted conditions (196). The transcriptome coexpression networks analysis based on the “guilt-by-association” principle is excellent for delimiting candidate genes involved in certain coincidental biological processes, in particular, when applied to a large-scale microarray data set across more than 1000 different experiments (174). Via this approach, the involvement of MYB28 and MYB29 in controlling glucosinolate synthesis was further confirmed (74), and the gene coding for a flavin-monooxygenase responsible for S-oxygenation in aliphatic glucosinolate biosynthesis was predicted (62). Similarly a plastidic UDP-glucose pyrophosphorylase committed to the first step of sulfolipid biosynthesis was successfully delimited from an orphan gene family (151). Genomic analysis of transcripts as phenotypes led to the expression quantitative trait loci (eQTL) analysis of the natural intraspecific genetic variation in Arabidopsis (97). An eQTL investigation together with glucosinolate profiling and metabolome analysis of Arabidopsis Bay × Sha recombinant inbred lines (RIL) identified AOP2, the last enzyme in the biosynthetic pathway, as a major regulator of both transcript levels for the entire pathway and aliphatic glucosinolate accumulation (172, 199). This suggests that natural variation in metabolites or their synthesis rates can feedback control the transcripts. The same RIL population was used to show that a single nucleotide polymorphism in an APR2 isoform of APR, resulting in strong differences in catalytic efficiency of the enzyme, is a major determinant for sulfate accumulation in plants (125). Interestingly, this QTL showed a strong interaction with nitrogen availability, thus confirming the coordination of the two assimilatory pathways (125).

Sulfur Acclimation Signals in Chlamydomonas The green alga Chlamydomonas reinhardtii provides an interesting model of the molecular mechanisms of the sulfur-limitation response (Figure 4). Unlike higher plants, Chlamydomonas is capable of hydrolyzing sulfate esters by using arylsulfatase to obtain sulfur under sulfate-deplete conditions. Regulatory components controlling the sulfur-limitation response were identified from a genetic screen for sac (sulfur acclimation) mutants showing abnormalities in arylsulfatase expression (28, 29). A membrane-bound putative sulfur-sensor protein, SAC1, was a causal gene initially identified from the screening (28). SAC1 induces the expression of proteins associated with the acclimation response to sulfur starvation and reduces photosynthesis that needs to be downregulated synchronously with decreased capacity to assimilate sulfur (28). Regulation under SAC1 is significant for Chlamydomonas cells to acclimate to a low-sulfur environment, although high-affinity sulfate transporters remained active in sac1 mutants. More recent analyses of SAC1-like transporters, SLT1 and SLT2, indicate these SAC1 homologues are required for high-affinity sulfate transport activities as sodium/sulfate cotransporters regulated under the components of sulfur-sensing machineries (156). Besides SAC1, SNRK2 family Ser/Thr protein kinases, SNRK2.1 and SNRK2.2 (SAC3), are involved in signaling pathways that control sulfur-deprivation responses in Chlamydomonas (29, 52). SNRK2.1 controls the acclimation response when cells are deprived of sulfur. Transcriptome analysis indicates SNRK2.1 is a direct upstream regulatory component of sulfur-responsive genes (51). When sulfur supply is sufficient, SNRK2.2 controls SNRK2.1 and represses the acclimation response. In contrast, SAC1 negatively controls SNRK2.2 under sulfur-deprived conditions, which allows for SNRK2.1 function (51, 52) (Figure 4). Genes controlled downstream of SNRK2.1 are related to sulfur acquisition, sulfur metabolism, substitution of sulfolipids and sulfur-containing


171

PP62CH07-Takahashi

ARI

4 April 2011

14:47

cell wall constituents, balancing of carbon and nitrogen metabolisms, and regulation of photosynthesis (51). When cells acclimate to a low-sulfur environment, these metabolic processes are necessarily controlled to maintain viability. The downstream transcriptional networks require further investigation.


IMPACT OF SULFUR AVAILABILITY ON THE EVOLUTION OF PHOTOSYNTHETIC ORGANISMS The basal effects of sulfur in photolitotrophic evolution can be found in aquatic primary producers, because the phylogenetic diversification of photosynthetic organisms occurred mostly in the oceans (49, 145). Sulfur availability and chemical form are among the features of oceanic chemistry that underwent the greatest changes. There is a striking temporal coincidence between, on the one hand, the waning of widespread euxenia and removal of sulfide in the euphotic zone (together with a return to anoxic and Fe2+ -rich subsurface waters) with, on the other hand, eukaryote diversification and the establishment of the dominance of oxygenic photosynthesis (20, 82). Before these Neoproterozoic events, only cyanobacteria were capable of appreciable oxygenic photosynthesis, although, when sulfide was abundant, they probably largely substituted H2 O with hydrogen sulfide as an electron source (25, 85). The appearance of cyanobacteria presumably occurred in the Archean ocean, whose sulfate content was below 200 μM (58). For most of the Proterozoic, ocean sulfate concentration was ∼1–5 mM (19, 87). During that time, green algae acquired ecological relevance (82, 98), but not until the Ediacaran did sulfate concentrations reach ∼15 mM. The trend toward increasing sulfate concentration halted in the Cambrian when sulfate concentration was 3– 12 mM (78). It stayed below 10 mM and possibly closer to the lower end of this range until the Carboniferous when it increased to >15 mM (48). The 28-mM sulfate concentration of extant oceans is likely the historical maximum. 172

Takahashi et al.

Interestingly, biomarker and molecularclock data indicate that the most abundant primary producers in extant oceans [i.e., diatoms, coccolithophorids, photosynthetic dinoflagellates, subsequently referred to as Chl (chlrorophyll) a+c algae] rose to dominance from the late Paleozoic through the Mesozoic (earlier evidence for dinoflagellates in the Silurian; no evidence of prominent diatoms and coccolithophorids until the Mesozoic) (36, 98), concomitant with the rise of sulfate concentrations to >15 mM (48). Experimental results show that extant Chl a+c algae indeed have higher growth rates with increasing sulfate, throughout the range of estimated values from the Proterozoic to the Mesozoic oceans, whereas prasinophyte green algae do not show a similar response. When Chl a+c phytoplankters and prasinophytes are inoculated together in media mimicking oceans at different points in the Earth’s history, the green algae outcompete the others in the Paleozoic treatment, but they are essentially eliminated in conditions resembling the Mesozoic and current oceans. These results are consistent with the hypothesis that the rise of sulfate concentration was not contradictory to the shift of algal group dominance in the oceans. The comparison with freshwaters reinforces this idea: In freshwaters, where sulfate concentrations are one to two orders of magnitude lower than in the sea (0.01 to 1 mM), green algae constitute a much greater proportion of primary producers than in the oceans (50, 76). Chl a+c algae have lower C:S ratios than green algae and cyanobacteria (145). It has been proposed that the success of these algal taxa is related to the possibility afforded by the greater S cell quota to produce larger amounts of DMSP (49, 145). A relationship between the cell C:S ratio and the ability to produce DMSP appears to exist, at least for phytoplankton and with the exception of some diatoms (91, 160). Terrestrial plants have mostly lost the ability to produce DMSP; only some angiosperms belonging to the unrelated genera Saccharum, Spartina and Wollastonia reacquired it, but with biosynthetic pathways that are different from those of algae (167). DMSP is widespread and


PP62CH07-Takahashi

ARI

4 April 2011

14:47

possibly highly conserved among algae, suggesting that its appearance precedes the expansion of Chl a+c algae and the concomitant increase in sulfur availability. It is an excellent alternative osmolyte to its N-containing analogue glycine betaine (18, 92, 167). The ability to produce DMSP may have thus represented an adaptive advantage in N-limited oceans (35). Regardless of its original role, DMSP later acquired a number of functions (160). Among these, the antigrazing activity that DMSP mainly exerts through its degradation to acrylate may have been especially important in shaping the destiny of photosynthetic lineages (145, 202, 203, 206). When oceanic sulfur availability increased above basic growth requirements, the algae capable of accumulating sulfur and allocating it to DMSP acquired a potent weapon against grazers, whereas this anti-grazing ability was absent from their low C:S competitors. This may have become especially useful at the beginning of the Mesozoic, when an increase in copepod grazing pressure may have favored the modern phytoplankton biota in a sort of microscopic version of Vermeij’s Mesozoic marine revolution (194). Although not necessarily a strict alternative to the influences of other factors and events, a potential physiological link is suggested between the Phanerozoic increase in marine sulfate concentrations, the evolutionary expansion of low C:S Chl a+c algae, and the confinement of most green algae and their descendents to low-sulfate environments such as freshwaters and the emerged lands.

CONCLUSIONS The past decade brought a quantum leap in our understanding of the biochemistry and physiology of plant sulfate acquisition and metabolism, especially their organ and compartment specificities, responsiveness to environmental and intrinsic signals, and gene-to-metabolite relationships of the processes. However, the molecular mechanisms associated with pathway regulation as well as sulfur sensing and signaling are still far from being fully understood. Recent findings of key regulatory components, SLIM1, the MYBs, and miR395, from Arabidopsis are important milestones allowing us to build up the first concepts regarding the regulatory networks (Figure 4). Metabolite-dependent control mechanisms for CSC organization and posttranscriptional degradation of the CGS1 transcript represent additional regulation modes for cysteine and methionine biosynthesis and reveal the great variety of concurrent mechanisms to fine-tune the regulation of the pathway. However, it should be noted that both upstream and downstream components of the regulatory networks of the known components are not yet fully characterized, suggesting that we are still at the beginning of the road to understanding the entire control mechanism of sulfur metabolism. In-depth omics analysis together with an application of natural variation and new genetic screens are necessary to uncover additional components of these networks and to elucidate their interactions.

FUTURE ISSUES 1. Sensing machineries that recognize fluctuations of the cellular sulfur status are not well understood. CSC organization is under the control of its substrates, and posttranscriptional regulation of CGS1 is dependent on SAM. It is necessary to resolve how these metabolite-sensing mechanisms or other independent sensing machineries may control sulfate transport and assimilation especially in response to the availability of external sulfate. SAC1 and SNRK2s are key sulfur-sensing and -signaling components in Chlamydomonas. Orthologous systems or alternative mechanisms for sulfur sensing are still unknown in higher plants.


173

PP62CH07-Takahashi

ARI

4 April 2011

14:47

2. Findings of the first transcriptional regulators opened a way to understand how the primary sulfur assimilatory pathway and glucosinolate biosynthesis are regulated in Arabidopsis. Currently these essential regulatory components are still uncoupled with upstream signaling events and the sulfur-sensing pathways of Chlamydomonas remain disconnected from the transcriptional networks. Additional transcription factors regulating gene expression of the components of sulfur metabolism await discovery.


3. Omics analysis indicated that the response to sulfur availability integrates a wide spectrum of biological events in both metabolic and developmental contexts. Numerous networks can be drawn for the complex interactions between genes, proteins, and metabolites that integrate sulfur metabolism with a general plant life cycle. Detailed dissection of these network models may provide unexpected findings, further pointing to a biological significance and indispensability of sulfur in nature. 4. The very recent attempt to investigate experimentally the role of sulfur on the evolution of photosynthetic organisms also provides new perspectives to enable an understanding of the interactions of sulfur acquisition and assimilation with the environment and its perturbations.

DISCLOSURE STATEMENT The authors are not aware of any affiliations, memberships, funding, or financial holdings that might be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS We thank all our colleagues who delivered significant amounts of scientific input to this research area. We are also grateful to the following granting organizations for financial support: Ministry of Education, Culture, Sports, Science, and Technology of Japan; Japan Society for the Promotion of Science; Japan Science and Technology Agency; and Bio-oriented Technology Research Advancement Institution (H.T. and K.S.); British Biotechnology and Biological Sciences Research Council (S.K.); Deutsche Forschungsgemeinschaft and CellNetworks Cluster of the German Excellence Initiative (R.H.); as well as Fondazione Cariverona and Università Politecnica delle Marche Internal funding, Italy (M.G.). We also gratefully acknowledge Andrew H. Knoll, Harvard University, and John A. Raven, University of Dundee, for their input regarding the S-related evolutionary issues.

LITERATURE CITED 1. Alvarez C, Calo L, Romero LC, Garcia I, Gotor C. 2010. An O-acetylserine(thiol)lyase homolog with L-cysteine desulfhydrase activity regulates cysteine homeostasis in Arabidopsis. Plant Physiol. 152:656– 69 2. Amano Y, Tsubouchi H, Shinohara H, Ogawa M, Matsubayashi Y. 2007. Tyrosine-sulfated glycopeptide involved in cellular proliferation and expansion in Arabidopsis. Proc. Natl. Acad. Sci. USA 104:18333– 38 3. Amir R, Hacham Y, Galili G. 2002. Cystathionine γ-synthase and threonine synthase operate in concert to regulate carbon flow towards methionine in plants. Trends Plant Sci. 7:153–56 174

Takahashi et al.


PP62CH07-Takahashi

ARI

4 April 2011

14:47

4. Awazuhara M, Fujiwara T, Hayashi H, Watanabe-Takahashi A, Takahashi H, et al. 2005. The function of SULTR2;1 sulfate transporter during seed development in Arabidopsis thaliana. Physiol. Plant 125:95– 105 5. Barberon M, Berthomieu P, Clairotte M, Shibagaki N, Davidian JC, et al. 2008. Unequal functional redundancy between the two Arabidopsis thaliana high-affinity sulphate transporters SULTR1;1 and SULTR1;2. New Phytol. 180:608–19 6. Barroso C, Romero LC, Cejudo FJ, Vega JM, Gotor C. 1999. Salt-specific regulation of the cytosolic O-acetylserine(thiol)lyase gene from Arabidopsis thaliana is dependent on abscisic acid. Plant Mol. Biol. 40:729–36 7. Bashandy T, Guilleminot J, Vernoux T, Caparros-Ruiz D, Ljung K, et al. 2010. Interplay between the NADP-linked thioredoxin and glutathione systems in Arabidopsis auxin signaling. Plant Cell 22:376–91 8. Baxter I, Muthukumar B, Park HC, Buchner P, Lahner B, et al. 2008. Variation in molybdenum content across broadly distributed populations of Arabidopsis thaliana is controlled by a mitochondrial molybdenum transporter (MOT1). PLoS Genet. 4:e1000004 9. Bednarek P, Pilewska-Bednarek M, Svato A, Schneider B, Doubsk J, et al. 2009. A glucosinolate metabolism pathway in living plant cells mediates broad-spectrum antifungal defense. Science 323:101– 6 10. Benning C. 1998. Biosynthesis and function of the sulfolipid sulfoquinovosyl diacylglycerol. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49:53–75 11. Bermudes MA, Páez-Ochoa MA, Gotor C, Romero LC. 2010. Arabidopsis S-sulfocysteine synthase ac´ tivity is essential for chloroplast function and long-day light-dependent redox control. Plant Cell 22:403– 16 12. Bianchini F, Vainio H. 2001. Allium vegetables and organosulfur compounds: Do they help prevent cancer? Environ. Health Perspect. 109:893–902 ˚ 13. Bick JA, Aslund F, Chen Y, Leustek T. 1998. Glutaredoxin function for the carboxyl-terminal domain of the plant-type 5 -adenylylsulfate reductase. Proc. Natl. Acad. Sci. USA 95:8404–9 14. Bick JA, Setterdahl AT, Knaff DB, Chen Y, Pitcher LH, et al. 2001. Regulation of the plant-type 5 -adenylyl sulfate reductase by oxidative stress. Biochemistry 40:9040–48 15. Bourgis F, Roje S, Nuccio ML, Fisher DB, Tarczynski MC, et al. 1999. S-methylmethionine plays a major role in phloem sulfur transport and is synthesized by a novel type of methyltransferase. Plant Cell 11:1485–98 16. Brunold C, Schiff JA. 1976. Studies of sulfate utilization by algae: 15. Enzymes of assimilatory sulfate reduction in Euglena and their cellular localization. Plant Physiol. 57:430–36 17. Brychkova G, Xia Z, Yang G, Yesbergenova Z, Zhang Z, et al. 2007. Sulfite oxidase protects plants against sulfur dioxide toxicity. Plant J. 50:696–709 18. Bucciarelli E, Sunda WG. 2003. Influence of CO2 , nitrate, phosphate, and silicate limitation on intracellular dimethylsulfoniopropionate in batch cultures of the coastal diatom Thalassiosira pseudonana. Limnol. Oceanogr. 48:2256–65 19. Canfield DE. 2004. The evolution of the Earth surface sulfur reservoir. Am. J. Sci. 304:839–61 20. Canfield DE, Poulton SW, Knoll AH, Narbonne GM, Ross G, et al. 2008. Ferruginous conditions dominated later Neoproterozoic deep water chemistry. Science 321:949–52 21. Celenza JL, Quiel JA, Smolen GA, Merrikh H, Silvestro AR, et al. 2005. The Arabidopsis ATR1 Myb transcription factor controls indolic glucosinolate homeostasis. Plant Physiol. 137:253–62 22. Chiba Y, Ishikawa M, Kijima F, Tyson RH, Kim J, et al. 1999 Evidence for autoregulation of cystathionine γ-synthase mRNA stability in Arabidopsis. Science 286:1371–74 23. Chiba Y, Sakurai R, Yoshino M, Ominato K, Ishikawa M, et al. 2003. S-adenosyl-L-methionine is an effector in the posttranscriptional autoregulation of the cystathionine γ-synthase gene in Arabidopsis. Proc. Natl. Acad. Sci. USA 100:10225–30 24. Clarkson DT, Smith FW, Vanden Berg PJ. 1983. Regulation of sulphate transport in a tropical legume, Macroptilium atropurpureum cv. Siratro. J. Exp. Bot. 34:1463–83 25. Cohen Y, Jorgensen BB, Revsbech NP, Poplawski R. 1986. Adaptation to hydrogen sulfide of oxygenic and anoxygenic photosynthesis among cyanobacteria. Appl. Environ. Microbiol. 51:398–407 www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

175

PP62CH07-Takahashi

ARI

4 April 2011

14:47

26. Cook PF, Wedding RT. 1978. Cysteine synthetase from Salmonella typhimurium LT-2. Aggregation, kinetic behavior, and effect of modifiers. J. Biol. Chem. 253:7874–79 27. Curien G, Dumas R, Ravanel S, Douce R. 1996. Characterization of an Arabidopsis thaliana cDNA encoding an S-adenosylmethionine-sensitive threonine synthase. Threonine synthase from higher plants. FEBS Lett. 390:85–90


28. The first identification of a putative sulfur sensor essential for acclimation of Chlamydomonas cells to sulfur deprivation.

176

28. Davies JP, Yildiz FH, Grossman A. 1996. Sac1, a putative regulator that is critical for survival of Chlamydomonas reinhardtii during sulfur deprivation. EMBO J. 15:2150–59 29. Davies JP, Yildiz FH, Grossman AR. 1999. Sac3, an Snf1-like serine/threonine kinase that positively and negatively regulates the responses of Chlamydomonas to sulfur limitation. Plant Cell 11:1179–90 30. Dominguez-Solis JR, He Z, Lima A, Ting J, Buchanan BB, et al. 2008. A cyclophilin links redox and light signals to cysteine biosynthesis and stress responses in chloroplasts. Proc. Natl. Acad. Sci. USA 105:16386–91 31. Droux M, Ruffet ML, Douce R, Job D. 1998. Interactions between serine acetyltransferase and Oacetylserine (thiol) lyase in higher plants-structural and kinetic properties of the free and bound enzymes. Eur. J. Biochem. 255:235–45 32. Eichel J, González JC, Hotze M, Matthews RG, Schroder J. 1995. Vitamin-B12 -independent me¨ thionine synthase from a higher plant (Catharanthus roseus). Molecular characterization, regulation, heterologous expression, and enzyme properties. Eur. J. Biochem. 230:1053–58 33. Eilers T, Schwarz G, Brinkmann H, Witt C, Richter T, et al. 2001. Identification and biochemical characterization of Arabidopsis thaliana sulfite oxidase. A new player in plant sulfur metabolism. J. Biol. Chem. 276:46989–94 34. Falkenberg B, Witt I, Zanor MI, Steinhauser D, Mueller-Roeber B, et al. 2008. Transcription factors relevant to auxin signalling coordinate broad-spectrum metabolic shifts including sulphur metabolism. J. Exp. Bot. 59:2831–46 35. Falkowski PG, Barber RT, Smetacek V. 1998. Biogeochemical controls and feedbacks on ocean primary production. Science 281:200–6 36. Falkowski PG, Katz ME, Knoll AH, Quigg A, Raven JA, et al. 2004. The evolution of modern eukaryotic phytoplankton. Science 305:354–60 37. Feldman-Salit A, Wirtz M, Hell R, Wade RC. 2009. A mechanistic model of the cysteine synthase complex. J. Mol. Biol. 386:37–59 38. Ferretti M, Destro T, Tosatto SCE, La Rocca L, Rascio N, et al. 2009. Gamma-glutamyl transferase in the cell wall participates in extracellular glutathione salvage from the root apoplast. New Phytol. 181:115–26 39. Fodor J, Gullner G, Adam AL, Barna B, Komives T, et al. 1997. Local and systemic responses of antioxidants to tobacco mosaic virus infection and to salicylic acid in tobacco (role in systemic acquired resistance). Plant Physiol. 114:1443–51 40. Foyer CH, Noctor G. 2009. Redox regulation in photosynthetic organisms: signaling, acclimation, and practical implications. Antioxid. Redox. Signal. 11:861–905 41. Francois JA, Kumaran S, Jez JM. 2006. Structural basis for interaction of O-acetylserine sulfhydrylase and serine acetyltransferase in the Arabidopsis cysteine synthase complex. Plant Cell 18:3647–55 42. Freeman JL, Garcia D, Kim D, Hopf A, Salt DE. 2005. Constitutively elevated salicylic acid signals glutathione-mediated nickel tolerance in Thlaspi nickel hyperaccumulators. Plant Physiol. 137:1082–91 43. Frottin F, Espagne C, Traverso JA, Mauve C, Valot B, et al. 2009. Cotranslational proteolysis dominates glutathione homeostasis to support proper growth and development. Plant Cell 21:3296–314 44. Gidda SK, Miersch O, Levitin A, Schmidt J, Wasternack C, et al. 2003. Biochemical and molecular characterization of a hydroxyjasmonate sulfotransferase from Arabidopsis thaliana. J. Biol. Chem. 278:17895–900 45. Gidda SK, Varin L. 2006. Biochemical and molecular characterization of flavonoid 7-sulfotransferase from Arabidopsis thaliana. Plant Physiol. Biochem. 44:628–36 46. Gigolashvili T, Berger B, Mock HP, Muller C, Weisshaar B, et al. 2007. The transcription factor ¨ HIG1/MYB51 regulates indolic glucosinolate biosynthesis in Arabidopsis thaliana. Plant J. 50:886–901 Takahashi et al.


PP62CH07-Takahashi

ARI

4 April 2011

14:47

47. Gigolashvili T, Yatusevich R, Berger B, Muller C, Flugge UI. 2007. The R2R3-MYB transcription ¨ ¨ factor HAG1/MYB28 is a regulator of methionine-derived glucosinolate biosynthesis in Arabidopsis thaliana. Plant J. 51:247–61 48. Gill BC, Lyons TW, Saltzman MR. 2007. Parallel, high-resolution carbon and sulfur isotope records of the evolving Paleozoic marine sulfur reservoir. Palaeogeogr. Palaeoclimatol. Palaeoecol. 256:156–73 49. Giordano M, Norici A, Hell R. 2005. Sulfur and phytoplankton: acquisition, metabolism and impact on the environment. New Phytol. 166:371–82 50. Giordano M, Norici A, Ratti S, Raven JA. 2008. Role of sulfur for algae: acquisition, metabolism, ecology and evolution. See Hell et al., pp. 397–415 51. Gonzalez-Ballester D, Casero D, Cokus S, Pellegrini M, Merchant SS, et al. 2010. RNA-seq ´ analysis of sulfur-deprived Chlamydomonas cells reveals aspects of acclimation critical for cell survival. Plant Cell 22:2058–84 52. Gonzalez-Ballester D, Pollock SV, Pootakham W, Grossman AR. 2008. The central role of a SNRK2 kinase in sulfur deprivation responses. Plant Physiol. 147:216–27 53. Gromes R, Hothorn M, Lenherr ED, Rybin V, Scheffzek K, et al. 2008. The redox switch of γglutamylcysteine ligase via a reversible monomer-dimer transition is a mechanism unique to plants. Plant J. 54:1063–75 54. Grzam A, Martin M, Hell R, Meyer A. 2007. γ-Glutamyl transpeptidase GGT4 initiates vacuolar degradation of glutathione S-conjugates in Arabidopsis. FEBS Lett. 581:3131–38 55. Guo H, Ecker JR. 2004. The ethylene signaling pathway: new insights. Curr. Opin. Plant Biol. 7:40–49 56. Gutierrez-Marcos JF, Roberts MA, Campbell EI, Wray JL. 1996. Three members of a novel small genefamily from Arabidopsis thaliana able to complement functionally an Escherichia coli mutant defective in PAPS reductase activity encode proteins with a thioredoxin-like domain and “APS reductase” activity. Proc. Natl. Acad. Sci. USA 93:13377–82 57. Haas FH, Heeg C, Queiroz R, Bauer A, Wirtz M, et al. 2008. Mitochondrial serine acetyltransferase functions as a pacemaker of cysteine synthesis in plant cells. Plant Physiol. 148:1055–67 58. Habicht KS, Gade M, Thamdrup B, Berg P, Canfield DE. 2002. Calibration of sulfate levels in the Archean ocean. Science 298:2372–74 59. Hacham Y, Schuster G, Amir R. 2006. An in vivo internal deletion in the N-terminus region of Arabidopsis cystathionine γ-synthase results in CGS expression that is insensitive to methionine. Plant J. 45:955–67 60. Halkier BA, Gershenzon J. 2006. Biology and biochemistry of glucosinolates. Annu. Rev. Plant Biol. 57:303–33 61. Hänsch R, Lang C, Riebeseel E, Lindigkeit R, Gessler A, et al. 2006. Plant sulfite oxidase as novel producer of H2 O2 : combination of enzyme catalysis with a subsequent non-enzymatic reaction step. J. Biol. Chem. 281:6884–88 62. Hansen BG, Kliebenstein D, Halkier BA. 2007. Identification of a flavin-monooxygenase as the Soxygenating enzyme in aliphatic glucosinolate biosynthesis in Arabidopsis. Plant J. 50:902–10 63. Hanson AD, Rathinasabapathi B, Rivoal J, Burnet M, Dillon MO, et al. 1994. Osmoprotective compounds in the Plumbaginaceae: a natural experiment in metabolic engineering of stress tolerance. Proc. Natl. Acad. Sci. USA 91:306–10 64. Harada E, Kusano T, Sano H. 2000. Differential expression of genes encoding enzymes involved in sulfur assimilation pathways in response to wounding and jasmonate in Arabidopsis thaliana. J. Plant Physiol. 156:272–76 65. Hatzfeld Y, Maruyama A, Schmidt A, Noji M, Ishizawa K, et al. 2000. β-Cyanoalanine synthase is a mitochondrial cysteine cynthase-like protein in spinach and Arabidopsis. Plant Physiol. 123:1163–72 66. Heeg C, Kruse C, Jost R, Gutensohn M, Ruppert T, et al. 2008. Analysis of the Arabidopsis Oacetylserine(thiol)lyase gene family demonstrates compartment-specific differences in the regulation of cysteine synthesis. Plant Cell 20:168–85 67. Hell R, Bergmann L. 1990. γ-Glutamylcysteine synthetase in higher plants: catalytic properties and subcellular localization. Planta 180:603–12 67a. Hell R, Dahl C, Knaff DB, Leustek T, eds. 2008. Sulfur Metabolism in Phototrophic Organisms. Dordrecht, The Neth.: Springer www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

51. Large-scale transcriptomic analysis provides evidence that SNRK2.1 is a key component for sulfur limitation response in Chlamydomonas.

57. This paper describes mitochondrion as the major subcellular compartment synthesizing OAS in cysteine biosynthesis.

177

PP62CH07-Takahashi

ARI

4 April 2011

14:47


68. Hell R, Wirtz M. 2008. Metabolism of cysteine in plants and phototrophic bacteria. See Hell et al. 2008, pp. 59–91 69. Herschbach C, Rennenberg H. 1991. Influence of glutathione (GSH) on sulphate influx, xylem loading and exudation in excised tobacco roots. J. Exp. Bot. 42:1021–29 70. Hesse H, Hoefgen R. 2003. Molecular aspects of methionine biosynthesis. Trends Plant Sci. 8:259–62 71. Hicks LM, Cahoon RE, Bonner ER, Rivard RS, Sheffield J, et al. 2007. Thiol-based regulation of redox-active glutamate-cysteine ligase from Arabidopsis thaliana. Plant Cell 19:2653–61 72. Hirai MY, Fujiwara T, Awazuhara M, Kimura T, Noji M, et al. 2003. Global expression profiling of sulfur-starved Arabidopsis by DNA macroarray reveals the role of O-acetyl-L-serine as a general regulator of gene expression in response to sulfur nutrition. Plant J. 33:651–63 73. Hirai MY, Klein M, Fujikawa Y, Yano M, Goodenowe DB, et al. 2005. Elucidation of gene-to-gene and metabolite-to-gene networks in Arabidopsis by integration of metabolomics and transcriptomics. J. Biol. Chem. 280:25590–95 74. An excellent example of multi-omics approach that finds MYB-family transcription factors for activation of glucosinolate biosynthesis.

178

74. Hirai MY, Sugiyama K, Sawada Y, Tohge T, Obayashi T, et al. 2007. Omics-based identification of Arabidopsis Myb transcription factors regulating aliphatic glucosinolate biosynthesis. Proc. Natl. Acad. Sci. USA 104:6478–83 75. Hirai MY, Yano M, Goodenowe DB, Kanaya S, Kimura T, et al. 2004. Integration of transcriptomics and metabolomics for understanding of global responses to nutritional stresses in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 101:10205–10 76. Holmer M, Storkholm P. 2001. Sulphate reduction and sulphur cycling in lake sediments: a review. Freshwater Biol. 46:431–51 77. Hopkins L, Parmar S, Blaszczyk A, Hesse H, Hoefgen R, et al. 2005. O-acetylserine and the regulation of expression of genes encoding components for sulfate uptake and assimilation in potato. Plant Physiol. 138:433–40 78. Horita J, Zimmermann H, Holland HD. 2002. Chemical evolution of seawater during the Phanerozoic: Implications from the record of marine evaporites. Geochim. Cosmochim. Acta 66:3733–56 79. Hothorn M, Wachter A, Gromes R, Stuwe T, Rausch T, et al. 2006. Structural basis for the redox control of plant glutamate cysteine ligase. J. Biol. Chem. 281:27557–65 80. Huang B, Vetting MW, Roderick SL. 2005. The active site of O-acetylserine sulfhydrylase is the anchor point for bienzyme complex formation with serine acetyltransferase. J. Bacteriol. 187:3201–5 81. Jez JM, Cahoon RE, Chen S. 2004. Arabidopsis thaliana glutamate-cysteine ligase: functional properties, kinetic mechanism, and regulation of activity. J. Biol. Chem. 279:33463–70 82. Johnston, DT, Wolfe-Simon F, Pearson A, Knoll AH. 2009. Anoxygenic photosynthesis modulated Proterozoic oxygen and sustained Earth’s middle age. Proc. Natl. Acad. Sci. USA 106:16925–29 83. Jones MG, Hughes J, Tregova A, Milne J, Tomsett AB, et al. 2004. Biosynthesis of the flavour precursors of onion and garlic. J. Exp. Bot. 55:1903–18 84. Jones-Rhoades MW, Bartel DP. 2004. Computational identification of plant microRNAs and their targets, including a stress-induced miRNA. Mol. Cell 14:787–99 85. Jorgensen BB, Cohen Y, Revsbech NP. 1986. Transition from anoxygenic to oxygenic photosynthesis in a Microcoleus chtonoplastes cyanobacterial mat. Appl. Environ. Microbiol. 51:408–17 86. Jost R, Altschmied L, Bloem E, Bogs J, Gershenzon J, et al. 2005. Expression profiling of metabolic genes in response to methyl jasmonate reveals regulation of genes of primary and secondary sulfur-related pathways in Arabidopsis thaliana. Photosynth. Res. 36:491–508 87. Kah LC, Lyons TW, Frank TD. 2004. Low marine sulphate and protracted oxygenation of the Proterozoic biosphere. Nature 431:834–38 88. Kataoka T, Hayashi N, Yamaya T, Takahashi H. 2004. Root-to-shoot transport of sulfate in Arabidopsis: evidence for the role of SULTR3;5 as a component of low-affinity sulfate transport system in the root vasculature. Plant Physiol. 136:4198–204 89. Kataoka T, Watanabe-Takahashi A, Hayashi N, Ohnishi M, Mimura T, et al. 2004. Vacuolar sulfate transporters are essential determinants controlling internal distribution of sulfate in Arabidopsis. Plant Cell 16:2693–704 Takahashi et al.


PP62CH07-Takahashi

ARI

4 April 2011

14:47

90. Kawashima CG, Yoshimoto N, Maruyama-Nakashita A, Tsuchiya YN, Saito K, et al. 2009. Sulphur starvation induces the expression of microRNA-395 and one of its target genes but in different cell types. Plant J. 57:313–21 91. Keller MD, Bellows WK, Guillard RRL. 1989. Dimethyl sulfide production in marine phytoplantkon. In Biogenic Sulfur in the Environment, ed. ES Saltzman, WJ Cooper, pp. 167–82. Washington, DC: Am. Chem. Soc. 92. Keller MD, Kiene RP, Matrai PA, Bellows WK. 1999. Production of glycine betaine and dimethylsulfoniopropionate in marine phytoplankton. I. Batch cultures. Mar. Biol. 135:237–48 93. Khan MS, Haas FH, Samami AA, Gholami AM, Bauer A, et al. 2010. Sulfite reductase defines a newly discovered bottleneck for assimilatory sulfate reduction and is essential for growth and development in Arabidopsis thaliana. Plant Cell 22:1216–31 94. Kim SK, Rahman A, Conover RC, Johnson MK, Mason JT, et al. 2006. Properties of the cysteine residues and the iron-sulfur cluster of the assimilatory 5 -adenylyl sulfate reductase from Enteromorpha intestinalis. Biochemistry 45:5010–18 95. Klapheck S, Latus C, Bergmann L. 1987. Localization of glutathione synthetase and distribution of glutathione in leaf cells of Pisum sativum L. J. Plant Physiol. 131:123–31 96. Klein M, Papenbrock J. 2004. The multi-protein family of Arabidopsis sulphotransferases and their relatives in other plant species. J. Exp. Bot. 55:1809–20 97. Kliebenstein D. 2009. Quantitative genomics: analyzing intraspecific variation using global gene expression polymorphisms or eQTLs. Annu. Rev. Plant Biol. 60:93–114 98. Knoll AH, Summons RE, Waldbauer J, Zumberge J. 2007. The geological succession of primary producers in the oceans. In The Evolution of Primary Producers in the Sea, ed. Falkowski P, Knoll AH, pp. 133–63. Burlington: Elsevier 99. Komori R, Amano Y, Ogawa-Ohnishi M, Matsubayashi Y. 2009. Identification of tyrosylprotein sulfotransferase in Arabidopsis. Proc. Natl. Acad. Sci. USA 106:15067–72 100. Kopriva S, Buchert T, Fritz G, Suter M, Weber M, et al. 2001. Plant adenosine 5 -phosphosulfate ¨ reductase is a novel iron-sulfur protein. J. Biol. Chem. 276:42881–86 101. Kopriva S, Buchert T, Fritz G, Suter M, Benda R, et al. 2002. The presence of an iron-sulfur cluster in ¨ adenosine 5 -phosphosulfate reductase separates organisms utilizing adenosine 5 -phosphosulfate and phosphoadenosine 5 -phosphosulfate for sulfate assimilation. J. Biol. Chem. 277:21786–91 102. Kopriva S, Fritzemeier K, Wiedemann G, Reski R. 2007. The putative moss 3 -phosphoadenosine-5 phosphosulfate reductase is a novel form of adenosine-5 -phosphosulfate reductase without an ironsulfur cluster. J. Biol. Chem. 282:22930–38 103. Kopriva S, Koprivova A. 2004. Plant adenosine 5 -phosphosulphate reductase: the past, the present, and the future. J. Exp. Bot. 55:1775–83 104. Kopriva S, Suter M, von Ballmoos P, Hesse H, Krähenbuhl ¨ U, et al. 2002. Interaction of sulfate assimilation with carbon and nitrogen metabolism in Lemna minor. Plant Physiol. 130:1406–13 105. Kopriva S, Wiedemann G, Reski R. 2007. Sulfate assimilation in basal land plants—What does genomic sequencing tell us? Plant Biol. 9:556–64 106. Koprivova A, Meyer A, Schween G, Herschbach C, Reski R, et al. 2002. Functional knockout of the adenosine 5 -phosphosulfate reductase gene in Physcomitrella patens revives an old route of sulfate assimilation. J. Biol. Chem. 277:32195–201 107. Koprivova A, Mugford ST, Kopriva S. 2010. Arabidopsis root growth dependence on glutathione is linked to auxin transport. Plant Cell Rep. 29:1157–67 108. Koprivova A, North KA, Kopriva S. 2008. Complex signaling network in regulation of adenosine 5 phosphosulfate reductase by salt stress in Arabidopsis roots. Plant Physiol. 146:1408–20 109. Koprivova A, Suter M, Op den Camp R, Brunold C, Kopriva S. 2000. Regulation of sulfate assimilation by nitrogen in Arabidopsis. Plant Physiol. 122:737–46 110. Kredich NM. 1996. Biosynthesis of cysteine. Cellular and molecular biology. In Escherichia coli and Salmonella typhimurium, ed. FC Neidhardt, R Curtiss, JL Ingraham, ECC Lin, KB Low, et al., pp. 514– 27. Washington, DC: ASM Press 111. Kreft O, Hoefgen R, Hesse H. 2003. Functional analysis of cystathionine γ-synthase in genetically engineered potato plants. Plant Physiol. 131:1843–54 www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

179


PP62CH07-Takahashi

ARI

4 April 2011

14:47

112. Kruger RJ, Siegel LM. 1982. Evidence for siroheme-Fe4 S4 interaction in spinach ferredoxin-sulfite ¨ reductase. Biochemistry 21:2905–9 113. Kruger S, Niehl A, Martin MCL, Steinhauser D, Donath A, et al. 2009. Analysis of cytosolic and ¨ plastidic serine acetyltransferase mutants and subcellular metabolite distributions suggests interplay of the cellular compartments for cysteine biosynthesis in Arabidopsis. Plant Cell Environ. 32:349–67 114. Krusell L, Krause K, Ott T, Desbrosses G, Krämer U, et al. 2005. The sulfate transporter SST1 is crucial for symbiotic nitrogen fixation in Lotus japonicus root nodules. Plant Cell 17:1625–36 115. Kumaran S, Yi H, Krishnan HB, Jez JM. 2009. Assembly of the cysteine synthase complex and the regulatory role of protein-protein interactions. J. Biol. Chem. 284:10268–75 116. Kutz A, Muller A, Hennig P, Kaiser WM, Piotrowski M, et al. 2002. A role for nitrilase 3 in the ¨ regulation of root morphology in sulphur-starving Arabidopsis thaliana. Plant J. 30:95–106 117. Lappartient AG, Vidmar JJ, Leustek T, Glass ADM, Touraine B. 1999. Inter-organ signaling in plants: regulation of ATP sulfurylase and sulfate transporter genes expression in roots mediated by phloemtranslocated compound. Plant J. 18:89–95 118. Lass B, Ullrich-Eberius CL. 1984. Evidence for proton/sulfate cotransport and its kinetics in Lemna gibba G1. Planta 161:53–60 119. Laudenbach DE, Grossman AR. 1991. Characterization and mutagenesis of sulfur-regulated genes in a cyanobacterium: evidence for function in sulfate transport. J. Bacteriol. 173:2739–50 120. Leggett JE, Epstein E. 1956. Kinetics of sulfate absorption by barley roots. Plant Physiol. 31:222–26 121. Leustek T, Martin MN, Bick JA, Davies JP. 2000. Pathways and regulation of sulfur metabolism revealed through molecular and genetic studies. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51:141–65 122. Liang G, Yang F, Yu D. 2010. MicroRNA395 mediates regulation of sulfate accumulation and allocation in Arabidopsis thaliana. Plant J. 62:1046–57 123. Lindberg P, Melis A. 2008. The chloroplast sulfate transport system in the green alga Chlamydomonas reinhardtii. Planta 228:951–61 124. Liu F, Yoo B-C, Lee J-Y, Pan W, Harmon AC. 2006. Calcium-regulated phosphorylation of soybean serine acetyltransferase in response to oxidative stress. J. Biol. Chem. 281:27405–15 125. QTL analysis shows APS reductase, APR2, is a determinant for sulfate utilization under low nitrogen environment.

132. The first report describing identification of an upstream transcriptional regulator of sulfur limitation response in Arabidopsis.

180

125. Loudet O, Saliba-Colombani V, Camilleri C, Calenge F, Gaudon V, et al. 2007. Natural variation for sulfate content in Arabidopsis thaliana is highly controlled by APR2. Nat. Genet. 39:896–900 126. Lunn JE, Droux M, Martin J, Douce R. 1990. Localization of ATP sulfurylase and Oacetylserine(thiol)lyase in spinach leaves. Plant Physiol. 94:1345–52 127. Marsolais F, Boyd J, Paredes Y, Schinas AM, Garcia M, et al. 2007. Molecular and biochemical characterization of two brassinosteroid sulfotransferases from Arabidopsis, AtST4a (At2g14920) and AtST1 (At2g03760). Planta 225:1233–44 128. Martin MN, Saladores PH, Lambert E, Hudson AO, Leustek T. 2007. Localization of members of the γ-glutamyl transpeptidase family identifies sites of glutathione and glutathione S-conjugate hydrolysis. Plant Physiol. 144:1715–32 129. Martinoia E, Maeshima M, Neuhaus HE. 2007. Vacuolar transporters and their essential role in plant metabolism. J. Exp. Bot. 58:83–102 130. Marty L, Siala W, Schwarzlander M, Fricker MD, Wirtz M, et al. 2009. The NADPH-dependent thioredoxin system constitutes a functional backup for cytosolic glutathione reductase in Arabidopsis. Proc. Natl. Acad. Sci. USA 106:9109–14 131. Maruyama-Nakashita A, Inoue E, Watanabe-Takahashi A, Yamaya T, Takahashi H. 2003. Transcriptome profiling of sulfur-responsive genes in Arabidopsis reveals global effects of sulfur nutrition on multiple metabolic pathways. Plant Physiol. 132:597–605 132. Maruyama-Nakashita A, Nakamura Y, Tohge T, Saito K, Takahashi H. 2006. Arabidopsis SLIM1 is a central transcriptional regulator of plant sulfur response and metabolism. Plant Cell 18:3235– 51 133. Maruyama-Nakashita A, Nakamura Y, Watanabe-Takahashi A, Inoue E, Yamaya T, et al. 2005. Identification of a novel cis-acting element conferring sulfur deficiency response in Arabidopsis roots. Plant J. 42:305–14 Takahashi et al.


PP62CH07-Takahashi

ARI

4 April 2011

14:47

134. Maruyama-Nakashita A, Nakamura Y, Yamaya T, Takahashi H. 2004. A novel regulatory pathway of sulfate uptake in Arabidopsis roots: implication of CRE1/WOL/AHK4-mediated cytokinin-dependent regulation. Plant J. 38:779–89 135. Matsubayashi Y, Ogawa M, Kihara H, Niwa M, Sakagami Y. 2006. Disruption and overexpression of Arabidopsis phytosulfokine receptor gene affects cellular longevity and potential for growth. Plant Physiol. 142:45–53 136. Matsubayashi Y, Sakagami Y. 1996. Phytosulfokine, sulfated peptides that induce the proliferation of single mesophyll cells of Asparagus officinalis L. Proc. Natl. Acad. Sci. USA 93:7623–27 137. Maughan SC, Pasternak M, Cairns N, Kiddle G, Brach T, et al. 2010. Plant homologs of the Plasmodium falciparum chloroquine-resistance transporter, PfCRT, are required for glutathione homeostasis and stress responses. Proc. Natl. Acad. Sci. USA 107:2331–36 138. Mithen R, Faulkner K, Magrath R, Rose P, Williamson G, et al. 2003. Development of isothiocyanateenriched broccoli, and its enhanced ability to induce phase 2 detoxification enzymes in mammalian cells. Theor. Appl. Genet. 106:727–34 139. Mugford SG, Yoshimoto N, Reichelt M, Wirtz M, Hill L, et al. 2009. Disruption of adenosine-5 phosphosulfate kinase in Arabidopsis reduces levels of sulfated secondary metabolites. Plant Cell 21:910– 27 140. Neuenschwander U, Suter M, Brunold C. 1991. Regulation of sulfate assimilation by light and Oacetyl-L-serine in Lemna minor L. Plant Physiol. 97:253–58 141. Nikiforova V, Freitag J, Kempa S, Adamik M, Hesse H, et al. 2003. Transcriptome analysis of sulfur depletion in Arabidopsis thaliana: interlacing of biosynthetic pathways provides response specificity. Plant J. 33:633–50 142. Nikiforova VJ, Daub CO, Hesse H, Willmitzer L, Hoefgen R. 2005. Integrative gene-metabolite network with implemented causality deciphers informational fluxes of sulphur stress response. J. Exp. Bot. 56:1887–96 143. Nikiforova VJ, Kopka J, Tolstikov V, Fiehn O, Hopkins L, et al. 2005. Systems rebalancing of metabolism in response to sulfur deprivation, as revealed by metabolome analysis of Arabidopsis plants. Plant Physiol. 138:304–18 144. Noji M, Inoue K, Kimura N, Gouda A, Saito K. 1998. Isoform-dependent differences in feedback regulation and subcellular localization of serine acetyltransferase involved in cysteine biosynthesis from Arabidopsis thaliana. J. Biol. Chem. 273:32739–45 145. Norici A, Hell R, Giordano M. 2005. Sulfur and primary production in aquatic environments: an ecological perspective. Photosynth. Res. 86:409–17 146. Ohkama N, Takei K, Sakakibara H, Hayashi H, Yoneyama T, et al. 2002. Regulation of sulfurresponsive gene expression by exogenously applied cytokinins in Arabidopsis thaliana. Plant Cell Physiol. 43:1493–501 147. Ohkama-Ohtsu N, Kasajima I, Fujiwara T, Naito S. 2004. Isolation and characterization of an Arabidopsis mutant that overaccumulates O-Acetyl-L-Ser. Plant Physiol. 136:3209–22 148. Ohkama-Ohtsu N, Oikawa A, Zhao P, Xiang C, Saito K, et al. 2008. A γ-glutamyl transpeptidaseindependent pathway of glutathione catabolism to glutamate via 5-oxoproline in Arabidopsis. Plant Physiol. 148:1603–13 149. Ohkama-Ohtsu N, Radwan S, Peterson A, Zhao P, Badr A, et al. 2007. Characterization of the extracellular gamma-glutamyl transpeptidases, GGT1 and GGT2, in Arabidopsis. Plant J. 49:865–77 150. Ohkama-Ohtsu N, Zhao P, Xiang C, Oliver D. 2007. Glutathione conjugates in the vacuole are degraded by gamma-glutamyl transpeptidase GGT3 in Arabidopsis. Plant J. 49:878–88 151. Okazaki Y, Shimojima M, Sawada Y, Toyooka K, Narisawa T, et al. 2009. A chloroplastic UDPglucose pyrophosphorylase from Arabidopsis is the committed enzyme for the first step of sulfolipid biosynthesis. Plant Cell 21:892–909 152. Onouchi H, Nagami Y, Haraguchi Y, Nakamoto M, Nishimura Y, et al. 2005. Nascent peptidemediated translation elongation arrest coupled with mRNA degradation in the CGS1 gene of Arabidopsis. Genes Dev. 19:1799–810 153. Pasternak M, Lim B, Wirtz M, Hell R, Cobbett CS, et al. 2008. Restricting glutathione biosynthesis to the cytosol is sufficient for normal plant development. Plant J. 53:999–1012 www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

137. An important finding that fills the gap of glutathione biosynthesis in cytosol.

152. An elaborate work describing the mRNA destabilizing mechanism of cystathionine γ-synthase.

181

ARI

4 April 2011

14:47

154. Patron NJ, Durnford DG, Kopriva S. 2008. Sulfate assimilation in eukaryotes: fusions, relocations and lateral transfers. BMC Evol. Biol. 8:39 155. Piotrowski M, Schemenewitz A, Lopukhina A, Muller A, Janowitz T, et al. 2004. Desulfoglucosinolate ¨ sulfotransferases from Arabidopsis thaliana catalyze the final step in the biosynthesis of the glucosinolate core structure. J. Biol. Chem. 279:50717–25 156. Pootakham W, González-Ballester D, Grossman AR. 2010. Identification and regulation of plasma membrane sulfate transporters in Chlamydomonas. Plant Physiol. 153:1653–68 156a. Popper Z, Michel G, Hervé C, Domozych D, Willats WGT, et al. 2011. Evolution and diversity of plant cell walls: from algae to flowering plants. Annu. Rev. Plant Biol. 62:567–90 157. Pye VE, Tingey AP, Robson RL, Moody PC. 2004. The structure and mechanism of serine acetyltransferase from Escherichia coli. J. Biol. Chem. 279:40729–36 158. Queval G, Thominet D, Vanacker H, Miginiac-Maslow M, Gakiere B, et al. 2009. H2 O2 -activated up-regulation of glutathione in Arabidopsis involves induction of genes encoding enzymes involved in cysteine synthesis in the chloroplast. Mol. Plant 2:344–56 159. Rae AL, Smith FW. 2002. Localisation of expression of a high-affinity sulfate transporter in barley roots. Planta 215:565–68 160. Ratti S, Giordano M. 2008. Allocation of sulfur to sulfonium compounds in microalgae. In Sulfur Assimilation and Abiotic Stress in Plants, ed. NA Khan, S Singh, S Umar, pp. 317–33. Heidelberg: Springer 161. Rausch T, Wachter A. 2005. Sulfur metabolism: a versatile platform for launching defence operations. Trends Plant Sci. 10:503–9 162. Ravanel S, Block MA, Rippert P, Jabrin S, Curien G, et al. 2004. Methionine metabolism in plants: chloroplasts are autonomous for de novo methionine synthesis and can import S-adenosylmethionine from the cytosol. J. Biol. Chem. 279:22548–57 163. Ravanel S, Gakière B, Job D, Douce R. 1998. Cystathionine γ-synthase from Arabidopsis thaliana: purification and biochemical characterization of the recombinant enzyme overexpressed in Escherichia coli. Biochem J. 331:639–48 164. Ravanel S, Gakière B, Job D, Douce R. 1998. The specific features of methionine biosynthesis and metabolism in plants. Proc. Natl. Acad. Sci. USA 95:7805–12 165. Rea P. 2007. Plant ATP-binding cassette transporters. Annu. Rev. Plant Biol. 58:347–75 166. Renosto F, Patel HC, Martin RL, Thomassian C, Zimmerman G, et al. 1993. ATP sulfurylase from higher plants: kinetic and structural characterization of the chloroplast and cytosol enzymes from spinach leaf. Arch. Biochem. Biophys. 307:272–85 167. Rhodes D, Hanson AD. 1993. Quaternary ammonium and tertiary sulfonium compounds in higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 44:357–84 168. Rodr´ıguez VM, Chételat A, Majcherczyk P, Farmer EE. 2010. Chloroplastic phosphoadenosine phosphosulfate metabolism regulates basal levels of the prohormone jasmonic acid in Arabidopsis leaves. Plant Physiol. 152:1335–45 169. Rotte C, Leustek T. 2000. Differential subcellular localization and expression of ATP sulfurylase and 5 -adenylylsulfate reductase during ontogenesis of Arabidopsis leaves indicates that cytosolic and plastid forms of ATP sulfurylase may have specialized functions. Plant Physiol. 124:715–24 170. Rouached H, Berthomieu P, El Kassis E, Cathala N, Catherinot V, et al. 2005. Structural and functional analysis of the C-terminal STAS (sulfate transporter and anti-sigma antagonist) domain of the Arabidopsis thaliana sulfate transporter SULTR1.2. J. Biol. Chem. 280:15976–83 171. Rouached H, Wirtz M, Alary R, Hell R, Arpat AB, et al. 2008. Differential regulation of the expression of two high-affinity sulfate transporters, SULTR1.1 and SULTR1.2, in Arabidopsis. Plant Physiol. 147:897–911 172. Rowe HC, Hansen BG, Halkier BA, Kliebenstein DJ. 2008. Biochemical networks and epistasis shape the Arabidopsis thaliana metabolome. Plant Cell 20:1199–216 173. Saito K. 2004. Sulfur assimilatory metabolism. The long and smelling road. Plant Physiol. 136:2443–50 174. Saito K, Hirai MY, Yonekura-Sakakibara K. 2008. Decoding genes with coexpression networks and metabolomics—‘majority report by precogs.’ Trends Plant Sci. 13:36–43


PP62CH07-Takahashi

182

Takahashi et al.


PP62CH07-Takahashi

ARI

4 April 2011

14:47

175. Saito K, Kurosawa M, Tatsuguchi K, Takagi Y, Murakoshi I. 1994. Modulation of cysteine biosynthesis in chloroplasts of transgenic tobacco overexpressing cysteine synthase [O-acetylserine(thiol)-Iyase]. Plant Physiol. 106:887–95 176. Saito K, Matsuda F. 2010. Metabolomics for functional genomics, systems biology, and biotechnology. Annu. Rev. Plant Biol. 61:463–89 177. Sasaki-Sekimoto Y, Taki N, Obayashi T, Aono M, Matsumoto F, et al. 2005. Coordinated activation of metabolic pathways for antioxidants and defence compounds by jasmonates and their roles in stress tolerance in Arabidopsis. Plant J. 44:653–68 178. Scheerer U, Haensch R, Mendel RR, Kopriva S, Rennenberg H, et al. 2010. Sulphur flux through the 1 sulphate assimilation pathway is differently controlled by adenosine 5 -phosphosulphate reductase under stress and in transgenic poplar plants overexpressing γ-ECS, SO or APR. J. Exp. Bot. 61:609–22 179. Setya A, Murillo M, Leustek T. 1996. Sulfate reduction in higher plants—molecular evidence for a novel 5 -adenylylsulfate reductase. Proc. Natl. Acad. Sci. USA 93:13383–88 180. Shibagaki N, Grossman AR. 2004. Probing the function of STAS domains of the Arabidopsis sulfate transporters. J. Biol. Chem. 279:30791–99 181. Shibagaki N, Grossman AR. 2010. The binding of cysteine synthase to the STAS domain of sulfate transporter and its regulatory consequences. J. Biol. Chem. 285:25094–102 182. Shibagaki N, Rose A, Mcdermott JP, Fujiwara T, Hayashi H, et al. 2002. Selenate-resistant mutants of Arabidopsis thaliana identify SULTR1;2, a sulfate transporter required for efficient transport of sulfate into roots. Plant J. 29:475–86 183. Sirko A, Blaszczyk A, Liszewska F. 2004. Overproduction of SAT and/or OASTL in transgenic plants: a survey of effects. J. Exp. Bot. 55:1881–88 184. Sirko A, Hryniewicz M, Hulanicka D, Bock ¨ A. 1990. Sulfate and thiosulfate transport in Escherichia coli K-12: nucleotide sequence and expression of the cysTWAM gene cluster. J. Bacteriol. 172:3351–57 185. Smith FW, Ealing PM, Hawkesford MJ, Clarkson DT. 1995. Plant members of a family of sulfate transporters reveal functional subtypes. Proc. Natl. Acad. Sci. USA 92:9373–77 186. Smith FW, Hawkesford MJ, Ealing PM, Clarkson DT, Vanden Berg PJ, et al. 1997. Regulation of expression of a cDNA from barley roots encoding a high affinity sulfate transporter. Plant J. 12:875– 84 187. Smith FW, Hawkesford MJ, Prosser IM, Clarkson DT. 1995. Isolation of a cDNA from Saccharomyces cerevisiae that encodes a high affinity sulphate transporter at the plasma membrane. Mol. Gen. Genet. 247:709–15 188. Takahashi H. 2010. Sulfate transport and assimilation in plants. Int. Rev. Cell Mol. Biol. 281:129–59 189. Takahashi H, Watanabe-Takahashi A, Smith FW, Blake-Kalff M, Hawkesford MJ, et al. 2000. The role of three functional sulfate transporters involved in uptake and translocation of sulfate in Arabidopsis thaliana. Plant J. 23:171–82 190. Takahashi H, Yamazaki M, Sasakura N, Watanabe A, Leustek T, et al. 1997. Regulation of sulfur assimilation in higher plants: a sulfate transporter induced in sulfate starved roots plays a central role in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 94:11102–7 191. Talalay P, Fahey JW. 2001. Phytochemicals from cruciferous plants protect against cancer by modulating carcinogen metabolism. J. Nutr. 131:3027S–33S 192. Tomatsu H, Takano J, Takahashi H, Watanabe-Takahashi A, Shibagaki N, et al. 2007. An Arabidopsis thaliana high-affinity molybdate transporter required for efficient uptake of molybdate from soil. Proc. Natl. Acad. Sci. USA 104:18807–12 193. Vauclare P, Kopriva S, Fell D, Suter M, Sticher L, et al. 2002. Flux control of sulphate assimilation in Arabidopsis thaliana: Adenosine 5 -phosphosulphate reductase is more susceptible to negative control by thiols than ATP sulphurylase. Plant J. 31:729–40 194. Vermeij GJ. 1977. The Mesozoic marine revolution: evidence from snails, predators and grazers. Paleobiology 3:245–58 195. Wachter A, Wolf S, Steininger H, Bogs J, Rausch T. 2005. Differential targeting of GSH1 and GSH2 is achieved by multiple transcription initiation: implications for the compartmentation of glutathione biosynthesis in the Brassicaceae. Plant J. 41:15–30 www.annualreviews.org • Sulfur Assimilation in Photosynthetic Organisms

183

PP62CH07-Takahashi

ARI


198. The serat multiple knockout mutants confirm the essential role of mitochondrial SAT in synthesis of OAS. 199. The enzyme catalyzing the last step of aliphatic glucosinolate synthesis is shown to feedback control gene expression of the entire biosynthetic pathway.

184

4 April 2011

14:47

196. Watanabe M, Hubberten HM, Saito K, Hoefgen R. 2010. General regulatory patterns of plant mineral nutrient depletion as revealed by serat quadruple mutants disturbed in cysteine synthesis. Mol. Plant. 3:438–66 197. Watanabe M, Kusano M, Oikawa A, Fukushima A, Noji M, et al. 2008. physiological roles of the β-substituted alanine synthase gene family in Arabidopsis. Plant Physiol. 146:310–20 198. Watanabe M, Mochida K, Kato T, Tabata S, Yoshimoto N, et al. 2008. Comparative genomics and reverse genetics analysis reveal indispensable functions of the serine acetyltransferase gene family in Arabidopsis. Plant Cell 20:2484–96 199. Wentzell AM, Rowe HC, Hansen BG, Ticconi C, Halkier BA, et al. 2007. Linking metabolic QTLs with network and cis-eQTLs controlling biosynthetic pathways. PLoS Genet. 3:1687–701 199a. Wirtz M, Birke H, Heeg C, Throm C, Hosp F, et al. 2010. Structure and function of the heterooligomeric cysteine synthase complex in plants. J. Biol. Chem. 285:32810–17 200. Wirtz M, Hell R. 2006. Functional analysis of the cysteine synthase protein complex from plants: structural, biochemical and regulatory properties. J. Plant Physiol. 163:273–86 201. Wirtz M, Hell R. 2007. Dominant-negative modification reveals the regulatory function of the multimeric cysteine synthase protein complex in transgenic tobacco. Plant Cell 19:625–39 202. Wolfe GV, Steinke M, Kirst GO. 1997. Grazing-activated chemical defence in a unicellular marine alga. Nature 387:894–97 203. Wolfe GV, Strom SL, Holmes JL, Radzio T, Olson MB. 2002. Dimethylsulfoniopropionare cleavage by marine phytoplankton in response to mechanical, chemical, or dark stress. J. Phycol. 34:948–60 204. Xiang C, Oliver DJ. 1998. Glutathione metabolic genes coordinately respond to heavy metals and jasmonic acid in Arabidopsis. Plant Cell 10:1539–50 205. Yatusevich R, Mugford SG, Matthewman C, Gigolashvili T, Frerigmann H, et al. 2010. Genes of primary sulfate assimilation are part of the glucosinolate biosynthetic network in Arabidopsis thaliana. Plant J. 62:1–11 206. Yoch DC. 2002. Dimethylsulfoniopropionate: its sources, role in the marine food web, and biological degradation to dimethylsulfide. Appl. Environ. Microbiol. 68:5804–15 207. Yoshimoto N, Inoue E, Saito K, Yamaya T, Takahashi H. 2003. Phloem-localizing sulfate transporter, Sultr1;3, mediates re-distribution of sulfur from source to sink organs in Arabidopsis. Plant Physiol. 131:1511–17 208. Yoshimoto N, Inoue E, Watanabe-Takahashi A, Saito K, Takahashi H. 2007. Posttranscriptional regulation of high-affinity sulfate transporters in Arabidopsis by sulfur nutrition. Plant Physiol. 145:378–88 209. Yoshimoto N, Takahashi H, Smith FW, Yamaya T, Saito K. 2002. Two distinct high-affinity sulfate transporters with different inducibilities mediate uptake of sulfate in Arabidopsis roots. Plant J. 29:465–73

Takahashi et al.

PP62-FrontMatter

ARI

15 April 2011

10:42

Annual Review of Plant Biology


Contents

Volume 62, 2011

It Is a Long Way to GM Agriculture Marc Van Montagu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 1 Anion Channels/Transporters in Plants: From Molecular Bases to Regulatory Networks Hélène Barbier-Brygoo, Alexis De Angeli, Sophie Filleur, Jean-Marie Frachisse, Franco Gambale, Sébastien Thomine, and Stefanie Wege p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p25 Connecting the Plastid: Transporters of the Plastid Envelope and Their Role in Linking Plastidial with Cytosolic Metabolism Andreas P.M. Weber and Nicole Linka p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p53 Organization and Regulation of Mitochondrial Respiration in Plants A. Harvey Millar, James Whelan, Kathleen L. Soole, and David A. Day p p p p p p p p p p p p p p p p p79 Folate Biosynthesis, Turnover, and Transport in Plants Andrew D. Hanson and Jesse F. Gregory III p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 105 Plant Nucleotide Sugar Formation, Interconversion, and Salvage by Sugar Recycling Maor Bar-Peled and Malcolm A. O’Neill p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 127 Sulfur Assimilation in Photosynthetic Organisms: Molecular Functions and Regulations of Transporters and Assimilatory Enzymes Hideki Takahashi, Stanislav Kopriva, Mario Giordano, Kazuki Saito, and Rudiger ¨ Hell p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 157 Signaling Network in Sensing Phosphate Availability in Plants Tzyy-Jen Chiou and Shu-I Lin p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 185 Integration of Nitrogen and Potassium Signaling Yi-Fang Tsay, Cheng-Hsun Ho, Hui-Yu Chen, and Shan-Hua Lin p p p p p p p p p p p p p p p p p p p p 207 Roles of Arbuscular Mycorrhizas in Plant Nutrition and Growth: New Paradigms from Cellular to Ecosystem Scales Sally E. Smith and F. Andrew Smith p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 227

v

PP62-FrontMatter

ARI

15 April 2011

10:42

The BioCassava Plus Program: Biofortification of Cassava for Sub-Saharan Africa Richard Sayre, John R. Beeching, Edgar B. Cahoon, Chiedozie Egesi, Claude Fauquet, John Fellman, Martin Fregene, Wilhelm Gruissem, Sally Mallowa, Mark Manary, Bussie Maziya-Dixon, Ada Mbanaso, Daniel P. Schachtman, Dimuth Siritunga, Nigel Taylor, Herve Vanderschuren, and Peng Zhang p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 251 In Vivo Imaging of Ca2+ , pH, and Reactive Oxygen Species Using Fluorescent Probes in Plants Sarah J. Swanson, Won-Gyu Choi, Alexandra Chanoca, and Simon Gilroy p p p p p p p p p p p p 273


The Cullen-RING Ubiquitin-Protein Ligases Zhihua Hua and Richard D. Vierstra p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 299 The Cryptochromes: Blue Light Photoreceptors in Plants and Animals Inês Chaves, Richard Pokorny, Martin Byrdin, Nathalie Hoang, Thorsten Ritz, Klaus Brettel, Lars-Oliver Essen, Gijsbertus T.J. van der Horst, Alfred Batschauer, and Margaret Ahmad p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 335 The Role of Mechanical Forces in Plant Morphogenesis Vincent Mirabet, Pradeep Das, Arezki Boudaoud, and Olivier Hamant p p p p p p p p p p p p p p p p 365 Determination of Symmetric and Asymmetric Division Planes in Plant Cells Carolyn G. Rasmussen, John A. Humphries, and Laurie G. Smith p p p p p p p p p p p p p p p p p p p p p p 387 The Epigenome and Plant Development Guangming He, Axel A. Elling, and Xing Wang Deng p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 411 Genetic Regulation of Sporopollenin Synthesis and Pollen Exine Development Tohru Ariizumi and Kinya Toriyama p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 437 Germline Specification and Function in Plants Frédéric Berger and David Twell p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 461 Sex Chromosomes in Land Plants Ray Ming, Abdelhafid Bendahmane, and Susanne S. Renner p p p p p p p p p p p p p p p p p p p p p p p p p p p p 485 Evolution of Photosynthesis Martin F. Hohmann-Marriott and Robert E. Blankenship p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 515 Convergent Evolution in Plant Specialized Metabolism Eran Pichersky and Efraim Lewinsohn p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 549 Evolution and Diversity of Plant Cell Walls: From Algae to Flowering Plants Zoë Popper, Gurvan Michel, Cécile Hervé, David S. Domozych, William G.T. Willats, Maria G. Tuohy, Bernard Kloareg, and Dagmar B. Stengel p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 567

vi

Contents