Photosynthesis Research 82: 203–220, 2004.
2004 Kluwer Academic Publishers. Printed in the Netherlands.

203

Review

The thioredoxin superfamily in Chlamydomonas reinhardtii Ste´phane D. Lemaire* & Myroslawa Miginiac-Maslow Institut de Biotechnologie des Plantes, Universite´ Paris-Sud, UMR 8618 CNRS, Baˆtiment 630, 91405 Orsay Cedex, France; *Author for correspondence (e-mail: [email protected]; fax: +33-1-69153425) Received 27 November 2003; accepted in revised form 23 February 2004

Key words: Chlamydomonas, gene family, genome, glutaredoxin, protein disulfide isomerase, redox, thioredoxin

Abstract The thioredoxin (TRX) superfamily includes redox proteins such as thioredoxins, glutaredoxins (GRXs) and protein disulfide isomerases (PDI). These proteins share a common structural motif named the thioredoxin fold. They are involved in disulfide oxido-reduction and/or isomerization. The sequencing of the Arabidopsis genome revealed an unsuspected multiplicity of TRX and GRX genes compared to other organisms. The availability of full Chlamydomonas genome sequence offers the opportunity to determine whether this multiplicity is specific to higher plant species or common to all photosynthetic eukaryotes. We have previously shown that the multiplicity is more limited in Chlamydomonas for TRX and GRX families. We extend here our analysis to the PDI family. This paper presents a comparative analysis of the TRX, GRX and PDI families present in Arabidopsis, Chlamydomonas and Synechocystis. The putative subcellular localization of each protein and its relative expression level, based on EST data, have been investigated. This analysis provides a large overview of the redox regulatory systems present in Chlamydomonas. The data are discussed in view of recent results suggesting a complex cross-talk between the TRX, GRX and PDI redox regulatory networks. Abbreviations: EST – expressed sequence tag; GRX – glutaredoxin; GSH – Glutathione; GST – glutathione-S-transferase; PDI – protein disulfide isomerase; PRX – peroxiredoxin, RNR – Ribonucleotide reductase; ROS – reactive oxygen species; TRX – thioredoxin

Introduction The thioredoxin superfamily encompasses proteins sharing a common structural motif named a thioredoxin domain or ‘thioredoxin fold’ (Martin 1995). Thioredoxins (TRXs) are small ubiquitous redox proteins found in all free-living organisms. They reduce disulfide bridges of various target proteins by thiol–disulfide exchange reactions and are involved in a large variety of functions (reviewed in Vlamis-Gardikas and Holmgren 2002). TRXs are reduced by NADPH through an NADPH thioredoxin reductase (NTR) located in the cytosol and mitochondria. In the chloroplast

they are reduced from photosynthetically reduced ferredoxin by ferredoxin thioredoxin reductase (FTR). TRX structure is characterized by the following succession of secondary structural elements: b1, a1, b2, a2, b3, a3, b4, b5, a4. The ‘thioredoxin fold’, as defined by Martin (1995) corresponds to the same succession without a1 and b1. All members of the TRX superfamily contain one or several thioredoxin domains. Five major groups of proteins are part of this superfamily: thioredoxin, glutaredoxin (GRX), protein disulfide isomerase (PDI), glutathione peroxidase and glutathione-S-transferase (GST). The present study will focus on the TRX, GRX and PDI families.

204 The sequencing of Arabidopsis revealed an unsuspected multiplicity of TRX genes. A total of 19 TRXs have now been identified in Arabidopsis while only four are present in the cyanobacterium Synechocystis PCC6803 (Lemaire et al. 2003a). This multiplicity raises questions about the redundancy and specificity of TRXs in photosynthetic organisms. One of the strategies used to try to answer these questions was the complementation of a yeast TRX deficient strain, exhibiting a pleiotropic phenotype, by plant TRXs. Complementation experiments by Arabidopsis cytosolic (Mouaheb et al. 1998; Bre´he´lin et al. 2000) or chloroplastic (Issakidis-Bourguet et al. 2001) TRXs revealed that some specificities can be distinguished but also that a partial redundancy is observed. The multiplicity of TRXs is also a problem for reverse genetic approaches since most single knock-out mutants of TRX in Arabidopsis do not exhibit phenotypes probably due to functional complementation and compensation by other TRXs and/or other redox systems. One could also wonder if this multiplicity is a particularity of Arabidopsis, of higher plants or of all photosynthetic eukaryotes. The availability of Chlamydomonas reinhardtii genome sequence offers the opportunity to answer this question. We have recently shown that Chlamydomonas contains only eight different TRXs (Lemaire et al. 2003b). The analysis of the GRX families also revealed a much lower number of GRXs in Chlamydomonas compared to Arabidopsis with only six GRX genes in the green alga compared to 30 in the higher plant genome (Lemaire 2004). GRXs are small redox proteins that are also able to reduce disulfide bridges and can have functions partially redundant with those of TRXs (Vlamis-Gardikas and Holmgren 2002). However, GRXs are reduced by glutathione generated from NADPH by glutathione reductase (GR). In this paper we extend our analysis to the protein disulfide isomerase family. These proteins are found in eukaryotes and are composed of four TRX motifs, two of which are redox-active. The vicinal dithiol of their active site (WCGHC) is able to promote the formation, reduction and/or isomerization of disulfide bridges. In addition, they have a chaperone activity and assist the folding of proteins (reviewed in Ferrari and So¨ling 1999). They are usually located in the endoplasmic reticulum (ER). However, the Rb60 protein, which is the only PDI

characterized in Chlamydomonas, is a chloroplastic protein involved in the control of psbA translation by light (Trebitsh et al. 2000; Trebitsh and Danon 2001). This indicates that the functions of PDIs are not restricted to the ER but that they can also participate in redox signaling. While PDIs have been extensively studied in mammals (Ferrari and So¨ling 1999; Clissold and Bicknell 2003), very little is known about these proteins in photosynthetic organisms where the number and diversity of PDIs remains to be examined. We have analyzed the genomes of Arabidopsis and Chlamydomonas in order to identify members of the PDI family. In this paper we present an overview of the TRX, GRX and PDI gene families present in Chlamydomonas reinhardtii. Each gene family is presented in detail including putative sub-cellular localization of the proteins, and their relative abundance based on EST data. The Chlamydomonas gene families are compared to those of Arabidopsis and Synechocystis PCC6803. The results are discussed in view of recent data suggesting a complex cross-talk between the TRX, GRX and PDI regulatory networks.

The thioredoxin family The biochemical function of thioredoxins is fulfilled through the vicinal cysteines of their protruding active site. The sequence of this conserved active site is WCGPC or WCPPC. Thioredoxin was first identified as a hydrogen donor to ribonucleotide reductase (RNR) in E. coli (Laurent et al. 1964). Since then, they have been identified in all free-living organisms. Non-photosynthetic eukaryotes contain a limited number of TRXs with usually one or two isoforms in the cytosol and one in the mitochondria. These TRXs are reduced by an NADPH thioredoxin reductase (NTR) present in the cytosol or the mitochondria. They have been shown to reduce methionine sulfoxide by way of methionine sulfoxide reductases, participate in the detoxification of reactive oxygen species (ROS) by way of peroxiredoxins (PRX), participate in sulfate assimilation by reducing PAPS reductase and control the DNA-binding activity of numerous transcription factors (VlamisGardikas and Holmgren 2002). In plants, two types of thioredoxins named TRX f and TRX m were initially identified (reviewed in Schu¨rmann

205 and Jacquot 2000; Jacquot et al. 2002). These TRXs are located in the chloroplasts and are reduced in the light by photoreduced ferredoxin and ferredoxin thioredoxin reductase. Once reduced, these TRXs are able to reduce regulatory disulfides on their target enzymes. The first TRX targets identified were key enzymes of the carbon metabolism such as Calvin-cycle enzymes (fructose-1,6-bisphosphatase (FBPase), phosphoribulokinase, glyceraldehyde-3-phosphate dehydrogenase), NADP-malate dehydrogenase (NADP-MDH), Rubisco activase or the c subunit of ATP synthase. These enzymes are inactive in the dark and are activated under illumination by the ferredoxin/thioredoxin system. A decade later a third TRX type, located in the cytosol, was discovered and named TRX h (for heterotrophic) (reviewed in Rouhier et al. 2002b). These TRXs are reduced by NADPH and NTR and are involved in the mobilization of seed reserves during germination (Kobrehel et al. 1992; Besse et al. 1996; Cho et al. 1999). They are also involved in self-incompatibility mechanisms (Cabrillac et al. 2001). However, their function in green leaves remains to be determined. Biochemical studies led more than 10 years ago, allowed identifying all three TRX isoforms in Chlamydomonas, (Huppe and Buchanan 1989; Decottignies et al. 1990, 1991; Stein et al. 1995) along with their respective reductases, i.e., FTR (Huppe et al. 1990) and NTR (Huppe et al. 1991). The expression of Chlamydomonas TRXs m and h was shown to be regulated by light, circadian clock and heavy metals (Lemaire et al. 1999a, b, 2002). This simple picture of the TRX system in plants has become incredibly more complicated with the availability of genomic data. The sequencing of Arabidopsis genome revealed the existence of multiple isoforms of TRX. This led to the discovery of mitochondrial TRXs (TRX o) (Laloi et al. 2001) and a new type of TRX named TRX x (Mestres-Ortega and Meyer 1999). We have recently shown that TRX x is localized in the chloroplast and exhibits almost no activity in vitro in the activation of classical TRX targets such as FBPase or NADP-MDH (Collin et al. 2003). On the other hand, TRX x appears to be particularly efficient for reduction of chloroplastic 2-cys peroxiredoxin (2-cys PRX), a TRX-dependent peroxidase. The analysis of EST data available in Chlamydomonas allowed us to identify a new type of TRX, previously undescribed, that we named

TRX y. Two genes encoding TRX y are present in Arabidopsis but were not initially considered as true TRXs. The biochemical characterization of Chlamydomonas reinhardtii TRX y (CrTRXy) showed that this TRX exhibits specificities completely different from other TRX types but its target probably remains to be identified. A phylogenetic tree including TRXs from Arabidopsis, Chlamydomonas and Synechocystis PCC6803 is presented on Figure 1. Arabidopsis contains 19 TRXs (8 TRX h, 2 TRX o, 4 TRX m, 2 TRX f, 1 TRX x and 2 TRX y) compared to 8 TRXs in Chlamydomonas (2 TRX h, 1 TRX o, 1 TRX m, 2 TRX f, 1 TRX x, 1 TRX y). Two additional proteins related to TRXs (DLC14 and DLC16) are specific to Chlamydomonas and are located in the outer arm dynein, in the flagella (Patel-King et al. 1996). They are suggested to participate in a redox-based control of dynein activity (Harrison et al. 2002). In contrast, Synechocystis only contains four TRXs, three of which correspond to TRX types present in eukaryotes: TRX m, TRX x and TRX y. The fourth TRX (sll1057) does not correspond to any TRX type and must have been lost in eukaryotes (data not shown). The multiplicity of TRX genes in Arabidopsis raises numerous questions: are these TRXs redundant or do they present some specificities? The comparison with the situation in Chlamydomonas provides some clues concerning this question. The number of TRX genes appears much more limited in Chlamydomonas. This is especially striking for h-type TRXs for which eight isoforms are present in Arabidopsis while only two are found in Chlamydomonas. This is consistent with the fact that several functions of TRX h are specific to higher plants (germination, self-incompatibility). TRX h is also known to function in oxidative stress defenses through reduction of PRX (Verdoucq et al. 1999; Rouhier et al. 2001) TRX h has also been shown to be one of the major proteins in rice phloem sap (Ishiwatari et al. 1995), leading to the suggestion that TRXs might participate in longrange signaling in plants, another function that is probably not required in Chlamydomonas. All TRX types present in Arabidopsis are also found in Chlamydomonas but with fewer isoforms for each type. The results obtained for each TRX by analysis of the genomic data are summarized in Table 1, which includes putative subcellular localizations and EST-abundance data. The

206

Figure 1. The thioredoxin family. Phylogenetic tree of the TRX families in Chlamydomonas reinhardtii, Arabidopsis thaliana and Synechocystis PCC6803. The unrooted tree was constructed with the Clustal X program. Gaps were excluded. Accession numbers: Cr: Chlamydomonas reinhardtii h1, P80028; h2, AY184797; o, AY184798; m, P23400; f1, AY184800; f 2, AV622215; x, AY184799; y, AY184796; At: Arabidopsis thaliana h1, Z14084; h2, Z35475; h3, Z35474; h4, Z35473; h5, Z35476; h7, AAD39316; h8, AAG52561; h9, AAG51342; o1, AAC12840; o2, AF396650; m1, O48737; m2, Q9SEU8; m3, Q9SEU7; m4, Q9SEU6; f1, Q9XFH8; f2, Q9XFH9; x, AAF15952; y1, AAF04439 corrected for intron splicing; y2, AAM91085 corrected with EST AY128276; Synechocystis PCC6803 slr0623, slr1139 and slr0233.

Table 1. The thioredoxin family in Chlamydomonas reinhardtii Genewise

Green genie

Contig ber

num-

Polypeptide length (ami-

Putative subcellular localization

Total number of EST

no acids) CrTRXh1

2405.1.1

2405.0

20021010.8550.1 113

C

69

CrTRXh2 CrTRXo

623.NO 1091.8.1

623.NO 1091.1

20021010.4921.1 109 20021010.550.2 150

C M

11 4

CrTRXm

929.NO

929.2

20021010.3153.2 140

P

86 21

20021010.6554.2 CrTRXx

217.15.1 (partial)

217.1 (partial)

20021010.1360.1 145

P

CrTRXy

54.NO

54.NO

20021010.7432.1 152

P

23

CrTRXf1

1839.8.1 (partial)

1839.0 (partial)

20021010.5798.1 173

P

60

CrTRXf2

1759.NO

1759.1

DLC14 DLC16

631.35.1 (wrong splicing) 631.NO 15.21.1 15.NO

20021010.8994.2 199

P

15

20021010.8730.1 129 20021010.3210.1 156

FL FL

19 13

Genewise and greengenie correspond to gene predictions in Chlamydomonas reinhardtii genome version 1.0 (http://genome.jgi-psf.org/ chlre1). The first number corresponds to the scaffold number and is followed by NO when the presence of the gene is not predicted. Partial indicates a truncation of the predicted coding sequence. Wrong splicing refers to splicing errors in the predicted coding sequence based on cDNA or EST data. Contig accession numbers deduced from assembly of EST sequences available at (http:// www.biology.duke.edu/chlamy_genome/). The putative subcellular localizations are deduced from those demonstrated experimentally for TRXs of the same type in Arabidopsis thaliana. C: cytosolic; M: mitochondrial; P: plastidial; FL: flagellar protein. Total number of EST last estimated on 1 November 2003.

207 subcellular localization of at least one protein of each TRX type has been determined experimentally using Arabidopsis sequences (Laloi et al. 2001; Collin et al. 2003 and unpublished results). These studies revealed that TRX h and TRX o are localized in the cytosol and mitochondria respectively while all the other TRX types are targeted to the chloroplast. The number of EST sequences available for each TRX gene can be used as a rough estimation of the expression level of the corresponding genes. The three major TRXs in Chlamydomonas are TRX h1, TRX m and TRX f1, the TRX types that had been identified using standard biochemical approaches. The other TRX types are expressed at much lower levels. However, the analysis of the distribution of EST sequences in the different libraries suggests that some of these genes are expressed in specific stress conditions (Lemaire et al. 2003b). For example, ESTs corresponding to Chlamydomonas TRX x are overrepresented in stress-II libraries that include oxidative-stress treatments. This is consistent with the measured efficiency of TRX x as an electron donor to chloroplastic 2-cys peroxiredoxin (Collin et al. 2003). For f- and h-type TRXs, two genes are present in Chlamydomonas for each type but the relative expression level of each isoform is different. TRXs f1 and h1 are expressed at much higher levels than TRX f2 and h2, respectively. This suggests that the two isoforms of each type might not be equivalent. Indeed, the ESTs corresponding to TRX h2 are overrepresented in the stress-II libraries, while it is not the case for TRX h1 (Lemaire et al. 2003b). This suggests that TRX h2 might be involved in the reduction of cytosolic PRXs. A recent breakthrough in our knowledge of TRX functions in the chloroplast came from the development of new strategies. These techniques are based on the fact that a single cysteine mutant of TRX, where the second cysteine of the active site has been replaced by serine, is able to form stable mixed disulfides with target proteins. We have used this strategy to identify Chlamydomonas 2-cys PRX as a target of TRXs in the chloroplast (Goyer et al. 2002). A monocysteinic mutant TRX was immobilized on a column and loaded with Chlamydomonas total extracts. The bound proteins could then be eluted by dithiothreitol (DTT). The number of known TRX targets recently increased due to the combination of this technique with

proteomic approaches. In the chloroplast it has risen up to 37 distinct proteins (Motohashi et al. 2001; Balmer et al. 2003). A modified approach recently led to the identification of 23 TRX targets in the starchy endosperm of wheat seeds (Wong et al. 2003). Initially chloroplastic TRXs were thought to be involved in the control of carbon metabolism by light and more recently in the scavenging of ROS by way of PRXs. The new targets suggest a broader range of functions including translation, protein assembly/folding, vitamin biosynthesis, DNA replication/transcription, isoprenoid biosynthesis, plastid division, tetrapyrrole biosynthesis or protein degradation.

The glutaredoxin family Glutaredoxins are ubiquitous oxidoreductases of approximately 12 kDa, like thioredoxins. The major difference between the two redox systems is that GRXs are reduced by glutathione (GSH) instead of a specific reductase in the case of TRXs. GRXs were much less extensively studied than TRXs. GRXs were originally identified as alternative hydrogen donors to RNR in E. coli TRX mutants (Holmgren 1976). The active site of classical GRXs consists of a dithiol/disulfide in the conserved sequence CPYC. The midpoint redox potential of this disulfide is less electronegative (approx. )200 mV) than the TRX active site disulfide ()280 mV). GRXs are able to reduce disulfides but also GSH-mixed disulfides. The latter reaction is called deglutathionylation and can be performed by GRX but not by TRX (Jung and Thomas 1996; Nulton-Persson et al. 2003). Glutathionylation of cysteine residues corresponds to the formation of a mixed disulfide between GSH and the thiol group of cysteine. This post-translational modification has been shown to regulate the activity of HIV-1 protease (Davis et al. 1997), human a ketoglutarate dehydrogenase (NultonPersson et al. 2003), E coli 3¢ PAPS reductase (Lillig et al. 2003) and the polymerization of actin in human cells (Wang et al. 2003). In plants, very little is known on these regulations but it has recently been shown that two Calvin-cycle enzymes are glutathionylated in Arabidopsis (Ito et al. 2003). GRXs play a major role in the response to oxidative stress and their expression is induced in presence of oxidants (Prieto-Alamo et al. 2000;

208 Grant 2001). GRXs have been shown to possess several activities including dehydroascorbate reductase (Park and Levine 1996; Washburn and Wells 1999), GST (Collinson and Grant 2003) and glutathione peroxidase activity (Collinson et al. 2002). They are also able to reduce certain types of PRXs (Rouhier et al. 2001, 2002a) and they participate in the regulation of several transcription factors related to oxidative stress signaling in mammals (reviewed in Rouhier et al. 2002b). E. coli contains three distinct GRXs. GRX1 and GRX3 are classical CPYC GRXs, while GRX2 is an atypical GRX of 24 kDa (Xia et al. 2001). The latter is in fact a GST-like protein containing a GRX CPYC active site. In S. cerevisiae five distinct GRXs have been identified. GRXs 1 and 2 are classical CPYC GRX while GRX3, GRX4 and GRX5 belong to a different type and possess a CGFS active site (Belli et al. 2002). GRX3 and GRX4 are cytosolic proteins and GRX5 is mitochondrial. Deletion of GRX5 leads to the loss of iron–sulfur cluster assembly in yeast mitochondria (Rodriguez-Manzaneque et al. 2002).

Contrary to TRXs, very little information is available on GRXs in photosynthetic organisms despite the availability of cDNA sequences for many years (Minakuchi et al. 1994; Szederkenyi et al. 1997). GRXs are, like TRXs and PRXs, major proteins of the phloem sap (Ishiwatari et al. 1995; Balachandran et al. 1997; Szederkenyi et al. 1997). The first biochemical characterization in plants was recently performed on poplar GRX. This revealed that GRXs are able to reduce a new type of poplar PRX (Rouhier et al. 2001, 2002a). We have recently found that Arabidopsis contains 30 different GRXs while only 6 isoforms exist in Chlamydomonas (Lemaire 2004). A phylogenetic tree of the GRX families present in Chlamydomonas, Arabidopsis, and Synechocystis is presented on Figure 2. GRXs from E. coli and S. cerevisiae have been included as references. Sequences homologous to GST-like E. coli GRX2 were not included. GRX-like sequences corresponding to GRX fused to other unrelated domains, like the well-characterized 5¢ adenylylsulfate reductase from Arabidopsis (Bick et al. 1998), were also

Figure 2. The glutaredoxin family. Phylogenetic tree of the glutaredoxin family in Chlamydomonas reinhardtii, Arabidopsis thaliana and Synechocystis PCC6803 (reproduced from Lemaire 2004). The unrooted tree was constructed with the Clustal X program. Gaps were excluded. Sequences from E. coli and S. cerevisiae were included for comparison purposes. Accession numbers: At: Arabidopsis thaliana as in Table 1, the names correspond to the locus accession numbers; Chlamydomonas reinhardtii (Cr) as in Table 2; Escherichia coli (Ec) GRX1, P00277; GRX3, P37687; Saccharomyces cerevisiae (Sc) GRX1, S19363; GRX2, P17695; GRX3, NP_010383; GRX4, NP_011101; GRX5, NP_015266; Synechocystis PCC6803 slr1562, slr1846 and ssr2061.

209 Table 2. The glutaredoxin family in Chlamydomonas reinhardtii GRX type

Gene-

Green

wise

genie

Contig number

Polypeptide

Putative

Total number

length

subcellular

of EST

(amino acids)

localization

CrGRX1

CPYC

6.74.1

6.16

20020630.7451

128

C

20

CrGRX2

CPYC

1116.9.1

1166.1

20021010.7553

107

C

11

CrGRX3

CGFS Group III

115.16

115.11

20021010.7618

148

M or P

42

CrGRX4

CGFS Group I

620.15.1

620.3

20020630.4199.4

234

C

CrGRX5

CGFS Group II

344.7.1

344.NO

NO

?

M or P

CrGRX6

CGFS Group IV 29.77.1

29.29

20021010.6117

343

P

4 0 14

Refer to the legend of Table 1 for details on genewise, greengenie and contig number columns. Uncertainties in the prediction of intron/ exon boundaries on the 5¢ part of CrGRX5 gene do not allow deducing the complete amino acid sequence of the corresponding polypeptide. Putative subcellular localizations are based on the predictions performed on homologous Arabidopsis sequences using a combination of several programs: Predotar, TargetP, Psort and iPsort (links available at www.expasy.ch) (Lemaire 2004). C: cytosolic; M: mitochondrial; P: plastidial. Total number of EST last estimated on 1 November 2003.

excluded. The accession numbers, GRX types, polypeptide length, putative subcellular localizations and EST abundance of the six glutaredoxins present in Chlamydomonas are presented in Table 2. Prediction programs are not well suited for Chlamydomonas sequences while they are quite accurate for Arabidopsis. Thus, the proposed subcellular localizations are based on the predictions performed on homologous GRXs from Arabidopsis using a combination of several programs: Predotar, TargetP, Psort and iPsort (links available at www.expasy.ch) (Lemaire 2004). The prokaryotic and eukaryotic sequences of CPYC-type GRXs form distinct clusters. The CPYC group contains six Arabidopsis GRXs, only four of which correspond to classical GRXs with a CPYC active site sequence. The two other proteins (At4g28730 and At2g20270), which are predicted to be targeted to the chloroplast, must have evolved recently and may have lost the active site sequence. Two Arabidopsis CPYC GRXs (At1g77370 and At5g20500) contain an N-terminal sequence resembling a secretion signal. These GRXs might thus represent the GRX isoforms known to be present in the phloem sap. The two remaining Arabidopsis proteins are predicted to be localized in the cytosol and are homologous to the two Chlamydomonas CPYC GRXs (Table 2). The CC type was recently identified and corresponds to proteins containing a CCXC or CCXS active site (Lemaire 2004). This new type appears to be specific of higher plant species and contains 20 members in Arabidopsis. A large majority of these proteins (18 out of 20) are predicted to be localized

in the cytosol while the two remaining sequences might be targeted, though no clear prediction could be established. Although CC-type GRXs exhibit characteristic sequence features of GRXs, their biochemical properties will have to be determined in vitro. The CGFS type contains four GRXs, both in Chlamydomonas and Arabidopsis. Each Chlamydomonas GRX clusters with a homologous protein in Arabidopsis, thereby defining four distinct subgroups (Figure 2). Group I corresponds to multipartite proteins with several GRX and/or TRX domains. These proteins are predicted to be localized in the cytosol. This is consistent with the presence in the same group of ScGRX3 and ScGRX4, two proteins which are not localized in yeast mitochondria (Rodriguez-Manzaneque et al. 2002). Group II corresponds to proteins that are most probably localized in the mitochondria. Prediction programs suggest a mitochondrial or chloroplastic localization but the presence in this group of ScGRX5, a yeast mitochondrial GRX, strongly suggests a mitochondrial localization. The absence of EST sequences corresponding to CrGRX5 is also an argument in favor of a mitochondrial targeting (Table 2). Indeed, in the case of the TRX family, mitochondrial isoforms were also found to be expressed at very low levels (Lemaire et al. 2003b). Group III contains GRXs predicted to be targeted to the chloroplast or mitochondria with similar probabilities. However, the presence of a Synechocystis GRX in the same group is rather indicative of a chloroplastic localization for these proteins. Synechocystis only

210 contains three GRXs: two CPYC prokaryotic sequences that cluster with E. coli GRXs and this group II CGFS GRX. CrGRX3 is the most highly expressed GRX in Chlamydomonas, much higher than the so-called classical CPYC GRXs (Table 2). This indicates that these proteins probably have important functions in oxygenic photosynthetic organisms that remain to be examined. GRXs in group IV are unequivocally predicted to be targeted to the chloroplast. These proteins are longer than GRXs found in group III, the C-terminal GRX domain being preceded by an N-terminal domain sharing strong homology with the Synechocystis slr1035 sequence of unknown function. Group IV GRXs might thus represent a fusion between a CGFS GRX and a homologue of slr1035, an argument in favor of a chloroplastic localization. All the subcellular localizations suggested here are based on prediction programs and will have to be confirmed experimentally. As in the case of TRXs, the GRX family appears much more limited in Chlamydomonas with only six isoforms compared to 30 in Arabidopsis. Contrary to TRXs, where all TRX types are present but with fewer isoforms for each type, the difference in the GRX families from both organisms is essentially due to the absence of the CC type in the green alga. CGFS GRXs were initially thought to be able to reduce GSH-mixed disulfides through a monothiol mechanism. The GSH-mixed disulfide is cleaved by GRX and, in turn, GSH forms a mixed disulfide with the active site cysteine of GRX. This mixed disulfide is cleaved by a second GSH to yield oxidized glutathione (GSSG) and reduced GRX. This mechanism was deduced from the analysis of monocysteinic mutants of CPYC GRXs, with a CPYS active site. The first biochemical characterization of a CGFS type GRX was recently performed on yeast GRX5 and revealed unexpected properties (Tamarit et al. 2003). ScGRX5 was shown to contain a disulfide between the active site cysteine (Cys60) and a second cysteine residue (Cys117). Apparently, once a mixed disulfide is formed between Cys60 and GSH, it is not attacked by a second GSH molecule as in the monothiol mechanism. Instead, it appears that the mixed disulfide is attacked by Cys117 which forms a disulfide bond with Cys60, thereby releasing reduced GSH. Interestingly, GRX5 disulfide is not efficiently reduced by GSH while TRX appears

more efficient. This provides an unexpected link between the TRX and GRX systems. Many CGFS GRXs present in Arabidopsis and Chlamydomonas also contain a second cysteine in the same position as Cys117 in ScGRX5 (Lemaire 2004). In Chlamydomonas, only GRX6 is lacking this additional cysteine while it is present in GRX3, GRX4 and GRX5. Future studies should be aimed at characterizing the CGFS GRXs present in the different subcellular compartments of photosynthetic eukaryotic cells. It will be particularly interesting to determine if these proteins might also contain a disulfide bond and to assess the possible reduction of this disulfide by TRX. In the latter case, the determination of the specificity of the different TRX types for reduction of GRX disulfide may shade light on the possible interactions between the TRX and GRX systems. Recently, a poplar TRX h was suggested to be reduced by GRX (Gelhaye et al. 2003) and the activity of human TRX was reported to be modulated by glutathionylation (Casagrande et al. 2002). These data are additional strong indications that the cross-talk between both redox systems deserves further analysis.

The PDI family Protein disulfide isomerases constitute a large family of enzymes with two interrelated activities. They exhibit disulfide oxidoreductase activity allowing the reduction, the oxidation or the isomerization of disulfides but they are also molecular chaperones which assist the folding of proteins (reviewed in Ferrari and So¨ling 1999). Contrary to TRXs and GRXs which are ubiquitous, PDIs are only found in eukaryotes and are usually located in the endoplasmic reticulum (ER). PDIs have a modular structure with different combinations of TRX domains called the a and b domains. The PDI active site WCGHC is only present in the a (or catalytic) domains while the b domains have a TRX-like folding but no sequence homology with the a domains. Consequently, the b domains cannot usually be recognized by alignment programs such as blastP. These b domains have an important role in the specificity of binding to substrate proteins (Freedman et al. 2002). Typical PDIs contain four TRX domains and follow the a–b–b¢–a¢ architecture. Several proteins

211 homologous to classical PDI but with distinct domain distribution have been identified in mammals. At least six families of proteins have been defined: PDI, ERp57, ERp72, P5, PDI-R and PDI-D. A detailed analysis of human genome recently allowed the identification of four additional PDI-like proteins (Clissold and Bicknell 2003). In yeast deletion of PDI1 is lethal (Scherens et al. 1991), its essential role being the isomerization of disulfides (Laboissie`re et al. 1995). While PDIs have been extensively studied in mammals, very little is known about these proteins in photosynthetic organisms. One of the most studied PDI from a photosynthetic organism is the Chlamydomonas Rb60 protein. Rb60 is part of a high molecular multiprotein complex involved in the control of psbA translation by light. Rb60 was originally identified, along with Rb38, Rb47 and Rb55, as part of the psbA 5¢-PC complex which specifically binds to the 5¢ UTR of the chloroplastic psbA mRNA (Danon and Mayfield 1991). The level of bonding of the complex is correlated with light-enhanced translation of psbA mRNA. The binding of 5¢-PC is mediated by Rb47 which presents a high homology with poly-A binding proteins (Yohn et al. 1998). The light regulation of 5¢PC operates through Rb60 by several mechanisms. ADP-dependent phosphorylation of Rb60, at ADP levels only found in the dark, appears to inactivate psbA 5¢PC (Danon and Mayfield 1994a). Additionally, Rb60 appears to be activated by reduction of vicinal dithiols, possibly by thioredoxin (Danon and Mayfield 1994b) but a counteracting oxidative component also appears to be involved (Trebitsh et al. 2000). However, this thiol regulation requires an initial step mediated by a light signal transduction pathway originating from the plastoquinone pool (Trebitsh and Danon 2001). The cloning of Rb60 revealed that it is a classical PDI (Kim and Mayfield, 1997). PDIs usually contain an N-terminal secretion signal for export to the ER and a KDEL retention signal. Surprisingly, the Rb60 amino-acid sequence also exhibits these features while its function strongly suggests a chloroplastic localization. Subcellular fractionation experiments have indeed confirmed the plastidic localization of Rb60 but it is still not known if it might also be present in the ER (Trebitsh et al. 2001). Recently, the knock-out of three PDI genes in the moss Physcomitrella patens

indicated that these proteins are not essential for viability (Meiri et al. 2002). However, to date, the number and diversity of PDIs in photosynthetic eukaryotes remains to be examined. We have analyzed the genomes of Arabidopsis and Chlamydomonas in order to identify all members of the PDI family. This analysis was restricted to PDIs containing at least one WCGHC active site with a tolerance for one amino-acid substitution within this sequence. Arabidopsis contains nine PDIs while only four are found in Chlamydomonas (Figure 3). If no restrictions are applied for the presence of one active site WCGHC sequence, the number of PDIs in Arabidopsis increases to 16, while only one additional PDI is found in Chlamydomonas (data not shown). Thus, as already observed for TRXand GRX-gene families, the multiplicity is more limited in the green alga. The phylogenetic tree allows the distinction of different subgroups which correspond in most cases to the modular architecture of the proteins and to PDI types already described in human (Figures 3 and 4 Tables 3 and 4). PDIs from human, alfalfa and Dictyostelium discoideum have been included as references. The first group corresponds to classical PDIs containing a typical a–b–b¢–a¢ domain (Figure 5). This group includes Rb60, four different Arabidopsis PDIs and four human PDIs. Three distinct subgroups can be distinguished. The first one contains human PDI and PDIp, the pancreatic specific PDI in human. Human PDI contains a C-terminal acidic region termed the c domain and has an a–b–b¢–a¢–c domain distribution. The second subgroup contains two Arabidopsis proteins (At1g77510 and At1g21750) and the human ERp57. These proteins have an a–b–b¢–a¢ structure but lack the c domain present in human PDI. ERp57 exhibits a lower redox activity than PDI and is part of a complex with calnexin and calreticulin involved in the maturation of glycosylated proteins (reviewed in High et al. 2000). This complex is involved in the folding of class-I major histocompatibility complex (reviewed in Bouvier, 2003). The third subgroup includes two Arabidopsis proteins (At5g60640 and At3g54960), Rb60 and human ERp72. ERp72 contains, like human PDI, a c domain located at the N-terminal end of the protein. An additional a domain is also present before the standard a–b–b¢–a¢ domains. The domain distribution of ERp72 is c–a0–a–b–b¢–a¢.

212

Figure 3. The protein disulfide isomerase family. Phylogenetic tree of the PDI family in Chlamydomonas reinhardtii and Arabidopsis thaliana. The unrooted tree was constructed with the Clustal X program. Gaps were excluded. Sequences from human, alfalfa (Medicago sativa) and Dictyostelium discoideum were included as references. Accession numbers: At: Arabidopsis thaliana as in Table 4, the names correspond to the locus accession numbers; Chlamydomonas reinhardtii (Cr) as in Table 3; Homo sapiens (Hs) PDI, P07237; ERp57, P30101; ERp72, P13667; P5, D49489; Medicago sativa (Ms) PDI-Da, P38661; Dictyostelium discoideum (Dd) PDI-Da, AAB86685.

Figure 4. The thioredoxin superfamily in Chlamydomonas reinhardtii. Phylogenetic tree of the TRX superfamily in Chlamydomonas reinhardtii. The unrooted tree was constructed with the Clustal X program. Gaps were excluded. Accession numbers as in Figure 1 and Tables 1–3.

213 Table 3. The protein disulfide isomerase family in Chlamydomonas reinhardtii Modular

Genewise

Green genie

Contig number

structure

CrPDI1

c–a–b–b¢–a¢ 436.19.1

436.8

(Rb60)

20021010.1793.1

Length

Four last

Putative

Total

(amino

amino

sorting

number

acids)

acids

signal

of EST

532

KDEL

S (but P

37

20021010.2858.1

experimental)

(partial) CrPDI2 CrPDI3

a–D A

CrPDI4

a0–a–b–c

2212.1.1

2212.0

(partial)

(wrong splicing)

1769.26.1

1769.1

(partial)

(wrong splicing)

440.13.1 (gap)

440.5 (gap + wrong

20021010.8415.1

286

EDEE

S?

33

20021010.4388.1

153

TTEA

S

11

20021010.8775.1 20021010.2496.1

484

NEEL

?

2

splicing) Refer to the legend of Table 1 for details on genewise, greengenie and contig number columns. Gap indicates the presence of a gap in the scaffold sequence. CrPDI4 coding sequence was reconstructed using a combination of EST and genome sequences. Putative sorting signals are based on the predictions performed using a combination of several programs: Predotar, TargetP, Psort and iPsort (links available at www.expasy.ch). P: plastidial; S: presence of an N-terminal secretion signal. For Rb60 prediction programs predict the presence of an N-terminal secretion sequence while the protein was shown to be present in the chloroplast (Trebitsh et al. 2001). Total number of EST last estimated on 1 November 2003.

Table 4. The protein disulfide isomerase family in Arabidopsis Locus

Modular structure

Polypeptide length

Four last amino acids

Putative sorting signal

(amino acids) At1g04980

a0–a–b–c

443

KDDL

S

At1g07960

a

146

DKEL

S

At1g21750

a–b–b¢–a¢

501

KDEL

S

At1g35620

a–b–b¢

440

KKED

S

At1g77510

a–b–b¢–a¢

508

KDEL

S

At2g32920

a0–a–b–c

440

KDEL

S

At2g47470

a0–a–D

361

VASS

S

At3g54960 At5g60640

c–a–b–b¢–a¢ c–a–b–b¢–a¢

579 597

KDEL KDEL

S S

Putative sorting signals are based on the predictions performed as described in the legend of Table 1. S: presence of an N-terminal secretion signal.

Rb60 and the two Arabidopsis PDIs of this subgroup also contain an acidic N-terminal domain but not the additional a domain. Their domain distribution is thus c–a–b–b¢–a¢. The presence of the c domain in these proteins might explain the presence of ERp72 in the same subgroup. ERp72 can complement yeast PDI-deficient strain (Gunther et al. 1993) and exhibits redox and isomerase activities (Rupp et al. 1994). ERp72 is part of a large ER complex of 10 molecular chaperones also including classical PDI and P5 PDI (Meunier et al. 2002). Rb60 is apparently the

only ‘true’ PDI in Chlamydomonas while four are present in Arabidopsis. These five PDIs are predicted to be localized in the ER since they possess N-terminal secretion peptides and they also all have a C-terminal KDEL retention signal, allowing retention of the protein in the lumen of the ER. However, as mentioned above, Rb60 is present in the chloroplast (Trebitsh et al. 2001). Thus, it is possible that some of the four Arabidopsis PDIs might also be present in the chloroplast, such as At5g60640 and At3g54960 which are more closely related to Rb60 than the two other Arabidopsis

214

Figure 5. Schematic representation of domain distributions in different types of PDI. Domains a0, a and a¢ (grey boxes) represent redox active thioredoxin domains. Domains b and b¢ (black boxes) represent redox inactive thioredoxin domains. D (white boxes) represents the a-helical domain specific of PDI-D family and c (white boxes) is a higly acidic region. The scale is arbitrary, thus the size of the boxes is not proportional to the length of the corresponding amino acid sequences. Species abbreviations: At: Arabidopsis thaliana; Cr: Chlamydomonas reinhardtii; Dd: Dictyostelium discoideum; Hs: Homo sapiens; Ms: Medicago sativa. Accession numbers as in Figure 3.

PDIs. Anyway, the subcellular localization of these proteins will have to be examined carefully for potential dual targeting. A second group of PDIs, named P5, is composed of CrPDI4, At1g04980 and At2g32920 which possess an a0–a–b–c architecture (Figure 5). These proteins are homologous to the human P5 and rat CaBP1 PDIs that also exhibit the same a0–a–b–c domain distribution (Fu¨llekrug et al. 1994; Ferrari and So¨ling, 1999). Both human and rat proteins are abundant ER resident proteins. CaBP1, which is the best characterized protein in the P5 group, is able, like classical PDIs, to catalyze the reduction of insulin (Nguyen Van et al. 1993; Fu¨llekrug et al. 1994) and the refolding of denatured RNase A III (Rupp et al. 1994). A detailed biochemical analysis recently revealed that CaBP1 can complement the yeast PDI1D lethal mutation but cannot catalyze the redox independent refolding of GAPDH (Kramer et al. 2001). CaBP1 apparently lacks the general chaperone and peptide binding activities of PDI generally attributed to the C-terminal end of the a¢ domain and the b¢ domain, respectively (Cai

et al. 1994; Klappa et al. 1998). This is consistent with the a0–a–b–c architecture of this protein. CaBP1 is, however, able to functionally replace PDI in its redox/isomerase activity and can catalyze the oxidative refolding of proteins as long as one functional TRX catalytic domain is present (Kramer et al. 2001). As mentioned above, CaBP1 belongs to a complex of 10 molecular chaperones together with PDI and ERp72 (Meunier et al. 2002). The role of the b domain present in all PDIs of the P5 type remains to be determined. The P5 type PDIs found in Arabidopsis and Chlamydomonas all contain a C-terminal ER retention signal while the presence of a putative N-terminal secretion signal is predicted for Arabidopsis proteins and uncertain for the Chlamydomonas protein (Tables 3 and 4). However, as already mentioned prediction programs are not always well-suited for Chlamydomonas sequences. All these P5-type proteins are thus likely to be localized in the ER like their mammalian homologues. Some proteins, originally defined as P5 PDIs, such as alfalfa (Medicago sativa) PDI (Shorrosh

215 and Dixon 1992) or Dictyostelium discoideum DdPDI (Monnat et al. 1997), have been reassigned to a distinct subfamily named PDI-D (Ferrari and So¨ling 1999). These PDIs are characterized by a long C-terminal a-helical domain of approximately 100 amino acids named the D domain. Proteins in the PDI-D family can either be redox active (PDIDa) like the alfalfa and Dictyostelium PDI-D or redox inactive (PDI-Db), like the human ERp28 (Ferrari et al. 1998) or rat ERp29 (Demmer et al. 1997; Mkrtchian et al. 1998). PDI-Db are ER resident proteins exhibiting a b–D structure. Genes encoding homologues of PDI-Db could not be identified in the genomes of Chlamydomonas and Arabidopsis (data not shown). However, this absence might be linked to the fact that the b domains are hardly detectable by standard BLAST alignments as already mentioned. PDI-Da from alfalfa and Dictyostelium have an a0–a–D structure. A single homologue of PDI-Da is found in Arabidopsis and Chlamydomonas. Both proteins possess the characteristic D domain but differ in their domain distribution. While Arabidopsis PDID exhibits the a0–a–D architecture also found in other member of PDI-Da family, Chlamydomonas PDI-D is lacking one TRX domain and exhibits an a–D organization (Tables 3 and 4, Figure 5). Indeed, the algal protein is approximately 100 amino acids shorter than its Arabidopsis counterpart, a size similar to the size of a TRX domain. Both proteins are predicted to contain an N-terminal secretion signal. While the C-terminal EDEE sequence found in CrPDI2 might function as an ER retention signal, it is not the case for the VASS sequence found in At2g47470. Similarly, no ER retention signal is present in PDI-Da from alfalfa or Dictyostelium while these proteins are known to be located in the ER (Shorrosh and Dixon 1992; Monnat et al. 1997). However, it was recently shown that the C-terminal part of the D domain is responsible for the ER localization of PDI-D in Dictyostelium (Monnat et al. 2000). The PDI-Da proteins found in Chlamydomonas and Arabidopsis also possess a C-terminal D domain which might also allow their import in the ER. An additional group corresponds to At1g07960 and CrPDI3. These proteins contain a single a domain with a WCGHC active site in Chlamydomonas and WCKHC in Arabidopsis (Figure 5). We thus propose to name this family PDI-A. The presence in other organisms of PDIs with similar

architecture has not been reported. However, analysis of EST databases indicates that homologues with high similarity are found in many other photosynthetic eukaryotes (data not shown). The PDI-A family thus appears to represent a new PDI family specific to photosynthetic eukaryotes. Finally the At1g35620 PDI appears to have no counterpart in Chlamydomonas reinhardtii or mammalian genomes. This PDI exhibits an a–b–b¢ structure which was recently reported to be present in the KIAA0573 human protein (Clissold and Bicknell 2003). However, the similarity between both proteins is quite low and phylogenetic analyses indicate that they cannot be considered as being part of the same subfamily (data not shown). Analyses of EST databases indicate that homologues of this protein are found in other higher plants. This suggests that this PDI might be part of a higher plant specific subfamily. The human ERp28 and PDI-R proteins found in mammals do not apparently have counterparts in photosynthetic eukaryotes. In contrast, the ERdj5 protein which is a DnaJ/PDI hybrid protein recently characterized is also present in Chlamydomonas and Arabidopsis (data not shown). The analysis of EST abundance in Chlamydomonas indicates that Rb60 and the PDI-Da CrPDI2 are the two most highly expressed PDIs. The PDI-A CrPDI3 and especially the P5 CrPDI4 are expressed at much lower levels. Many studies have been performed on Rb60 but its function still needs to be clarified. It will be especially interesting to determine which type of TRX is involved in the regulation of psbA translation by light through the reduction of Rb60. All the newly identified PDIs will have to be characterized biochemically and their subcellular localization will have to be determined experimentally. Their associations in complexes with other molecular chaperones, already shown for some of their mammalian homologues, will also have to be analyzed. Finally, it will be especially interesting to study the function of members of the PDI-A type, apparently specific to photosynthetic eukaryotes.

Chlamydomonas: a model organism to study the thioredoxin superfamily Analysis of the genome of Chlamydomonas reinhardtii indicates that the TRX, GRX and PDI

216 families contain 10, 6 and 4 members, respectively (Figure 4). We do not exclude the possibility that genes encoding additional members of this family are present in the genome. For example, scaffolds 922 and 22 (in Chlamydomonas reinhardtii genome version 1.0 from DOE Joint Genome Institute) contain nucleotide sequences that may encode polypeptides corresponding to fragments of PDI. No genes are predicted to be present in the corresponding regions but it is also the case for CrTRXh2 and CrTRXy while cDNAs are found for these sequences (Table 1). However, in the case of scaffolds 922 and 22, the absence or very low abundance of EST in the regions encoding fragments of PDI does not allow to confirm the presence of a PDI gene. Many errors in prediction of intron/exon boundaries by both genewise and green genie programs have been encountered and often led to errors and truncation in the translated polypeptides (Tables 1 and 3). Some members of the studied gene families might have been missed due to the presence of such errors. Additionally, many gaps are present in this first version of the Chlamydomonas genome and could contain genes corresponding to members of the TRX superfamily. Efforts are still in progress to sequence these gaps and the availability of the second version of the genome might allow identifying new members of the superfamily. The criteria used to determine if a protein is part of a gene family can significantly change the result of the genome analysis. These criteria are subject to changes with time depending on the progress made in the analysis of new members of the family. Many proteins containing one or several TRX domains are not currently considered as members of the TRX, GRX or PDI families. This was also initially the case for TRX y which is now considered as a true TRX (Lemaire et al. 2003a). A TRX from poplar containing a CXXS active site sequence was recently characterized biochemically (Gelhaye et al. 2003). Homologues with similarly altered active sites are present in other photosynthetic organisms. However, in the present study, these TRXs were considered as TRX-like proteins rather than true TRX. These examples illustrate the importance of the criteria used to determine the boundaries of the gene families. However, the same criteria being used for analysis of the different genomes, comparison of the gene families between different organisms is possible. The TRX,

Table 5. Members of the thioredoxin superfamily identified in Synechocystis, Chlamydomonas and Arabidopsis Organism

TRX

GRX

PDI

genes

genes

genes

Total

4

3

0

Chlamydomonas

10

6

4 (5)

20 (21)

Arabidopsis

19

30

9 (16)

58 (65)

Synechocystis

7

The total number of genes identified for the TRX, GRX and PDI families is presented. Numbers in parentheses correspond to the total number of PDI genes identified in the corresponding genome if no restrictions are applied for the presence of one active site WCGHC sequence (see text).

GRX and PDI families in Chlamydomonas contain 20 members compared to 58 in Arabidopsis (Table 5). This large discrepancy is mainly due to the GRX family which contains 30 members in the higher plant and only six in the green alga, essentially due to the absence of the CC type in Chlamydomonas. However, even without the GRX family, the total number of genes in Chlamydomonas still represents one-half of the genes present in Arabidopsis. Multiplicity thus appears much lower in Chlamydomonas. This is consistent with the fact that some of the functions described for these proteins are specific to higher plants. Consequently, if these functions are fulfilled by specialized proteins, these proteins will not be present in Chlamydomonas. The inverse situation is also found, as Chlamydomonas flagellar specific TRXs have no counterparts in higher plants. The redundancy of members of the TRX superfamily is a major problem for reverse genetic studies in Arabidopsis. The limited number of TRX, GRX and PDI isoforms in Chlamydomonas makes it a good model to unravel basic functions of these proteins in photosynthetic eukaryotic cells. Additionally, Chlamydomonas is a facultative autotroph, an exceptional feature for a photosynthetic eukaryote which could prove useful for the analysis of the functions of chloroplastic proteins of the superfamily. One of the advantages of Arabidopsis is the availability of large collections of knock out mutants. Similar collections will not be available in Chlamydomonas due to problems of strain storage and maintenance. However, the recent advances in the field of gene silencing, and especially the RNA interference (RNAi) technique, may provide a very

217 good alternative. RNA interference allows, after introduction of a transgene expressing a double stranded RNA homologous to a given gene, to block the expression of this gene through specific degradation of mRNA (Cerutti 2003). This technique can also be used to simultaneously inactivate the expression of highly homologous genes, a feature that could be useful for the analysis of the TRX superfamily. Chlamydomonas which can be easily transformed by electroporation and grows rapidly appears well suited for RNAi. A recent study allowed the inactivation of two Chlamydomonas rhodopsin genes by RNAi (Sineshchekov et al. 2002). The RNAi strategy thus appears to be applicable in Chlamydomonas. However, its efficiency in the alga is quite low and renders the isolation of knock-down mutants difficult without screening, i.e., without any presumptions on the expected phenotype (S.D. Lemaire and M. MiginiacMaslow, unpublished results). The development of an efficient RNAi strategy allowing direct selection of mutants thus appears as a prerequisite for the development of reverse genetic approaches in Chlamydomonas. The development of new techniques combined with proteomics allowed identifying an unexpectedly high number of new targets of TRX in chloroplasts (Motohashi et al. 2001; Balmer et al. 2003) and seeds (Wong et al. 2003). These new targets are involved in many different processes on which the influence of TRX regulation will have to be examined. These studies mainly focused on the targets of TRX f and m in the chloroplast and TRX h in seeds. Similar approaches with other TRX types or with different subcellular fractions should considerably increase the number of known TRX targets and TRX-regulated processes. They could also be used to identify the proteins interacting with GRXs or PDIs. The availability of Chlamydomonas genome will now allow using this kind of approach to unravel basic functions of TRX superfamily members in the cell of photosynthetic eukaryotes. The studies on members of the TRX superfamily in photosynthetic organisms have been mainly focused on TRXs while little is known on the function of GRXs and PDIs in these organisms. This is especially detrimental in view of recent studies suggesting a complex cross-talk between these proteins. Indeed, some GRXs appear to be reduced by TRX and not GSH (Tamarit et al.

2003). Inversely, a poplar TRX h was very recently shown to be reduced by GRX instead of NTR (Gelhaye et al. 2003). TRX is also likely to interact with the Rb60 PDI in Chlamydomonas (Trebitsh and Danon 2001). Moreover, the activity of human TRX has recently been shown to be modulated by glutathionylation (Casagrande et al. 2002). All these recent advances clearly indicate that the TRX, GRX and to a lesser extent PDIs are not independent redox protein families but rather constitute a complex regulatory network with cross-talk regulations. Unraveling this network will be the challenge of the oncoming years.

References Balachandran S, Xiang Y, Schobert C, Thompson GA and Lucas WJ (1997) Phloem sap proteins from Cucurbita maxima and Ricinus communis have the capacity to traffic cell to cell through plasmodesmata. Proc Natl Acad Sci USA 94: 14150–14155 Balmer Y, Koller A, del Val G, Manieri W, Schu¨rmann P and Buchanan BB (2003) Proteomics gives insight into the regulatory function of chloroplast thioredoxins. Proc Natl Acad Sci USA 100: 370–375 Belli G, Polaina J, Tamarit J, De La Torre MA, RodriguezManzaneque MT, Ros J and Herrero E (2002) Structurefunction analysis of yeast Grx5 monothiol glutaredoxin defines essential amino acids for the function of the protein. J Biol Chem 277: 37590–37596 Besse I, Wong JH, Kobrehel K and Buchanan BB (1996) Thiocalsin: a thioredoxin-linked, substrate-specific protease dependent on calcium. Proc Natl Acad Sci USA 93: 3169– 3175 Bick JA, Aslund F, Chen Y and Leustek T (1998) Glutaredoxin function for the carboxyl-terminal domain of the plant-type 5¢-adenylylsulfate reductase. Proc Natl Acad Sci USA 95: 8404–8409 Bouvier M (2003) Accessory proteins and the assembly of human class I MHC molecules: a molecular and structural perspective. Mol Immunol 39: 697–706 Bre´he´lin C, Mouaheb N, Verdoucq L, Lancelin JM and Meyer Y (2000) Characterization of determinants for the specificity of Arabidopsis thioredoxins h in yeast complementation. J Biol Chem 275: 31641–31647 Cabrillac D, Cock JM, Dumas C and Gaude T (2001) The S-locus receptor kinase is inhibited by thioredoxins and activated by pollen coat proteins. Nature 410: 220–223 Cai H, Wang CC and Tsou CL (1994) Chaperone-like activity of protein disulfide isomerase in the refolding of a protein with no disulfide bonds. J Biol Chem 269: 24550–24552 Casagrande S, Bonetto V, Fratelli M, Gianazza E, Eberini I, Massignan T, Salmona M, Chang G, Holmgren A and Ghezzi P (2002) Glutathionylation of human thioredoxin: a possible crosstalk between the glutathione and thioredoxin systems. Proc Natl Acad Sci USA 99: 9745–9749

218 Cerutti H (2003) RNA interference: traveling in the cell and gaining functions? Trends Genet 19: 39–46 Cho MJ, Wong JH, Marx C, Jiang W, Lemaux PG and Buchanan BB (1999) Overexpression of thioredoxin h leads to enhanced activity of starch debranching enzyme (pullulanase) in barley grain. Proc Natl Acad Sci USA 96: 14641–14646 Clissold PM and Bicknell R (2003) The thioredoxin-like fold: hidden domains in protein disulfide isomerases and other chaperone proteins. Bioessays 25: 603–611 Collin V, Issakidis-Bourguet E, Marchand C, Hirasawa M, Lancelin JM, Knaff DB and Miginiac-Maslow M (2003) The Arabidopsis plastidial thioredoxins: new functions and new insights into specificity. J Biol Chem 278: 23747–23752 Collinson EJ and Grant CM (2003) Role of yeast glutaredoxins as glutathione S-transferases. J Biol Chem 278: 22492–22497 Collinson EJ, Wheeler GL, Garrido EO, Avery AM, Avery SV and Grant CM (2002) The yeast glutaredoxins are active as glutathione peroxidases. J Biol Chem 277: 16712–16717 Danon A and Mayfield SP (1991) Light regulated translational activators: identification of chloroplast gene specific mRNA binding proteins. EMBO J 10: 3993–4001 Danon A and Mayfield SP (1994a) ADP-dependent phosphorylation regulates RNA-binding in vitro: implications in lightmodulated translation. EMBO J 13: 2227–2235 Danon A and Mayfield SP (1994b) Light-regulated translation of chloroplast messenger RNAs through redox potential. Science 266: 1717–1719 Davis DA, Newcomb FM, Starke DW, Ott DE, Mieyal JJ and Yarchoan R (1997) Thioltransferase (glutaredoxin) is detected within HIV-1 and can regulate the activity of glutathionylated HIV-1 protease in vitro. J Biol Chem 272: 25935–25940 Decottignies P, Schmitter JM, Jacquot JP, Dutka S, Picaud A and Gadal P (1990) Purification, characterization, and complete amino acid sequence of a thioredoxin from a green alga, Chlamydomonas reinhardtii. Arch Biochem Biophys 280: 112–121 Decottignies P, Schmitter JM, Dutka S, Jacquot JP and Miginiac-Maslow M (1991) Characterization and primary structure of a second thioredoxin from the green alga, Chlamydomonas reinhardtii. Eur J Biochem 198: 505–512 Demmer J, Zhou C and Hubbard MJ (1997) Molecular cloning of ERp29, a novel and widely expressed resident of the endoplasmic reticulum. FEBS Lett 402: 145–150 Ferrari DM and So¨ling HD (1999) The protein disulphideisomerase family: unravelling a string of folds. Biochem J 339: 1–10 Ferrari DM, Nguyen Van P, Kratzin HD and So¨ling HD (1998) ERp28, a human endoplasmic-reticulum-lumenal protein, is a member of the protein disulfide isomerase family but lacks a CXXC thioredoxin-box motif. Eur J Biochem 255: 570–579 Freedman RB, Klappa P and Ruddock LW (2002) Protein disulfide isomerases exploit synergy between catalytic and specific binding domains. EMBO Rep 3: 136–140 Fu¨llekrug J, So¨nnichsen B, Wunsch U, Arseven K, Nguyen Van P, So¨ling HD and Mieskes G (1994) CaBP1, a calcium binding protein of the thioredoxin family, is a resident KDEL protein of the ER and not of the intermediate compartment. J Cell Sci 107: 2719–2727

Gelhaye E, Rouhier N and Jacquot JP (2003) Evidence for a subgroup of thioredoxin h that require GSH/GRX for its reduction. FEBS Lett 555: 443–448 Goyer A, Hasleka˚s C, Miginiac-Maslow M, Klein U, Le Marechal P, Jacquot JP and Decottignies P (2002) Isolation and characterization of a thioredoxin-dependent peroxidase from Chlamydomonas reinhardtii. Eur J Biochem 269: 272– 282 Grant CM (2001) Role of the glutathione/glutaredoxin and thioredoxin systems in yeast growth and response to stress conditions. Mol Microbiol 39: 533–541 Gunther R, Srinivasan M, Haugejorden S, Green M, Ehbrecht IM and Kuntzel H (1993) Functional replacement of the Saccharomyces cerevisiae Trg1/Pdi1 protein by members of the mammalian protein disulfide isomerase family. J Biol Chem 268: 7728–7732 Harrison A, Sakato M, Tedford HW, Benashski SE, Patel-King RS and King SM (2002) Redox-based control of the gamma heavy chain ATPase from Chlamydomonas outer arm dynein. Cell Motil Cytoskeleton 52: 131–143 High S, Lecomte FJ, Russell SJ, Abell BM and Oliver JD (2000) Glycoprotein folding in the endoplasmic reticulum: a tale of three chaperones? FEBS Lett 476: 38–41 Holmgren A (1976) Hydrogen donor system for Escherichia coli ribonucleoside-diphosphate reductase dependent upon glutathione. Proc Natl Acad Sci USA 73: 2275–2279 Huppe HC and Buchanan BB (1989) Activation of a chloroplast type of fructose bisphosphatase from Chlamydomonas reinhardtii by light-mediated agents. Z Naturforsch 44: 487–494 Huppe HC, de Lamotte-Guery F, Jacquot J-P and Buchanan BB (1990) The ferredoxin–thioredoxin system of a green alga, Chlamydomonas reinhardtii: identification and characterization of thioredoxins and ferredoxin-thioredoxin reductase components. Planta 180: 341–351 Huppe HC, Picaud A, Buchanan BB and Miginiac-Maslow M (1991) Identification of an NADP/thioredoxin system in Chlamydomonas reinhardtii. Planta 186: 115–121 Ishiwatari Y, Honda C, Kawashima I, Nakamura S, Hirano H, Mori S, Fujiwara T, Hayashi H and Chino M (1995) Thioredoxin h is one of the major proteins in rice phloem sap. Planta 195: 456–463 Issakidis-Bourguet E, Mouaheb N, Meyer Y and MiginiacMaslow M (2001) Heterologous complementation of yeast reveals a new putative function for chloroplast m-type thioredoxin. Plant J 25: 127–135 Ito H, Iwabuchi M and Ogawa K (2003) The sugar-metabolic enzymes aldolase and triose-phosphate isomerase are targets of glutathionylation in Arabidopsis thaliana: detection using biotinylated glutathione. Plant Cell Physiol 44: 655–660 Jacquot JP, Rouhier N and Gelhaye E (2002) Redox control by dithiol–disulfide exchange in plants: I. The chloroplastic systems. Ann N Y Acad Sci 973: 508–519 Jung CH and Thomas JA (1996) S-glutathiolated hepatocyte proteins and insulin disulfides as substrates for reduction by glutaredoxin, thioredoxin, protein disulfide isomerase, and glutathione. Arch Biochem Biophys 335: 61–72 Kim J and Mayfield SP (1997) Protein disulfide isomerase as a regulator of chloroplast translational activation. Science 278: 1954–1957

219 Klappa P, Ruddock LW, Darby NJ and Freedman RB (1998) The b¢ domain provides the principal peptide-binding site of protein disulfide isomerase but all domains contribute to binding of misfolded proteins. EMBO J 17: 927–935 Kobrehel K, Wong JH, Balogh A, Kiss F, Yee BC and Buchanan BB (1992) Specific reduction of wheat storage proteins by thioredoxin h. Plant Physiol 99: 919–924 Kramer B, Ferrari DM, Klappa P, Pohlmann N and So¨ling HD (2001) Functional roles and efficiencies of the thioredoxin boxes of calcium-binding proteins 1 and 2 in protein folding. Biochem J 357: 83–95 Laboissiere MC, Sturley SL and Raines RT (1995) The essential function of protein-disulfide isomerase is to unscramble nonnative disulfide bonds. J Biol Chem 270: 28006–28009 Laloi C, Rayapuram N, Chartier Y, Grienenberger JM, Bonnard G and Meyer Y (2001) Identification and characterization of a mitochondrial thioredoxin system in plants. Proc Natl Acad Sci USA 98: 14144–14149 Laurent TC, Moore EC, and Reichard P (1964) Enzymatic synthesis of deoxyribonucleotides. IV. Isolation and characterization of thioredoxin, the hydrogen donor from Escherichia coli B. J Biol Chem 239: 3436–3444 Lemaire SD (2004) The glutaredoxin family in oxygenic photosynthetic organisms. Photosynthesis Res 79: 305– 318 Lemaire S, Keryer E, Stein M, Schepens I, Issakidis-Bourguet E, Ge´rard-Hirne C, Miginiac-Maslow M and Jacquot JP (1999a) Heavy-metal regulation of thioredoxin gene expression in Chlamydomonas reinhardtii. Plant Physiol 120: 773–778 Lemaire SD, Stein M, Issakidis-Bourguet E, Keryer E, Benoit V, Pineau B, Gerard-Hirne C, Miginiac-Maslow M and Jacquot JP (1999b) The complex regulation of ferredoxin/ thioredoxin-related genes by light and the circadian clock. Planta 209: 221–229 Lemaire SD, Miginiac-Maslow M and Jacquot JP (2002) Plant thioredoxin gene expression: control by light, circadian clock, and heavy metals. Meth Enzymol 347: 412–421 Lemaire SD, Collin V, Keryer E, Quesada A and MiginiacMaslow M (2003a) Characterization of thioredoxin y, a new type of thioredoxin identified in the genome of Chlamydomonas reinhardtii. FEBS Lett 543: 87–92 Lemaire SD, Collin V, Keryer E, Issakidis-Bourguet E, Lavergne D and Miginiac-Maslow M (2003b) Chlamydomonas reinhardtii: a model organism for the study of the thioredoxin family. Plant Physiol Biochem 41: 513–521 Lillig CH, Potamitou A, Schwenn JD, Vlamis-Gardikas A and Holmgren A (2003) Redox regulation of 3¢-phosphoadenylylsulfate reductase from Escherichia coli by glutathione and glutaredoxins. J Biol Chem 278: 22325–22330 Martin JL (1995) Thioredoxin – a fold for all reasons. Structure 3: 245–250 Meiri E, Levitan A, Guo F, Christopher DA, Schaefer D, Zryd JP and Danon A (2002) Characterization of three PDI-like genes in Physcomitrella patens and construction of knock-out mutants. Mol Genet Genomics 267: 231–240 Mestres-Ortega D and Meyer Y (1999) The Arabidopsis thaliana genome encodes at least four thioredoxins m and a new prokaryotic-like thioredoxin. Gene 240: 307–316 Meunier L, Usherwood YK, Chung KT and Hendershot LM (2002) A subset of chaperones and folding enzymes form

multiprotein complexes in endoplasmic reticulum to bind nascent proteins. Mol Biol Cell 13: 4456–4469 Minakuchi K, Yabushita T, Masumura T, Ichihara K and Tanaka K (1994) Cloning and sequence analysis of a cDNA encoding rice glutaredoxin. FEBS Lett 337: 157–160 Mkrtchian S, Fang C, Hellman U and Ingelman-Sundberg M (1998) A stress-inducible rat liver endoplasmic reticulum protein, ERp29. Eur J Biochem 251: 304–313 Monnat J, Hacker U, Geissler H, Rauchenberger R, Neuhaus EM, Maniak M and Soldati T (1997) Dictyostelium discoideum protein disulfide isomerase, an endoplasmic reticulum resident enzyme lacking a KDEL-type retrieval signal. FEBS Lett 418: 357–362 Monnat J, Neuhaus EM, Pop MS, Ferrari DM, Kramer B and Soldati T (2000) Identification of a novel saturable endoplasmic reticulum localization mechanism mediated by the Cterminus of a Dictyostelium protein disulfide isomerase. Mol Biol Cell 11: 3469–3484 Motohashi K, Kondoh A, Stumpp MT and Hisabori T (2001) Comprehensive survey of proteins targeted by chloroplast thioredoxin. Proc Natl Acad Sci USA 98: 11224–11229 Mouaheb N, Thomas D, Verdoucq L, Monfort P and Meyer Y (1998). In vivo functional discrimination between plant thioredoxins by heterologous expression in the yeast Saccharomyces cerevisiae. Proc Natl Acad Sci USA 95: 3312–3317 Nguyen Van P, Rupp K, Lampen A and So¨ling HD (1993) CaBP2 is a rat homologue of ERp72 with protein disulfide isomerase activity. Eur J Biochem 213: 789–795 Nulton-Persson AC, Starke DW, Mieyal JJ and Szweda LI (2003) Reversible inactivation of alpha-ketoglutarate dehydrogenase in response to alterations in the mitochondrial glutathione status. Biochemistry 42: 4235–4242 Park JB and Levine M (1996) Purification, cloning and expression of dehydroascorbic acid-reducing activity from human neutrophils: identification as glutaredoxin. Biochem J 315: 931–938 Patel-King RS, Benashki SE, Harrison A and King SM (1996) Two functional thioredoxins containing redox-sensitive vicinal dithiols from the Chlamydomonas outer dynein arm. J Biol Chem 271: 6283–6291 Prieto-Alamo MJ, Jurado J, Gallardo-Madueno R, MonjeCasas F, Holmgren A and Pueyo C (2000) Transcriptional regulation of glutaredoxin and thioredoxin pathways and related enzymes in response to oxidative stress. J Biol Chem 275: 13398–13405 Rodriguez-Manzaneque MT, Ros J, Cabiscol E, Sorribas A and Herrero E (1999) Grx5 glutaredoxin plays a central role in protection against protein oxidative damage in Saccharomyces cerevisiae. Mol Cell Biol 19: 8180–8190 Rodriguez-Manzaneque MT, Tamarit J, Belli G, Ros J and Herrero E (2002) Grx5 is a mitochondrial glutaredoxin required for the activity of iron/sulphur enzymes. Mol Biol Cell 13: 1109–1121 Rouhier N, Gelhaye E, Sautiere PE, Brun A, Laurent P, Tagu D, Gerard J, de Fay E, Meyer Y and Jacquot JP (2001) Isolation and characterization of a new peroxiredoxin from poplar sieve tubes that uses either glutaredoxin or thioredoxin as a proton donor. Plant Physiol 127: 1299–1309 Rouhier N, Gelhaye E and Jacquot JP (2002a) Glutaredoxindependent peroxiredoxin from poplar: protein-protein

220 interaction and catalytic mechanism. J Biol Chem 277: 13609–13614 Rouhier N, Gelhaye E and Jacquot JP (2002b) Redox control by dithiol–disulfide exchange in plants: II. The cytosolic and mitochondrial systems. Ann N Y Acad Sci 973: 520–528 Rupp K, Birnbach U, Lundstrom J, Van PN and So¨ling HD (1994) Effects of CaBP2, the rat analog of ERp72, and of CaBP1 on the refolding of denatured reduced proteins. Comparison with protein disulfide isomerase. J Biol Chem 269: 2501–2507 Scherens B, Dubois E and Messenguy F (1991) Determination of the sequence of the yeast YCL313 gene localized on chromosome III. Homology with the protein disulfide isomerase (PDI gene product) of other organisms. Yeast 7: 185– 193 Schu¨rmann P and Jacquot JP (2000) Plant thioredoxin systems revisited. Annu Rev Plant Physiol Plant Mol Biol 51: 371–400 Shorrosh BS and Dixon RA (1992) Sequence analysis and developmental expression of an alfalfa protein disulfide isomerase. Plant Mol Biol 19: 319–321 Shorrosh BS, Subramaniam J, Schubert KR and Dixon RA (1993) Expression and localization of plant protein disulfide isomerase. Plant Physiol 103: 719–726 Sineshchekov OA, Jung KH and Spudich JL (2002) Two rhodopsins mediate phototaxis to low- and high-intensity light in Chlamydomonas reinhardtii. Proc Natl Acad Sci USA 99: 8689–8694 Stein M, Jacquot JP, Jeannette E, Decottignies P, Hodges M, Lancelin JM Mittard V, Schmitter JM and Miginiac-Maslow M (1995) Chlamydomonas reinhardtii thioredoxins: structure of the genes coding for the chloroplastic m and cytosolic h isoforms; expression in Escherichia coli of the recombinant proteins, purification and biochemical properties. Plant Mol Biol 28: 487–503 Szederkenyi J, Komor E and Schobert C (1997) Cloning of the cDNA for glutaredoxin, an abundant sieve-tube exudate protein from Ricinus communis L. and characterisation of the glutathione-dependent thiol-reduction system in sieve tubes. Planta 202: 349–356

Tamarit J, Belli G, Cabiscol E, Herrero E and Ros J (2003) Biochemical characterization of yeast mitochondrial grx5 monothiol glutaredoxin. J Biol Chem 278: 25745–25751 Trebitsh T and Danon A (2001) Translation of chloroplast psbA mRNA is regulated by signals initiated by both Photosystems II and I. Proc Natl Acad Sci USA 98: 12289–12294 Trebitsh T, Levitan A, Sofer and Danon A (2000) Translation of chloroplast psbA mRNA is modulated in the light by counteracting oxidizing and reducing activities. Mol Cell Biol 20: 1116–1123 Trebitsh T, Meiri E, Ostersetzer O, Adam Z and Danon A (2001) The protein disulfide isomerase-like RB60 is partitioned between stroma and thylakoids in Chlamydomonas reinhardtii chloroplasts. J Biol Chem 276: 4564–4569 Verdoucq L, Vignols F, Jacquot JP, Chartier Y and Meyer Y (1999) In vivo characterization of a thioredoxin h target protein defines a new peroxiredoxin family. J Biol Chem 274: 19714–19722 Vlamis-Gardikas A and Holmgren A (2002) Thioredoxin and glutaredoxin isoforms. Meth Enzymol 347: 286–296 Wang J, Tekle E, Oubrahim H, Mieyal JJ, Stadtman ER and Chock PB (2003) Stable and controllable RNA interference: investigating the physiological function of glutathionylated actin. Proc Natl Acad Sci USA 100: 5103–5106 Washburn MP and Wells WW (1999) The catalytic mechanism of the glutathione-dependent dehydroascorbate reductase activity of thioltransferase (glutaredoxin). Biochemistry 38: 268–274 Wong JH, Balmer Y, Cai N, Tanaka CK, Vensel WH, Hurkman WJ and Buchanan BB (2003) Unraveling thioredoxin-linked metabolic processes of cereal starchy endosperm using proteomics. FEBS Lett 547: 151–156 Xia B, Vlamis-Gardikas A, Holmgren A, Wright PE and Dyson HJ (2001) Solution structure of Escherichia coli glutaredoxin-2 shows similarity to mammalian glutathione-S-transferases. J Mol Biol 310: 907–918 Yohn CB, Cohen A, Danon A and Mayfield SP (1998) A poly(A) binding protein functions in the chloroplast as a message-specific translation factor. Proc Natl Acad Sci USA 95: 2238–2243