Sulfur Metabolism

Hideki Takahashi

4.1 Introduction

Sulfur is an essential element and classified as one of the six macroelements that significantly affect plant growth (Marschner, 1995). Sulfur is found in both inorganic and organic compounds in nature. The sulfur cycle is globally balanced between assimilatory and dissimilatory functions of plants and microorganisms (Crawford etal., 2000;Leustek etal., 2000; Saito 2000, 2004; Kopriva, 2006). Plants and microorganisms assimilate inorganic sulfur to produce organic sulfur, and animals will utilise the organic forms of sulfur as nutrient sources. Plants normally utilise sulfate (SO42-), the most oxidised form of sulfur (+VI redox state), to synthesize organic sulfur-compounds (Leustek et al., 2000; Saito, 2000, 2004). Inorganic sulfur-compounds in reduced states such as hydrogen sulfide (H2S) and sulfur dioxide (SO2) are erupted from volcanoes and hot springs, although they are not the sulfur sources primarily utilised by higher plant species. Organic sulfur is present in soil as sulfated esters and sulfonates. Microorganisms in soil degrade these organic sulfur-compounds and release sulfate. In addition, wastes and remains from plants and animals enter the degradation pathways in soil microbes. Catabolic degradation of organic sulfur to inorganic sulfate is an essential step for recycling sulfur source for plants.

Sulfur is present in both major cellular constituents and specialised compounds. Cysteine and methionine are the sulfur-containing essential amino acids. For folding of proteins, the thiol residue of cysteine serves as a moiety to form S-S bonds. It also serves as active sites for electron transfer in enzyme reactions. Glutathione, ferredoxins and thioredoxins are peptides and proteins which contain cysteine to function in redox regulation (Buchanan and Balmer, 2005). Sulfur can be also found in sulfolipid which is an essential component

Plant Metabolism and Biotechnology, First Edition. Edited by Hiroshi Ashihara, Alan Crozier, and Atsushi Komamine. © 2011 John Wiley & Sons, Ltd. Published 2011 by John Wiley & Sons, Ltd. ISBN: 978-0-470-74703-2

of chloroplast membranes (Benning, 1998). Other than being cellular constituents like amino acids, proteins and lipids, sulfur is present in secondary sulfur-metabolites having specific biological functions. Vitamins and cofactors such as thiamine, biotin and coenzyme A contain sulfur. Some secondary sulfur-metabolites are important as being beneficial for human. Glucosinolates in Brassicaceae plants and S-alkylcysteine sulfoxides in Alliaceae plants are known to induce detoxifying enzymes and prevent tumour formation (Talalay and Fahey, 2001; Bianchini and Vainio, 2001). They also have special odours and pungency, and are repellants against insects and microorganisms (Jones et al., 2004; Halkier and Gerschenzon, 2006; Grubb and Abel, 2006). Sulfated forms of N-acylated chitooligosac-charides are known as Nod factors of symbiotic nitrogen-fixing rhizobacteria promoting nodule formation in legumes (Fisher and Long, 1992).

As mentioned here, plants synthesize a wide variety of sulfur-compounds that serve for maintenance of cell viability. Plants are autotrophic for sulfur and able to assimilate inorganic sulfate to synthesize all these metabolites. In other words, sulfur in all metabolites present in plants derives from sulfate (Figure 4.1). The input of sulfate therefore influences the remaining metabolic pathways. This review will focus on how sulfate is transported and metabolised to cysteine and methionine, and how sulfur metabolisms are controlled under regulatory components that optimise sulfur use efficiencies, responding to environmental sulfur availabilities and intrinsic signals.

4.2 Sulfate Transport

4.2.1 Sulfate Transport Mechanisms

Uptake of sulfate is the entry step of sulfur metabolism (Figure 4.1). Plants have eukaryotic-type membrane-bound sulfate transport proteins that facilitate uptake of sulfate from extracellular space to cytoplasm across plasma membranes. At a whole-plant level, sulfate moves from cell to cell and transfers from roots to above-ground organs through xylems to fulfill the requirements of sulfur in photosynthetic tissues. Multiple transport systems eventually mediate the intercellular and interorgan transport processes. Following uptake to the cell, sulfate accumulates in vacuoles for storage or enters chloroplast/plastids.

A plasma membrane-bound protein that facilitates the uptake of sulfate to the cell requires a driving force to import sulfate, a negatively charged ion, against the gradients of membrane potential and sulfate concentration (Figure 4.2). Under these circumstances, influx of sulfate can be mediated by a secondary active co-transport system that utilises proton gradients as motive force or by an anion exchange system that needs to be coupled with an outward movement of another anion. Current understanding indicates that a proton/sulfate co-transport system probably mediates the influx of sulfate in plant cells (Lass and Ullrich-Eberius, 1984; Hawkesford et al., 1993). Proton pumping by H+-ATPase is suggested to be coupled with the influx of sulfate, but at the same time the outside positive potential can facilitate the efflux of sulfate through an unknown passive transport mechanism. This mechanism applies to accumulation of sulfate in vacuoles where H+-ATPase and H+-pyrophosphatase at tonoplast form positive potentials on vacuolar lumen side (Martinoia et al., 2000, 2007). It is known that both saturable and linear components may work as facilitators for influx of sulfate to vacuoles (Kaiser et al., 1989). Efflux of sulfate from

Glutathione Sulfate

Figure 4.1 Pathways for primary sulfur metabolism from sulfate uptake to cysteine and methionine biosynthesis in higher plants. The figure illustrates pathways for sulfate transport across plasma membranes and tonoplast, and sulfur metabolism in cytosol, chloroplasts/plastids and mitochondria in higher plants. Fluxes of metabolites are indicated by thicknesses of the lines. Putative metabolite transport pathways are indicated by dashed lines. Abbreviations: APR, APS reductase; APS, adenosine 5'-phosphosulfate; ATPS, ATP sulfurylase; CBL, cystathionine fi-lyase; CGS, cystathionine y -synthase; Cys, cysteine; GSH, glutathione; Hcy, homocysteine; Met, methionine; MS, methionine synthase; OAS, O-acetylserine; OASTL, OAS(thiol)lyase; OPH, O-phosphohomoserine; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine; Ser, serine; SERAT, serine acetyltransferase; SIR, sulfite reductase; SULTR, sulfate transporter; TS, threonine synthase

Figure 4.1 Pathways for primary sulfur metabolism from sulfate uptake to cysteine and methionine biosynthesis in higher plants. The figure illustrates pathways for sulfate transport across plasma membranes and tonoplast, and sulfur metabolism in cytosol, chloroplasts/plastids and mitochondria in higher plants. Fluxes of metabolites are indicated by thicknesses of the lines. Putative metabolite transport pathways are indicated by dashed lines. Abbreviations: APR, APS reductase; APS, adenosine 5'-phosphosulfate; ATPS, ATP sulfurylase; CBL, cystathionine fi-lyase; CGS, cystathionine y -synthase; Cys, cysteine; GSH, glutathione; Hcy, homocysteine; Met, methionine; MS, methionine synthase; OAS, O-acetylserine; OASTL, OAS(thiol)lyase; OPH, O-phosphohomoserine; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine; Ser, serine; SERAT, serine acetyltransferase; SIR, sulfite reductase; SULTR, sulfate transporter; TS, threonine synthase

SO42- H+ H+ nH+/SO42-efflux H co-transport

SO42- H+ H+ nH+/SO42-efflux H co-transport

Figure 4.2 Mechanisms of sulfate transport across plasma membrane and tonoplast. Sulfate transporters (SULTR) mediate import of sulfate to the cell across plasma membrane and efflux of sulfate from vacuoles across tonoplast. H+-ATPase and H+-pyrophosphatase (PPase) are indicated by grey squares. Proteins facilitating efflux of sulfate from the cell, and influx of sulfate to vacuoles and chloroplasts, are not known

Figure 4.2 Mechanisms of sulfate transport across plasma membrane and tonoplast. Sulfate transporters (SULTR) mediate import of sulfate to the cell across plasma membrane and efflux of sulfate from vacuoles across tonoplast. H+-ATPase and H+-pyrophosphatase (PPase) are indicated by grey squares. Proteins facilitating efflux of sulfate from the cell, and influx of sulfate to vacuoles and chloroplasts, are not known vacuoles is equally as significant as the influx of sulfate across plasma membranes, in the sense that similar mechanisms can provide sulfate to sulfur metabolic pathways. It is considered that proton gradients generated by tonoplast-bound H+-ATPase and H+-pyrophosphatase drive a proton/sulfate co-transport system to release vacuolar storage of sulfate to cytosol (Figure 4.2).

4.2.2 Sulfate Uptake System

In plant roots, the kinetics of sulfate uptake activity follows the Michealis-Menten equation (Leggett and Epstein, 1956). Analysis of sulfate uptake kinetics has indicated that a high-affinity sulfate transport protein mediates the influx of sulfate particularly under sulfur-limited conditions (Clarkson et al., 1983; Deane-Drummond, 1987). This is equivalent to phase I transport systems for the uptake of other nutrients.

Molecular cloning of sulfate transporters was first demonstrated by complementation of a chromate/selenate resistant yeast mutant strain lacking sulfate transport activities (Smith et al., 1995a; Cherest et al., 1997). Subsequently, plant cDNAs encoding high-affinity sulfate transporters were identified from a leguminous plant, Stylosanthes hamata, by functional complementation of the yeast mutant (Smith et al., 1995b). Up to now, numbers of orthologous sulfate transporters have been identified from various plant species (Smith et al., 1997; Bolchi et al., 1999; Vidmar et al., 1999, 2000; Takahashi et al., 2000; Shibagaki et al., 2002; Yoshimoto et al., 2002; Howarth et al., 2003; Buchner et al., 2004a; Hopkins et al., 2005). Their major functions are suggested to be relevant to sulfate uptake as they are expressed in the roots of sulfur-starved plants. The SULTR gene family of a model plant, Arabidopsis thaliana, is the most well characterised group of sulfate transporters. The gene family consists of 12 distinct members classified into four subgroups (Buchner et al., 2004b; Takahashi et al., 2006; Takahashi, 2010). The components of the high-affinity sulfate uptake system of Arabidopsis roots are encoded by SULTR1;1 and SULTR1;2, and their biochemical and physiological functions have been precisely demonstrated (Takahashi et al., 2000; Vidmar et al., 2000; Shibagaki et al., 2002; Yoshimoto et al., 2002).

In Arabidopsis roots, both SULTR1;1 and SULTR1;2 are expressed in epidermal and cortical cell layers where nutrients are absorbed to roots through the functions of transport proteins (Takahashi et al., 2000; Shibagaki et al., 2002; Yoshimoto et al., 2002). SULTR1;1 and SULTR1;2 are regulated by sulfur at transcript and protein levels, and deletion of both genes results in loss of viability under low-sulfur conditions (Yoshimoto et al., 2007; Barberon et al., 2008). These results indicate the significance of two sulfate transport systems in sulfate uptake, although slight differences are present. SULTR1;2 is suggested to be the major form, while SULTR1;1 seems to represent a subsidiary or compensatory function (Yoshimoto et al., 2007; Barberon et al., 2008). The major contribution of SULTR1;2 was suggested from abundance of transcripts, characteristics in sulfate uptake, and tolerance of mutants to selenate (Shibagaki et al., 2002; Yoshimoto et al., 2002, 2007; Maruyama-Nakashita etal., 2003; El Kassis etal., 2007; Barberon etal., 2008). In contrast, SULTR1;1 has been featured for its potential contribution to high-affinity sulfate uptake under extreme conditions. Its specialised functions are the low Km value for sulfate, and strong and rapid induction of transcript accumulation in response to sulfur starvation (Takahashi et al., 2000; Yoshimoto etal., 2002).

4.2.3 Transport of Sulfate from Roots to Shoots

After entering epidermal and cortical cell layers, sulfate moves horizontally through the plasmodesmata between the cells and reaches xylem parenchyma cells (Figure 4.3). During this horizontal transfer through the symplastic pathway, sulfate can be leaked from the cells to apoplast (i.e. cell wall space) but quickly retrieved back to the cells as SULTR1;1 and SULTR1;2 are present (Figure 4.3). The sulfate efflux system remains unverified. The Casparian strip of the endodermal cell layer makes a barrier for apoplastic transfer of nutrients and water to the central cylinder. Once sulfate reaches pericycle layers inside the central cylinder, it can be released to apoplast. Transporters having retrieval functions would be necessary to bring sulfate back to symplastic pathways in parenchyma cells connected to xylem vessels. According to this model, the presence of barley HVST1 sulfate transporter in pericycle and xylem parenchyma cells suggests its function for this retrieval mechanism (Rae and Smith, 2002). In Arabidopsis, low-affinity sulfate transporters probably mediate this process. SULTR2;1 and SULTR3;5 are the components suggested to co-facilitate import of sulfate and prevent the leakage of sulfate to apoplastic space in the central cylinder (Kataoka etal., 2004a) (Figure4.3). Interestingly, the activity of SULTR2;1

Shoot mesophyll cells f~r xylem and phloem parenchyma cells vacuoles chloroplasts SO42- metabolism





SO, vacuoles ~

O42- SO42-SULTR4;1 SULTR4;2

é > SO42- —-» SO42- -—»SO42---> SO42----> SO4;

apoplastic SO42-

SULTR3;5 apoplastic SO42-

epidermis cortex endodermis pericycle & xylem parenchyma cells xylem

Figure 4.3 Uptake and internal transport of sulfate in plants. Pathways operated for the uptake and root-to-shoot transport of sulfate in Arabidopsis are illustrated. Sulfate transport pathways upregulated are indicated by thick solid lines with arrowheads. Circles on the lines indicate sulfate transporters, SULTR1;1 and SULTR1;2 (black circles), SULTR4;1 and SULTR4;2 (white circles), and SULTR2;1 and SULTR3;5 (grey circles), respectively. Pathways downregulated under low-sulfur conditions or those unfavourable for sulfate uptake processes are indicated by dashed lines low-affinity sulfate transporter (Takahashi et al., 1997, 2000) was shown to be enhanced in the presence of SULRT3;5, which by itself was non-functional (Kataoka et al., 2004a). In addition to the proposed retrieval mechanism, release of sulfate from vacuoles has been shown to increase the flux of sulfate transported to shoots (Kataoka et al., 2004b) (Figure 4.3). SULTR4;1 and SULTR4;2 are the tonoplast-localising transporters mediating sulfate remobilisation. Transporters involved in these two mechanisms are generally upregulated by sulfur limitation, suggesting that the processes are dependent upon the requirement of sulfur sources in shoots (Takahashi et al., 1997, 2000; Kataoka et al., 2004a,b).


SO42- storage




4.2.4 Subcellular Transport of Sulfate

Vacuoles are the storage compartments for sulfate. The positive electrical charge gradient across the tonoplast is the most likely driving force to facilitate entry of sulfate to the vacuoles (Martinoia et al., 2000,2007). However, channel or carrier proteins mediating the influx of sulfate to vacuoles are not known. By contrast, export of sulfate from vacuoles requires SULTR4;1 and SULTR4;2 (Kataoka et al., 2004b), suggesting that these components mediate proton/sulfate co-transport mechanisms at the tonoplast. Storage and release of the vacuolar sulfate pool depend upon demands for sulfur. Regulation of this process would be important for sulfur storage and efficient sulfur utilisation (Figures 4.2 and 4.3).

Sulfate in cytosol is metabolised by a cytosolic isoform of ATP sulfurylase or transported to chloroplasts/plastids to enter the reduction pathway. Transport systems for entry of sulfate to chloroplasts still remain elusive. It is known that sulfate inhibits the activity of phosphate/triose-phosphate translocator in chloroplasts, although the interference seems to occur under very high concentrations of sulfate (Gross et al., 1990). The contribution of this system to sulfate influx is probably limited. In Chlamydomonas reinhardii, a bacterial-type sulfate transporter complex mediates the influx of sulfate to the chloroplast (Lindberg and Melis, 2008). This complex consists of a sulfate-binding protein, membrane-anchored proteins, and an ATP-binding cassette protein homologous to their bacterial origins (Sirko et al., 1990; Laudenbach and Grossman, 1991). Orthologous proteins for this system have not been identified from higher plant species, suggesting that the mechanism is probably different from those in algae (Figure 4.2).

4.2.5 Redistribution of Sulfur

Redistribution of sulfur from source to sink organs occurs through phloem. Besides sulfate, glutathione and S-methylmethionine are present in phloem sap (Bourgis et al., 1999; Herschbach et al., 2000; Kuzuhara et al., 2000). In Arabidopsis leaves, SULTR2;1 low-affinity sulfate transporter gene is expressed in parenchyma cells around the phloem and xylem (Takahashi et al., 2000). SULTR2;1 was downregulated by sulfur limitation in leaf vasculature (Takahashi et al., 2000). This would prevent retrieval of sulfate back to xylem and to phloem, which allows sulfate to be transferred to mesophyll cells (Figure 4.3). Eventually, distribution of sulfur via phloem can be restricted under low-sulfur conditions. The model suggests that the mobility of sulfate and/or other sulfur-compounds from old to young leaves declines during sulfur starvation. SULTR2;1 also functions for translocation of sulfur source to developing seeds where its expression is found in funiculus and vasculature of seed pods (Awazuhara et al., 2005). It has been shown that developing seeds respond to sulfur deficiency and accumulate sulfurless seed storage proteins (Hirai et al., 1995). However, it has not been defined whether deficits of sulfate or other sulfur-metabolites trigger this response in seeds. The metabolic pathways for synthesis of sulfur-compounds are present in developing cotyledons of lupin (Tabe and Droux, 2001). On the other hand, glutathione synthesis is active in embyos and funiculus of Arabidopsis, allowing the possibility that organic sulfur-compounds are synthesized from sulfate and transported to seeds (Cairns et al., 2006).

Relevant to phloem transport, a high-affinity sulfate transporter, SULTR1;3, is shown to localise in companion cells of phloem (Yoshimoto et al., 2003). The knockout of SULTR1;3

restricted transport of sulfur from cotyledons to shoot meristem and roots in Arabidopsis, suggesting significance for this transporter in source-to-sink transport of sulfur (Yoshimoto etal., 2003). Accumulation of SULTR1;3 transcripts in sulfur-starved plants further suggests its importance under sulfur-limited conditions. The induction of SULTR1;3 transcripts on low sulfur would make a considerable contribution to increasing the amount of sulfate loaded to phloem. It is also possible that transportable forms of sulfur-metabolites were decreased by repression of SULTR2;1 in phloem parenchyma cells (Takahashi et al., 2000), and the induction of SULTR1;3 could have occurred in response to local sulfur deficiency. In addition to SULTR1;3, the low-affinity isoform, SULTR2;2, has been reported to localise in phloem companion cells (Takahashi et al., 2000).

Vacuolar sulfate transporters are also suggested to participate in redistribution of sulfur. Accumulation of SULTR4;1 and SULTR4;2 transcripts in senescing leaves of Brassica napus plants suggests that sulfate pools are remobilised during senescence and most likely transferred to sink organs (Dubousset et al., 2009). Sulfate in vacuole is considered to be temporal storage of sulfur, and will be released by induction of SULTR4;1 and SULTR4;2 during sulfur limitation or senescence to allow it to transfer to sink organs.

4.2.6 Regulation of Sulfate Uptake

Sulfate transporters in roots are primarily regulated by sulfate availability (Smith et al., 1995b, 1997; Takahashi etal., 2000; Vidmar etal., 2000; Shibagaki etal, 2002; Yoshimoto et al., 2002; Maruyama-Nakashita et al., 2004a). The amount of sulfate supply affects synthesis of downstream sulfur-metabolites such as cysteine and glutathione. These metabolites can feedback-regulate the expression of sulfate transporters (Herschbach and Rennenberg, 1991; Smith etal., 1997; Bolchi etal., 1999; Lappartient etal., 1999; Vidmar et al., 1999, 2000; Maruyama-Nakashita et al., 2004b). It is further suggested that sulfate to glutathione ratios of phloem sap correlate with rates of sulfate acquisition in roots (Herschbach and Rennenberg, 1991; Herschbach et al., 2000). In addition, interorgan repressive signals are possibly related to glutathione in phloem, regulating the amount of low-affinity sulfate transporter gene, SULTR2;1, expressed in roots (Lappartient and Touraine, 1996; Lappartient etal., 1999).

As for the positive signals, O-acetylserine (OAS) can induce the expression of sulfate transporters by external application (Smith et al., 1997; Maruyama-Nakashita et al., 2004b, 2005). OAS is the precursor of cysteine. It is suggested that OAS can override negative feedback effects of cysteine and glutathione accumulated in the cells (Smith et al., 1997). It is not known how these metabolites may stimulate signalling cascades, however, microarray studies suggest significant overlaps between transcriptomes of OAS treatment and sulfur starvation in Arabidopsis (Hirai et al., 2003, 2004; Nikiforova et al., 2003). The sulfur responsive element (SURE) present in the 5'-region of SULTR1;1 has been shown to respond not only to sulfate but also cysteine, glutathione and OAS (Maruyama-Nakashita et al., 2005). This provides additional evidence that metabolic effectors actually influence the sulfur-responsive regulatory pathway. However, as suggested by the absence of SURE in the 5'-region of SULTR1;2, the regulatory pathways are not always uniform among the two sulfur-responsive high-affinity sulfate transporters. Nevertheless, both of them show similar responses to sulfur limitation and effector metabolites.

In addition to sulfur specific regulations, sulfate transporters are known to be regulated by other general factors. As an intrinsic signal, a plant hormone cytokinin was shown to repress the expression of high-affinity sulfate transporters in Arabidopsis roots (Maruyama-Nakashita et al., 2004b). Signals downstream of cytokinin receptor CRE1 (Inoue et al., 2001) diminished the transcript levels of SULTR1;1 and SULRT1;2; however, their sulfur-deficiency responsiveness was not substantially affected (Maruyama-Nakashita et al., 2004b). It is suggested that cytokinin functions as a general repressive signal to modulate the levels of sulfate transporters and sulfate uptake rates in roots. Generality of the cytokinin signalling pathway in the regulation of nutrient acquisition has been suggested from its involvement in controlling phosphorus response (Martin et al., 2000; Franco-Zorrilla et al., 2002). Besides hormone signals, carbon and nitrogen status are general factors that may enhance the levels of high-affinity sulfate transporters in roots (Vidmar et al., 1999; Lejay et al., 2003; Wang et al., 2003; Maruyama-Nakashita et al., 2004c). In general, SULTR1;1 is regulated more specifically by sulfur conditions, whereas SULTR1;2 accepts broader environmental signals for regulation (Rouached et al., 2008).

The promoter analysis of sulfur responsiveness of SULTR1;1 gene expression indicates the presence of an auxin response factor binding sequence (Ulmasov et al., 1999; Hagen and Guilfoyle, 2002) within the cis-element (Maruyama-Nakashita etal., 2005). Although a potential binding sequence in the cis-element was relevant to a general growth regulator, the response mediated under this element was rather specific to sulfur (Maruyama-Nakashita et al., 2005). Identification of SLIM1/EIL3 from Arabidopsis indicates another unique upstream pathway significant for regulation of both SULTR1;1 and SULTR1;2 during sulfur limitation (Maruyama-Nakashita et al., 2006). The role of this transcription factor will be described in the last section of this chapter.

As for post-transcriptional regulation, SULTR1;1 and SULRT1;2 are shown to be regulated at protein levels (Yoshimoto et al., 2007). Over-accumulation of SULTR proteins occurred during sulfur limitation even though the genes were expressed under a constitutive promoter. This provides evidence for the significance of SULTR1;2 whose transcript accumulates only moderately in response to sulfur limitation under the native promoter. Another mode of post-transcriptional regulation involves the function of its C-terminus region. Sulfate transporter has a hydrophilic STAS (sulfate transporter and anti-sigma factor antagonist) domain in its C-terminus (Aravind and Koonin, 2000). Plasma membrane localisation and function of sulfate transporter are suggested to be largely influenced when mutations are introduced around a putative phosphorylated residue in the STAS domain (Shibagaki and Grossman, 2004, 2006; Rouached et al., 2005). Other than the STAS domain, polar residues present in the first and second predicted membrane spanning regions are shown to be necessary for the activity of sulfate transporter (Leves et al., 2008).

4.3 Sulfate Reduction 4.3.1 ATP Sulfurylase

ATP sulfurylase is the first committing enzyme for reduction of sulfate (Figure 4.1). The enzyme catalyzes the reaction, generating adenosine 5'-phosphosulfate (APS) and pyrophosphate from sulfate and ATP. Removal of pyrophosphate drives the reaction, otherwise the enzyme catalyzes the reverse reaction. ATP sulfurylase activity is present in both cytosol and chloroplasts (Lunn et al., 1990; Renosto et al., 1993; Rotte and Leustek, 2000). cD-NAs encoding these subcellular specific isoforms are identified from potato (Klonus et al., 1994). Arabidopsis has four ATP sulfurylase genes, ATPS1 to ATPS4. Proteins encoded by these four ATPS genes in Arabidopsis contain chloroplast-targeting transit peptides in their ^-terminal regions (Leustek et al., 1994; Murillo and Leustek, 1995; Logan et al., 1996; Hatzfeld et al., 2000a), although ATPS2 is predicted to be translated also as a cytosolic isoform (Hatzfeld et al., 2000a). It still remains unresolved how these four ATPS isoforms share their functions in planta.

4.3.2 APS Reductase

In higher plants, APS is directly reduced to sulfite by APS reductase (APR) (Gutierrez-Marcos et al., 1996; Setya et al., 1996; Prior et al., 1999; Suter et al., 2000) (Figure 4.1). The pathway is different from those operating in fungi where APS is first phosphorylated by APS kinase (APK) to form 3'-phosphoadenosine 5'-phosphosulfate (PAPS), then PAPS reductase subsequently serves for enzymatic conversion of PAPS to sulfite. Some bacteria and lower plants have both pathways (Koprivova et al., 2002). Thus, plant APR is suggested to derive evolutionally from PAPS reductase (Kopriva and Korpivova, 2004). Three APR genes, APR1, APR2 and APR3, are identified from Arabidopsis (Gutierrez-Marcos et al., 1996; Setya et al., 1996). APR is a nuclear encoded chloroplast/plastid-localising enzyme. Consistent with predicted subcellular localisations of APR isoforms, the enzyme activity was found only in chloroplasts/plastids (Rotte and Leustek, 2000). The mature APR enzyme consists of a catalytic domain similar to PAPS reductase, and a C-terminal thioredoxin-like domain which functions as a glutaredoxin and utilises reduced glutathione as a reductant (Bick et al., 1998; Prior et al., 1999; Koprivova et al., 2002).

In sulfate reduction pathways of plants, APR is the only metabolic enzyme strongly upregulated by sulfur starvation at transcript levels (Gutierrez-Marcos et al., 1996; Takahashi et al., 1997). APR is suggested to be the flux-controlling enzyme (Vauclare et al., 2002). The significance of this pathway has been suggested from studies with ectopic overexpression of a Pseudomonas APR gene as a chloroplast-targeted form in Arabidopsis (Tsakraklides et al., 2002). Sulfate reduction and cysteine synthesis were enhanced in overexpressors, suggesting that the step catalyzed by APR is rate-limiting under normal conditions (Tsakraklides et al., 2002). Furthermore, APR2 is identified as a locus determining sulfate accumulation in Arabidopsis natural variation, suggesting significance of this enzyme in driving sulfate reduction pathways (Loudet et al., 2007).

4.3.3 APS Kinase

In higher plants, APK and APR share a common substrate, APS, at the juncture of the sulfate reduction pathway (Leustek et al., 2000; Saito, 2000; Kopriva and Korpivova, 2004; Kopriva, 2006) (Figure 4.1). PAPS is not a substrate of sulfate reduction, but instead it is used exclusively for sulfation reactions in plants (Klein and Papenbrock, 2004; Piotrowski etal., 2004; Hirai etal., 2005; Klein etal., 2006). In fact, the amounts of sulfated metabolites including glucosinolates were reduced in the knockout of APK isoforms in Arabidopsis (Mugford et al., 2009). As mentioned in the following subsections, APK and APR were oppositely regulated to direct the flux of sulfur for use in cysteine biosynthesis or secondary sulfur metabolism, depending on sulfur availabilities (Takahashi et al., 1997; Maruyama-Nakashita et al., 2003; Hirai et al., 2003, 2004).

Arabidopsis has four APK isoforms (Lee and Leustek, 1998; Lillig et al., 2001). APK1, APK2 and APK4 are localised in chloroplasts/plastids, while APK3 encodes a cytosolic isoform (Mugford et al., 2009). Synthesis of its substrate, APS, occurs in both compartments. Arabidopsis ATPS2 is predicted to encode both chloroplastic and cytosolic isoforms (Hatzfeld et al., 2000a), and a cytosolic ATPS exists in potato (Klonus et al., 1994). Considering subcellular localisation of APK and ATPS isoforms, conversion of sulfate to APS and further phosphorylation to PAPS may occur partly in cytosol; however, two chloroplast/plastid-localising APK isoforms, APK1 and APK2, are the major components that make significant contributions to the generation of PAPS for sulfation reactions in Arabidopsis (Mugford et al., 2009). On the other hand, as for the use of PAPS in sulfation reactions, sulfotransferases are shown to localise in cytosol or are predicted to be in compartments other than chloroplasts (Klein and Papenbrock, 2004; Klein et al., 2006). These observations still leave us to consider cytosolic pathways for ATPS and APK, and the relevance of transport of PAPS across chloroplast membranes (Figure 4.1).

4.3.4 Sulfite Reductase

Sulfite reductase (SIR) is a chloroplasts/plastids-localising enzyme that generates sulfide from sulfite using reduced ferredoxin as a reductant (Krüger and Siegel, 1982) (Figure 4.1). Photosynthetic electron transfer provides reduced ferredoxin in leaves, while NADPH generated from the oxidative pentose phosphate pathway reduces ferredoxin in roots (Yonekura-Sakakibara etal., 1998,2000). The Arabidopsis SIR is a single copy gene (Brühl etal., 1996; Bork et al., 1998), suggesting that the reaction is a rate-limiting step of sulfate reduction. However, SIR transcripts are not regulated significantly by sulfur conditions (Takahashi et al., 1997). As mentioned previously, APR and APK are the enzymes regulated at transcript levels (Takahashi et al., 1997; Maruyama-Nakashita et al., 2003; Hirai et al., 2003). SIR appears to be present as an abundant enzyme that helps rapid conversion of toxic sulfite to sulfide. SIR is also known as a chloroplast DNA-binding protein. Nucleoid attachment of this enzyme is relevant to compaction of DNA and is suggested to regulate transcription in chloroplasts (Sekine et al., 2002, 2007; Chi-Ham et al., 2002).

4.3.5 Regulation of Sulfate Reduction

Sulfate reduction pathways are regulated under complex mechanisms in plants. As for the entry step of sulfate reduction, ATPS transcript levels are not always regulated positively for reduction of sulfate in sulfur-starved plants. The regulatory mechanism for ATPS appears to be complicated. In Arabidopsis, ATPS1 and ATPS3 are known as isoforms whose mRNAs are moderately accumulated by sulfur limitation and repressed by feedback regulation responding to demands for sulfur (Takahashi etal., 1997; Lappartient etal., 1999). However, recent studies indicate an additional mechanism of post-transcriptional regulation. Under sulfur deficiency, microRNA-395 (miR395) accumulates to destabilise ATPS1 and ATPS3 transcripts (Jones-Rhoades and Bartel, 2004; Allen et al., 2005; Kawashima et al., 2009). ATPS2 lacks the miR395 target sequence and is insensitive to this mechanism. ATPS4 is an isoform repressed under sulfur deficiency and is subject to miR395-mediated regulation (Kawashima et al., 2009). More importantly, SLIM1, a transcription factor that controls the sulfur limitation response (Maruyama-Nakashita et al., 2006), is an upstream regulator of miR395 which in turn negatively controls ATPS transcript accumulation (Kawashima etal., 2009).

Regulation of sulfate reduction essentially occurs at the juncture of primary and secondary sulfur metabolism in plants. As described in the previous subsection, APR and APK share a common substrate, APS. When supply of sulfate is limited, APR transcripts will increase (Gutierrez-Marcos et al., 1996; Takahashi et al., 1997), while those for APK decrease significantly (Maruyama-Nakashita et al., 2003; Hirai et al., 2003). Assuming that transcript levels correlate with abundance of enzymes being translated, the mechanism favours an increased sulfur flux in the reduction pathway. This would ensure the plants can primarily synthesize essential sulfur-metabolites such as cysteine and methionine under sulfur deficiency. By contrast, APK is likely to be an unfavourable enzyme for primary metabolism, as the metabolic pathway is branched for secondary metabolism. It appears that synthesis of PAPS is allowed mostly under sulfur-rich conditions.

APR is a key metabolic enzyme that drives the reduction of sulfate in plants. It is primarily regulated at transcript levels by the availability of sulfate (Gutierrez-Marcos et al., 1996; Takahashi et al., 1997). In addition, the isoforms of APR in Arabidopsis are responsive to OAS, cysteine and GSH as expected (Koprivova et al., 2000; Vauclare et al., 2002; Hesse et al., 2003). Besides regulation by sulfur, APR transcript accumulations are dependent on supplies of nitrogen and carbon sources that provide the backbone of cysteine (Kopriva et al., 1999, 2002; Koprivova et al., 2000; Hesse et al., 2003; Wang et al., 2003). Supply of carbon may increase the amount of reductant used for the APR reaction, as NADPH can eventually reduce glutathione. In addition to general metabolic balancing, plant hormones participate in the regulation of APR. Upregulation of APR by cytokinin is suggested to be caused by sucrose accumulation (Ohkama et al., 2002) or by reduced uptake of sulfate (Maruyama-Nakashita et al., 2004b). Jasmonate upregulates pathways of sulfate reduction and metabolism in addition to APR, suggesting a mechanism to alleviate oxidative stress and/or to induce defence response (Jost et al., 2005). At the level of enzyme activities, Arabidopsis APR1 was activated under oxidative conditions, suggesting an additional mode of regulation for this key enzyme (Bick et al., 2001).

4.4 Cysteine Biosynthesis 4.4.1 0-Acetylserine(thiol)lyase

Following reduction of sulfate, sulfide is transferred to O-acetylserine to form cysteine and acetate. This reaction is catalyzed by O-acetylserine(thiol)lyase (OASTL) which localises in cytosol, chloroplasts/plastids and mitochondria (Lunn et al., 1990) (Figure 4.1). Sulfide, the substrate of OASTL, needs to be transported between subcellular compartments as SIR is present exclusively in chloroplasts/plastids. Numbers of genes encoding OASTL have been identified and characterised (Saito et al., 1992, 1994a; Youssefian et al., 1993; Ruffet et al., 1994; Hell et al., 1994; Barroso et al., 1995; Jost et al., 2000). The chloroplast/plastid-localising isoform existed in excess amounts and was suggested to play major roles in cysteine biosynthesis, directly utilising sulfide from SIR (Ruffet et al., 1994). Cytosolic isoforms of OASTL would also accept atmospheric sulfide (Gotor et al., 1997). Loss-of-function mutants of OASTL isoforms further indicated the significance of cysteine biosynthesis in cytosol and chloroplasts (Heeg et al., 2008; Watanabe et al., 2008a). Importance was also suggested from the overexpression of OASTL that makes plants tolerant to sulfite and sulfide (Youssefian et al., 1993; Saito et al., 1994b; Noji et al., 2001). The functions of OASTL families are diverse. In mitochondria, cysteine generated by OASTL serves as a substrate for ß-cyanoalanine synthase, which is a close homologue of OASTL (Hatzfeld etal., 2000b; Yamaguchi etal., 2000; Maruyama et al., 2001). Recent findings indicate that sulfocysteine synthase, which utilises thiosulfate as a specific substrate, belongs to this family (Bermudes et al., 2010). Knockout of this isoform results in accumulating reactive oxygen species and impaired growth.

4.4.2 Serine Acetyltransferase

O-acetylserine (OAS), the precursor of cysteine biosynthesis, is synthesized by serine acetyltransferase (SERAT). SERAT activity was found in cytosol, chloroplasts/plastids and mitochondria (Ruffet et al., 1995) (Figure 4.1). Arabidopsis has five SERAT isoforms showing distinctive features for OAS biosynthesis and localisation (Ruffet et al., 1995; Noji et al., 1998; Hell et al., 2002; Kawashima et al., 2005; Haas et al., 2008; Watanabe et al., 2008b). As OASTL accumulates in excess in chloroplasts (Ruffet et al., 1994), cysteine can be readily formed following synthesis of OAS. However, OASTL does not exist abundantly in mitochondria. This allows OAS, the product of SERAT, to be exported from mitochondria to cytosol and converted to cysteine by cytosolic OASTL. Genetic reconstitution of each SERAT isoform in quintuple mutant of Arabidopsis indicated that mitochondrial SERAT2;2 and cytosolic SERAT1;1 make major contributions to OAS synthesis (Watanabe et al., 2008b). Current understandings indicate that mitochondrial SERAT2;2 (SAT3) most likely controls the rate of OAS synthesis and makes the largest contribution among the isoforms (Haas et al., 2008; Watanabe et al., 2008b). By contrast, contribution of chloro-plastic SERAT2;1 was suggested to be limited (Watanabe et al., 2008b). Consistent with these findings, it has been suggested that OAS can be limiting in chloroplasts. Some trans-genic studies provided supporting evidence. Biosynthesis of cysteine and glutathione was enhanced by overexpression of bacterial SERAT as a chloroplast-targeted form in plants (Blaszczyk et al., 1999; Harms et al., 2000). Overexpression of Arabidopsis chloroplastic SERAT2;1 to lupin led to an enhanced production of OAS, cysteine and glutathione in developing seeds (Tabe et al., 2010). Furthermore, external feeding of OAS significantly enhanced the rates of cysteine and glutathione production in transgenic plants overexpress-ing Pseudomonas-derived APR as a chloroplast-targeted form (Tsakraklides et al., 2002). Accordingly, transport of OAS to chloroplasts is likely to be a necessary process for cysteine biosynthesis by OASTL (Figure 4.1).

Cytosolic isoforms of SERAT were either sensitive to feedback inhibition by cysteine, or less active unless sufficient amount of substrates were supplied (Noji et al., 1998; Kawashima et al., 2005). Specific functions are suggested for cytosolic SERAT3;1 and SERAT3;2 under sulfur deficiency and in reproductive organs (Kawashima et al., 2005; Watanabe et al., 2008b). On the other hand, it was evident that another cytosolic isoform, SERAT1;1, makes a substantial contribution to OAS synthesis (Watanabe et al., 2008b), although the activity of this enzyme was feedback-inhibited by cysteine (Noji et al., 1998). These observations may indicate that cysteine concentration is strictly maintained below the inhibitory levels in the cytosol. Alternatively, the enzyme could be functional as a cysteine synthase complex (described in the following subsection). For this model, tight control of OAS concentration would become necessary, as the complex will dissociate when OAS accumulates in excess (Droux et al., 1998). OAS needs to be metabolised or sequestered to maintain the activities of SERAT isoforms in cytosol and mitochondria. OAS can be metabolised by OASTL in cytosol or transported to chloroplasts where a huge demand for cysteine biosynthesis should exist (Heeg et al., 2008; Watanabe et al., 2008a) (Figure 4.1).

4.4.3 Cysteine Synthase Complex

SERAT and OASTL form cysteine synthase complex, which tightly controls cysteine biosynthesis through changes of conformation (Droux et al., 1998; Berkowitz et al., 2002; Wirtz and Hell, 2006). Dissociation and reversible re-association of the complex occurs in the presence of OAS and sulfide, respectively (Droux et al., 1998) (Figure 4.4). When the complex dissociates in the presence of excess OAS, SERAT aggregates and becomes inactive, while unbound free OASTL converts excess OAS to cysteine. Under this mechanism, OAS will soon be depleted and sulfide accumulates. The complex re-associates in the presence of sulfide and serves as SERAT to produce OAS. The entire cycle controls cysteine biosynthesis depending upon OAS and sulfide supplied to the system. In addition, SERAT itself was feedback-inhibited by cysteine unless bound to OASTL (Figure 4.4). The feedback inhibition occurred particularly for cytosolic SERAT, playing a major role in cysteine synthesis (Noji et al., 1998; Watanabe et al., 2008b). Besides these control mechanisms, the amounts of SERAT and OASTL accumulated in each subcellular compartment are likely to vary, suggesting the significance of compartmentation and enzymatic properties of each high sulfur S2- in excess low sulfur OAS in excess

Cys high sulfur S2- in excess low sulfur OAS in excess


Figure 4.4 Regulatory mechanisms of cysteine biosynthesis. Under high-sulfur conditions where sulfide is in excess, cysteine synthase complex assembles to function as SERAT. The activity of free SERAT can be feedback-inhibited by cysteine unless it is bound to OASTL. Under low-sulfur conditions, OAS will accumulate and dissociates the cysteine synthase complex to stop OAS production. Abbreviations: Cys, cysteine; OAS, O-acetylserine; OASTL, OAS(thiol)lyase; Ser, serine; SERAT, serine acetyltransferase

Figure 4.4 Regulatory mechanisms of cysteine biosynthesis. Under high-sulfur conditions where sulfide is in excess, cysteine synthase complex assembles to function as SERAT. The activity of free SERAT can be feedback-inhibited by cysteine unless it is bound to OASTL. Under low-sulfur conditions, OAS will accumulate and dissociates the cysteine synthase complex to stop OAS production. Abbreviations: Cys, cysteine; OAS, O-acetylserine; OASTL, OAS(thiol)lyase; Ser, serine; SERAT, serine acetyltransferase isoform to fulfil the consecutive reactions for OAS and cysteine biosynthesis (Haas et al., 2008; Heeg et al., 2008; Watanabe et al., 2008a,b) (Figure 4.1).

4.5 Methionine Biosynthesis 4.5.1 Biosynthetic Pathways

Cystathionine y -synthase (CGS) catalyzes the first step of methionine biosynthesis (Figures 4.1 and 4.5). Cysteine and O-phosphohomoserine (OPH) are the substrates for synthesis of cystathionine in this step. Cystathionine p-lyase (CBL) subsequently catalytes the cleavage of cystathionine to form homocysteine. CGS and CBL are present only in chloroplasts (Figure 4.1). The significance of these chloroplast-localised metabolic pathways is suggested by the need for CGS and CBL for plant growth (Ravanel et al., 1998a; Maimann et al., 2000). CGS and CBL were encoded by single or few copies of genes, indicating that the pathways are indispensable (Kim and Leustek, 1996; Ravanel et al., 1995). Following consecutive reactions catalyzed by CGS and CBL, homocysteine is subsequently methylated to form methionine by methionine synthase (MS) using methyltetrahydrofolate as a methyl donor (Eichel et al., 1995; Ravanel et al., 1998a, 2004; Zeh et al., 2002). This final step is present in both chloroplasts and cytosol (Figure 4.1). Arabidopsis has three isoforms of MS showing distinct subcellular localisations. MS1 and MS2 are the cytosolic forms, while MS3 is located in chloroplasts (Ravanel et al., 2004). Methionine in cytosol is further metabolised to S-adenosylmethionine (SAM) by cytosolic SAM synthetase, and

Cys mRNA stability

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