Molecular Plant Advance Access published online on October 3, 2008
Molecular Plant, doi:10.1093/mp/ssn061
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Prompt and Easy Activation by Specific Thioredoxins of Calvin Cycle Enzymes of Arabidopsis thaliana Associated in the GAPDH/CP12/PRK Supramolecular Complex
a Laboratory of Molecular Plant Physiology, Department of Experimental Evolutionary Biology, University of Bologna, Via Irnerio 42, I-40126 Bologna, Italy
b Institut de Biotechnologie des Plantes, UMR 8618, CNRS/Univ. Paris-Sud 11, 91405 Orsay Cedex, France
1 To whom correspondence should be addressed. E-mail paolo.trost{at}unibo.it, fax +39051242576, tel. +390512091329.
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Abstract |
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 TOP Abstract INTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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The Calvin cycle enzymes glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and phosphoribulokinase (PRK) can form under oxidizing conditions a supramolecular complex with the regulatory protein CP12. Both GAPDH and PRK activities are inhibited within the complex, but they can be fully restored by reduced thioredoxins (TRXs). We have investigated the interactions of eight different chloroplast thioredoxin isoforms (TRX f1, m1, m2, m3, m4, y1, y2, x) with GAPDH (A4, B4, and B8 isoforms), PRK and CP12 (isoform 2), all from Arabidopsis thaliana. In the complex, both A4-GAPDH and PRK were promptly activated by TRX f1, or more slowly by TRXs m1 and m2, but all other TRXs were ineffective. Free PRK was regulated by TRX f1, m1, or m2, while B4- and B8-GAPDH were absolutely specific for TRX f1. Interestingly, reductive activation of PRK caged in the complex was much faster than reductive activation of free oxidized PRK, and activation of A4-GAPDH in the complex was much faster (and less demanding in terms of reducing potential) than activation of free oxidized B4- or B8-GAPDH. It is proposed that CP12-assembled supramolecular complex may represent a reservoir of inhibited enzymes ready to be released in fully active conformation following reduction and dissociation of the complex by TRXs upon the shift from dark to low light. On the contrary, autonomous redox-modulation of GAPDH (B-containing isoforms) would be more suited to conditions of very active photosynthesis.
Key Words: carbon metabolism • enzymology • light regulation • metabolic regulation • photosynthesis • Arabidopsis
Received for publication June 2, 2008. Accepted for publication August 19, 2008.
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INTRODUCTION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Thioredoxins (TRXs) are small ubiquitous redox proteins encoded by large gene families in oxygenic photosynthetic organisms (Lemaire et al., 2003; Buchanan and Balmer, 2005; Florencio et al., 2006; Meyer et al., 2006). Arabidopsis thaliana has been reported to contain at least 20 genes coding for classical TRX proteins of 10–14 kDa with a conserved WC(G/P)PC motif in the active site (Meyer et al., 2005; Lemaire et al., 2007), nine of which are targeted to plastids (Collin et al., 2003, 2004; Lemaire et al., 2007; Schürmann and Buchanan, 2008). Based on sequence similarities, plastid TRXs have been classified into four types (f, m, x, y), each including one or more isoforms. Apart from TRX y1, which is mainly expressed in non-photosynthetic organs (Collin et al., 2004), all other TRXs appear to be predominantly, but not exclusively, expressed in leaves (Schmid et al., 2005; de Dios Barajas-Lopez et al., 2007; Traverso et al., 2008).
In chloroplasts, TRXs are reduced in the light by Photosystem I via ferredoxin and ferredoxin:thioredoxin reductase (Dai et al., 2007; Schürmann and Buchanan, 2008). The active site dithiol of reduced TRXs can reduce disulfide bridges on target proteins, in most cases enzymes that are reversibly activated by reduction (Buchanan and Balmer, 2005). The multiplicity of TRX genes is paralleled by a multiplicity of TRX targets, most of which have been recently identified through redox proteomic approaches (Michelet et al., 2006; Lemaire et al., 2007). In chloroplasts, these include, for instance, all 11 enzymes of the Calvin cycle, besides Rubisco activase (Portis et al., 2008) and CP12 (Wedel et al., 1997), which are involved in Calvin cycle regulation. Only in few cases, however, has the specificity of TRX targets for different TRX isoforms been thoroughly investigated (Schürmann and Buchanan, 2008). In general, TRXs f and m were found to be principally involved in enzyme regulation, including light-modulation of photosynthetic metabolism, while TRXs x and y seem to serve mainly as hydrogen donors for antioxidant enzymes (e.g. peroxiredoxins: Collin et al., 2003, 2004; glutathione peroxidases: Navrot et al., 2006; methionine sulfoxides reductases: Vieira dos Santos et al., 2007).
In this work, we investigated the specificity of interactions between chloroplast TRXs and two enzymes of the Calvin cycle (glyceraldehyde-3-phosphate dehydrogenase, GAPDH, and phosphoribulokinase, PRK) which are known to form a supramolecular complex with CP12 under oxidizing conditions, those prevailing when the photosynthetic electron transport is decreased under limiting light or stress conditions or in the dark (Wedel et al., 1997; Scheibe et al., 2002; Graciet et al., 2004; Trost et al., 2006; Howard et al., 2008). All experiments were performed with recombinant proteins from Arabidopsis thaliana, namely eight chloroplast TRX isoforms (f1, m1, m2, m3, m4, x, y1, and y2; Collin et al., 2003, 2004) and five different targets (GAPDH-isoforms A4, B4, and B8; CP12-isoform 2 and PRK; Marri et al., 2005).
PRK is a dimer with one TRX-sensitive disulfide per subunit (Porter et al., 1988) and, although disulfide formation itself causes a significant inhibition of PRK activity (Hirasawa et al., 1999; Geck and Hartman, 2000; Hutchinson et al., 2000), inclusion of oxidized PRK into the supramolecular complex with GAPDH and CP12 leads to additional inhibition of enzyme activity (Marri et al., 2005). CP12 may contain two intramolecular disulfides and, similarly to PRK, has to be fully oxidized to promote the assembly of the supramolecular complex including also GAPDH (Wedel and Soll, 1998; Graciet et al., 2003; Marri et al., 2008).
Calvin cycle's GAPDH exists in different isoforms and may either have no disulfides (isoform A4) or contain two identical disulfides per tetramer, one in each of the C-terminal extensions specific for B-subunits (isoforms A2B2 or A8B8, Baalmann et al., 1996; Fermani et al., 2007). Recombinant GAPDH isoforms made of B-subunits only (B4 or B8) were never observed in vivo but found to display similar properties to native AB-GAPDH isoforms (A2B2 or A8B8), including the autonomous regulation by TRXs and metabolites such as NAD(P)(H) and 1,3-bisphosphoglycerate (Baalmann et al., 1996; Li and Anderson, 1997; Sparla et al., 2002, 2005). The A4-GAPDH isoform is instead completely insensitive to regulation by TRXs and metabolites (Baalmann et al., 1996; Scagliarini et al., 1998; Sparla et al., 2002), although catalytic cysteines (Cys-149) of A4-GAPDH can form mixed disulfides with glutathione under oxidative stress conditions (Zaffagnini et al., 2007). This post-translational modification leads to complete loss of enzyme activity, but can be reverted by glutaredoxins (Zaffagnini et al., 2008). On the other hand, complex formation with PRK and CP12 leads to reversible inhibition of A4-GAPDH activity under physiological conditions (Graciet et al., 2004; Marri et al., 2005) and has been envisioned as a mechanism for light-modulation of A4-GAPDH in land plants (Trost et al., 2006; Marri et al., 2008), similar to the regulatory system of lower photosynthetic organisms that do not contain AB-GAPDH (Wedel and Soll, 1998; Graciet et al., 2004; Tamoi et al., 2005; Oesterhelt et al., 2007). Due to the close similarity between A- and B-subunits, a similar regulation based on CP12 might also affect A2B2-GAPDH, which was observed to be partially complexed with CP12 and PRK in the dark or under low-light conditions in different higher plant species (Wedel and Soll, 1998; Scheibe et al., 2002; Howard et al., 2008).
Here, we show that the inhibition of the activities of enzymes complexed with CP12 (A4-GAPDH and PRK) can be rapidly and fully reversed by specific TRX isoforms. Interestingly, this process is much faster than the reductive activation of free enzymes (B-GAPDH or PRK) and, in the case of GAPDH, is also less demanding in terms of reducing potential. Since this regulatory mechanism based on the dissociation of a supramolecular complex results in the parallel activation of two Calvin cycle enzymes, we speculate that its physiological function might be to ensure a fast and coordinated onset of the Calvin cycle upon the transition from dark to light (Howard et al., 2008).
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RESULTS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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A supramolecular complex comprising A4-GAPDH, CP12 (isoform 2; Trost et al., 2006), and PRK of Arabidopsis thaliana was reconstituted in vitro by incubating the three purified recombinant proteins with NAD under oxidizing conditions (oxidized DTT, Marri et al., 2005). The 500-kDa complex included two GAPDH tetramers, two PRK dimers, and four CP12 monomers (Marri et al., 2008) and eluted in size-exclusion chromatography as a single symmetrical peak (Figure 1) with strongly inhibited enzyme activities. The NADPH-dependent GAPDH activity of the complex was 26 ± 6% (SD, n = 16) of the NADPH-activity of free A4-GAPDH. PRK activity of the complex was 5 ± 2% (SD, n = 10) of the activity of the fully reduced free enzyme.
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Thermodynamics of Complex Dissociation and GAPDH/PRK Reactivation
The reactivation process of GAPDH and PRK embedded in the A4-GAPDH/CP12/PRK complex was analyzed by equilibrium redox titrations in the presence of a cocktail of different Arabidopsis chloroplast thioredoxins and different ratios of oxidized/reduced DTT (Figure 2). In these experiments, TRXs acted as redox mediators, favoring full equilibration of the complex with the redox potential established by DTT (Hutchinson and Ort, 1995; Hirasawa et al., 1999). The measurements were performed at pH 7.9, corresponding to the pH of the chloroplast stroma in the light.
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Although A4-GAPDH contains no redox-sensitive disulfides, the NADPH-dependent GAPDH activity of the complex responded to the redox potential, while the NADH-dependent activity was constant. By moving from less negative redox potentials (mimicking low light intensities in vivo) to more negative ones (mimicking high light intensities), the NADPH-activity of GAPDH reached its maximal level (Figure 2A). Experimental data were fitted by nonlinear regression to a Nernst equation, including one thiol/disulfide equilibrium, resulting in a midpoint redox potential of –335 ± 10 mV. Equations including two or more thiol/disulfide equilibria did not result in better interpolations of experimental data.
For comparison, a similar redox titration analysis was performed on recombinant B4-GAPDH of Arabidopsis, known to be redox-regulated due to the presence of the C-terminal extension of B-subunits, which is closely homologous to the C-terminal sequence of CP12 (Pohlmeyer et al., 1996; Baalmann et al., 1996; Sparla et al., 2002). However, the redox response of B4-GAPDH was characterized by a more negative midpoint redox potential of –359 mV (Figure 2A), similar to both B4-GAPDH and A2B2-GAPDH from spinach (–357 and –353 mV, respectively; Sparla et al., 2005).
Redox titration analysis of PRK embodied in the A4-GAPDH/CP12/PRK complex resulted in a midpoint redox potential of –326 ± 3 mV (Figure 2B). The redox-response of PRK in the complex resembled that of free PRK (Em,7.9 –330 mV, Marri et al., 2005), although oxidized PRK within the complex was less active (5 ± 2% residual activity) than free PRK when oxidized under identical conditions (21 ± 3% residual activity after 3 h incubation with 20 mM oxidized DTT). This result clearly indicated that complex formation provided additional inhibition of PRK activity, over and above enzyme oxidation.
Kinetics and Specificity versus Thioredoxin Isoforms of Complex Dissociation and GAPDH/PRK Reactivation
The activation rates of both GAPDH and PRK activities embodied in the complex were studied in the presence of eight different chloroplast TRX isoforms from Arabidopsis (f1, m1, m2, m3, m4, y1, y2, x; Collin et al., 2003, 2004). Among classical TRXs known to be present in Arabidopsis chloroplasts, only TRX f2 was not tested. It should be noted, however, that TRX f1 and f2 are coded by recently duplicated paralogous genes (Meyer et al., 2005) and are almost identical in sequence, sharing more than 90% identity at the protein level. In time-course experiments of reductive activation, TRX isoforms (2 µM) were kept reduced by a fixed concentration of DTT (0.2 mM) that had limited effects by itself on either enzyme activities (Figure 3) or complex stability (not shown).
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The inhibited GAPDH activity (NADPH-dependent) of the complex was restored very rapidly to full activity by incubation with TRX f1 at saturating concentration (t1/2 0.26 ± 0.01 min). At the same concentration of 2 µM, TRX m1 and m2 required much longer incubations to fully activate GAPDH (t1/2 1.4 and 6 min, respectively). The effect of all other TRXs was negligible, namely within the experimental error if compared with DTT alone.
For comparison, activation of oxidized GAPDH formed by B-subunits only was analyzed and found to exhibit a more stringent specificity (Figure 3). In fact, B4-GAPDH, a partially activated form stabilized by NADP (Sparla et al., 2002), was absolutely specific for TRX f1 and reached full activation with slower kinetics (t1/2 2.2 ± 0.5 min) than complex-embedded A4-GAPDH (Figure 3B and Table 1). An NAD-stabilized aggregated form, possibly B8-GAPDH (Sparla et al., 2002), behaved identically in terms of absolute specificity for TRX f1, but displayed even slower kinetics of activation (t1/2 14.1 ± 0.3 min, Figure 3C and Table 1).
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Analysis of PRK activation rates after incubation of the complex with different TRX isoforms yielded qualitatively similar results to A4-GAPDH reactivation. Full PRK activity was recovered during incubations with TRX f1, m1, or m2 with decreasing efficiency in this order (Figure 3D), while other plastidial TRXs had little or no effect (less than 10% additional activation with respect to DTT alone after 30 min incubation). Within the experimental error, PRK activation rate by TRX f1 (t1/2 0.6 ± 0.4 min) or m1 (t1/2 2.2 ± 0.5 min) was similar to A4-GAPDH activation under identical conditions (Table 1). On the other hand, different activation rates were observed with TRX m2, which proved to be a much weaker activator of PRK (t1/2 25 ± 7 min) than of A4-GAPDH (6 ± 2 min; Table 1). This latter observation suggests that reductive activation of either A4-GAPDH or PRK embedded in the complex does not depend on one and the same mechanism.
Specificity of redox regulation of PRK in the free state was also tested. Again, reductive activation of free PRK was best performed by TRX f1 (t1/2 3.9 ± 0.9 min), followed by TRX m2 (14 ± 4 min) and TRX m1 (17 ± 6 min) (Figure 3E). All other TRX isoforms were as efficient as DTT alone within the experimental error. Interestingly, activation of free PRK by either TRX f1 or m1 was much slower than activation of complexed PRK (Table 1), suggesting that CP12-complex formation could be a means for both strongly inhibiting PRK activity and rapidly delivering the fully active enzyme upon reduction. A different oxidizing treatment with DTNB, as suggested by Geck and Hartman (2000), was also applied to free PRK. Under these conditions, the activity of PRK (2 ± 1%, n = 8) was similar to that of PRK in the complex. However, reductive activation of DTNB-treated PRK led to slow and partial recovery of enzyme activity even after 1 h incubation with reduced TRXs (not shown), preventing a detailed characterization of the reactivation process.
CP12 alone was also tested for its sensitivity to reduction by different TRX isoforms. Fully reduced and oxidized CP12, obtained by 3 h incubation with 20 mM DTT (reduced or oxidized), were resolved by analytical size-exclusion chromatography (Figure 4A). Although peaks largely overlapped, elution volumes corresponding to maximal absorption of each peak were reproducible: reduced CP12 eluted at 1.350 ± 0.009 ml and oxidized CP12 at 1.379 ± 0.003 ml (n = 3). Samples of oxidized CP12, after accurate desalting to remove excess DTT, were individually incubated with reduced DTT plus different plastidic TRX isoforms and analyzed as above. Due to limitations of the analytical method, time-course experiments of CP12 reduction could not be performed, and specificity was assessed by measuring the redox state of CP12 after incubations with varying concentrations of TRXs (1–15 µM). DTT alone (0.5 mM) was unable to reduce CP12. As shown in Figure 4, both TRX f1 and m1 were already efficient at 2–5 µM, whereas TRX m2 was efficient at higher concentrations.
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DISCUSSION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Several effectors including TRXs, pyridine nucleotides, and metabolic intermediates, contribute in higher plants to tune the Calvin cycle with the activity of photosynthetic electron transport, which, under non-stressed conditions, primarily depends on the variable amount of light intercepted by leaves. To this aim, several enzymes of the Calvin cycle are finely regulated, including PRK and GAPDH, which catalyze two of the three energy-consuming reactions of the cycle. Both enzymes are modulated in a coordinated manner by thiol-based redox regulation (Geck and Hartman, 2000; Fermani et al., 2007), but also by protein–protein interactions through the intrinsically unstructured protein CP12 (Wedel et al., 1997; Graciet et al., 2004). Supramolecular complex formation with CP12 leads to additional inhibition of both enzyme activities (Marri et al., 2005). However, since chloroplast CP12 also contains two intramolecular disulfides involved in its assembling activity (Wedel and Soll, 1998; Lebreton et al., 2006; Marri et al., 2008), CP12-dependent effects would also depend, in turn, on the redox state of TRXs. In chloroplasts, the redox state of TRXs depends on the electron pressure from photosystem I (Schürmann and Buchanan, 2008) such that the regulatory properties of both PRK, GAPDH, and CP12 seem perfectly suited to modulate the Calvin cycle under conditions of variable photosynthetic activity.
Here, we show that both GAPDH and PRK can be recovered as fully active enzymes upon incubation of the supramolecular complex GAPDH/CP12/PRK with specific TRX isoforms. Experiments were performed exclusively with recombinant proteins of Arabidopsis thaliana, enabling specificity to be assessed. Although it has been shown that GAPDH/CP12/PRK complexes may contain TRX-sensitive AB-GAPDH besides TRX-insensitive A4-GAPDH (Wedel and Soll, 1998; Scheibe et al., 2002; Howard et al., 2008), we have reconstituted the complex in vitro only with A4-GAPDH. Apart from reversible glutathionylation, a type of regulation that might be relevant under stress (Zaffagnini et al., 2007, 2008), A4-GAPDH is not itself redox-regulated, but it may represent a tool to investigate CP12-dependent effects in that the reductive activation of A4-GAPDH caged in the complex can be taken as a measure of complex dissociation.
Kinetic and structural properties of A4-GAPDH are very similar to those of reduced (activated) A2B2-GAPDH (Sparla et al., 2004), however, and both isoforms may be expected to behave similarly in the complex. The crystal structure of oxidized A2B2-GAPDH shows that the CTE bearing an internal disulfide tightly packs into a cleft between a pair of A/B-subunits, thereby causing inhibition of enzyme activity (Fermani et al., 2007). The C-terminal part of CP12, homologous to the CTE of GAPDH (Pohlmeyer et al., 1996), is involved in the interaction between CP12 and GAPDH (Wedel and Soll, 1998; Lebreton et al., 2006; Marri et al., 2008) and would possibly compete with oxidized CTE for the same interaction sites on GAPDH (Trost et al., 2006). We thus speculate that A2B2-GAPDH, in order to be engaged in the complex with CP12 and PRK, should be reduced, namely in a similar conformation to A4-GAPDH.
Although several effectors other than TRXs can also dissociate the GAPDH/CP12/PRK complex and activate both enzyme activities to some extent (e.g. 1,3-bisphosphoglycerate, NADP(H), ATP, Wedel and Soll, 1998; Graciet et al., 2004; Marri et al., 2005; Tamoi et al., 2005; Lebreton et al., 2006), specific TRXs proved to be highly efficient in recovering full activity of both GAPDH and PRK. Among the eight chloroplast TRXs tested, only TRX f1, m1, and m2 were found effective, TRX f1 being by far the most efficient. At equilibrium, activation of both enzymes occurred at comparable redox potentials (Em,7.9 –335 mV for GAPDH and –326 mV for PRK) corresponding to a minimal amount of reduced TRXs with respect to the total pool (e.g. –330 mV corresponds to 16% reduction of the TRX f pool, having an Em,7.9 of –351 mV; Collin et al., 2003). This suggests that initial reactivation of GAPDH and PRK embodied in the complex may be triggered in vivo at relatively low light intensities, allowing an easy start-up of the Calvin cycle well before full reduction of TRXs could be attained.
In agreement with our evidence in vitro, Howard et al. (2008) observed in darkened pea leaves that GAPDH/CP12/PRK complex dissociation and enzyme activation started at low light intensities (equivalent to not very negative redox potentials) and was completed well before saturation of photosynthesis by high light intensities (more negative redox potentials). Interestingly, while the redox response of free PRK (Em,7.9 –330 mV, Marri et al., 2005; Hirasawa et al., 1999; Hutchinson et al., 2000) equals the response of PRK in the complex (–326 mV), activation of free GAPDH (B-containing isoforms) requires more reducing conditions (–359 mV) than activation of A4-GAPDH (–335 mV). CP12 therefore seems to provide a mechanism for tight coupling of GAPDH and PRK responses to the redox conditions of the chloroplasts, particularly by making GAPDH sensitive to partial reduction of thioredoxin pools (f, m1, m2), such as under conditions of low light intensity. The autonomous regulation of AB-GAPDH (CTE-mediated) would be tuned instead with conditions of very active photosynthesis and specifically dependent on TRX f. Under these conditions, the strong GAPDH activator 1,3-bisphosphoglycerate (Trost et al., 1993) would also increase in chloroplasts (Baalmann et al., 1994), thereby contributing to enzyme activation in a synergetic mechanism with TRX f1 (Baalmann et al., 1995). In addition, the specific effect of TRX f1 could be further modulated by glutathionylation of TRX f, which limits its capability to reactivate oxidized B4-GAPDH. Interestingly, among all chloroplast TRX isoforms of Arabidopsis, this post-translational modification was found specific for TRX f (Michelet et al., 2005).
Besides requiring a limited reducing power, reductive activation of either GAPDH or PRK in the complex was relatively fast, particularly with TRX f1 (t1/2 0.26 min for GAPDH and 0.6 min for PRK; Table 1). As noted above, A4-GAPDH activation corresponded to complex dissociation while full activation of PRK required both enzyme reduction (oxidation being responsible for about 80% enzyme inhibition) and complex dissociation (additional 15% inhibition, Figure 1B). Within the complex, PRK activation by TRX f1 or m1 was not significantly slower than A4-GAPDH activation, indicating that activation by reduction of complexed PRK is fast enough to cope with dissociation of the complex. Surprisingly, however, reductive activation of free oxidized PRK required longer times than activation of PRK in the complex and qualitatively similar results were obtained by Gontero et al. (1993) with proteins purified from spinach, well before the discovery of CP12. Clearly, PRK in the complex is locked into a conformation that, although almost inactive, is more receptive to reductive activation than the free oxidized enzyme.
In a similar way, activation of A4-GAPDH in the complex was found to be much faster than reductive activation of free GAPDH isoforms (either B4 or B8). Recombinant B-isoforms of GAPDH behave very similarly to native AB-isoforms, in terms of both regulatory properties (Baalmann et al., 1996; Li and Anderson, 1997; Sparla et al., 2002) and kinetic parameters (Sparla et al., 2005). In particular, B4-GAPDH resembles light-activated A2B2-GAPDH and B8-GAPDH resembles dark-inhibited A8B8-GAPDH (Scagliarini et al., 1993; Baalmann et al., 1994). Although we cannot exclude subtle differences in TRX activation between recombinant and native GAPDH isoforms, our results suggest again that release of active enzymes from the complex can be faster than reductive activation of free enzymes (Figure 3). In full agreement with these findings, the very recent observation that complex dissociation in pea leaves could be triggered by light in a time frame of minutes (Howard et al., 2008) adds strong physiological relevance to this picture. In this work (Howard et al., 2008), it is also shown that PRK can exist in the reduced state within the complex, thus suggesting that complex dissociation would not depend on PRK reduction. We thus propose that complex dissociation depends mainly on the reduction of CP12, a process performed most efficiently by TRX f1, or by TRX m1 or TRX m2 with decreasing efficiency. All these TRX isoforms were indeed demonstrated to be able to reduce isolated CP12. In contrast, the redox-regulated B4-GAPDH is very specifically activated by thioredoxin f, m-type thioredoxins being totally inefficient. In this respect, it can be noted that neither thioredoxin f nor B-type GAPDH does exist in prokaryotes.
A sophisticated modulation of Calvin cycle enzymes is a necessity for land plants that cannot escape high light stress. Our findings support the conclusion that protein–protein interactions mediated by CP12 confer specific regulatory properties on two redox-regulated enzymes of the Calvin cycle, namely GAPDH and PRK. Thanks to CP12, redox-insensitive A4-GAPDH becomes redox-regulated, and behaves very similarly to PRK embedded in the same complex, both in terms of sensitivity to the redox potential and in terms of rapidity of reactivation under reducing conditions (particularly with TRX f1 and m1). Besides coordinate regulation, the complexed enzymes respond rapidly to a limited reduction of the TRX pool, suggesting that CP12-complexes may be mainly involved in short-term response of the Calvin cycle under relatively low light (for instance, the transition between night and dawn). On the other hand, it is tempting to speculate that free oxidized enzymes, particularly AB-GAPDH, which requires longer times for activation and stronger reducing potentials, might represent an inhibited form best suited for modulation at high light intensities. Indeed, Howard et al. (2008) showed that GAPDH total activity in pea chloroplasts further increased after full dissociation of the complex.
From an evolutionary point of view, it must be noted that the thioredoxin-regulated AB-GAPDH exists only in land plants, whereas A4-GAPDH and CP12 are present also in cyanobacteria, which are mobile prokaryotic organisms. A greater sophistication of enzyme redox regulation in higher plants has already been observed for NADP-malate dehydrogenase (NADP-MDH), which exhibits slow activation kinetics and two regulatory disulfides in higher plants (Em,7.9 –385 and –335 mV, respectively, for the sorghum enzyme, recalculated from Hirasawa et al., 2000), and fast activation kinetics and one regulatory disulfide (Em,7.9 –370 mV, Lemaire et al., 2005, recalculated) in the unicellular flagellate alga Chlamydomonas reinhardtii. In higher plants, the redox potential of the N-terminal disulfide of NADP-MDH (Em,7.9 –335 mV) suggests that, at low light intensities, the enzyme could be present in a ‘pre-reduced’ form, ready to become functional as soon as the ambient redox potential is decreased and allows reduction of the more negative C-terminal disulfide (Hirasawa et al., 2000). Thus, the increasing complexity of enzyme redox regulation seems to be a general evolutionary characteristic of higher plants.
It is notable that Trx f, existing only in eukaryotic photosynthetic organisms, activates very specifically not only fructose-1,6-bisphosphatase, but also A2B2-GAPDH and it is also the most efficient thioredoxin for activation of PRK. It seems, therefore, that thioredoxin f evolved as a general regulator of enzymes involved in photosynthetic carbon metabolism, ensuring the coordination of their activities.
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METHODS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODSFUNDING |
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Protein Expression and Purification
Heterologous expression and purification of Arabidopsis A4-GAPDH (GapA-1, At3g26650), CP12-2 (At3g62410), and PRK (At1g32060) were performed as in Marri et al. (2005). Arabidopsis chloroplast thioredoxin isoforms f1, m1, m2, m3, m4, x, y1, and y2 were heterologously expressed and purified as in Collin et al. (2003, 2004).
B-GAPDH was obtained by cloning the sequence encoding Arabidopsis GAPDH B-subunit (GapB, At1g42970, provided by TAIR) into a pET-29 expression vector (Novagen). The resulting plasmid was used to transform an E. coli BL21 (DE3) strain. The expression and purification of recombinant GAPDH were performed according to Sparla et al. (2002), except that the final anion-exchange chromatographic step was replaced by a hydrophobic interaction column (Phenyl Sepharose, Amersham Biosciences). Final preparations were electrophoretically pure, as judged by SDS–PAGE.
In order to obtain either B4- or B8-GAPDH, the protein was incubated at 4°C for 2 h in 25 mM potassium phosphate, pH 7.5, supplemented either with 0.2 mM NADP, or 0.2 mM NAD. The resulting B4- or B8-GAPDH was then purified on a Superdex 200 HR column (GE Healthcare) equilibrated in 25 mM potassium phosphate, pH 7.5, 1 mM EDTA, 150 mM KCl, and either 0.2 mM NADP for the tetrameric form, or 0.2 mM NAD for the aggregated form.
Reconstitution of the A4-GAPDH/CP12/PRK complex was obtained by incubating the purified recombinant proteins at 25°C for 3 h in 25 mM potassium phosphate, pH 7.5, 0.2 mM NAD, 2 mM oxidized DTT in a molar ratio of 1 (A4-GAPDH) : 2 (CP12-2) : 1 (PRK), according to Marri et al. (2008). The reconstituted complex was purified on a Superdex 200 HR column (GE Healthcare), equilibrated in 25 mM potassium phosphate, pH 7.5, 1 mM EDTA, 150 mM KCl, 0.2 mM NAD. Fractions containing the 500-kDa ternary complex (Figure 1) were collected, desalted with PD10 columns (GE Healthcare) and stored in 100 mM Tricine–NaOH, pH 7.9.
Redox Titrations
Redox titration experiments were performed with the reconstituted ternary complex in 100 mM Tricine–NaOH, pH 7.9. The A4-GAPDH/CP12/PRK complex (0.3 µM) was incubated for 3 h at 25°C with TRXs f1, m1, m2 (0.2 µM each) and 20 mM reduced/oxidized DTT in various dithiol/disulfide ratios in a final volume of 50 µl (Sparla et al., 2002). Following incubation, NAD(P)H-dependent GAPDH activities and PRK activity were assayed (Marri et al., 2005). Results were fitted by non-linear regression (CoStat, CoHort Software) to the Nernst equation (n = 2, one thiol/disulfide; Hirasawa et al., 2000; Sparla et al., 2002). Midpoint redox potentials are reported as means ± standard deviations of two independent experiments. Redox titration of B4-GAPDH (0.5 µM) was performed under the same conditions as complex A4-GAPDH/CP12/PRK, except that only TRX f1 (0.2 µM) was included in incubation media.
Reduction/Activation by Chloroplast Thioredoxin Isoforms of GAPDH, PRK, and CP12
To test the specificity of A4-GAPDH and PRK activation within the complex by chloroplast TRXs, 0.3 µM reconstituted complex was incubated in 100 mM Tricine–NaOH, pH 7.9, plus 0.2 mM reduced DTT with or without 2 µM TRXs. Aliquots were withdrawn over time (0–60 min) to assay GAPDH and PRK enzymatic activities. Activities are expressed as percentage of fully activated enzymes (Marri et al., 2005). Experimental data were fitted to an exponential equation and half-time of activation (t1/2) was graphically obtained. TRXs that, following 30 min incubation, did not provide more than 10% additional activation in respect to DTT alone were considered inactive (and respective data were not reported in Figure 3).
The same specificity tests were performed with 2 µM pure PRK in 100 mM Tricine–NaOH, pH 7.9, 0.2 mM reduced DTT with 5 µM TRXs. Oxidized PRK was obtained either by incubation with oxidized DTT (20 mM, 3 h) followed by desalting, or incubation with DTNB (5 µM, 5 min), following the procedure of Geck and Hartman (2000). Similar experiments were also performed with B4-GAPDH and B8-GAPDH (2.5 µM tetramer concentration).
Thioredoxin specificity versus CP12-2 was analyzed by monitoring CP12 behavior in size-exclusion chromatography. Fully reduced or fully oxidized CP12-2 were obtained by incubation of 100 µM recombinant CP12-2 in 60 µl of 25 mM potassium phosphate, pH 7.5, including 20 mM reduced or oxidized DTT. After 3 h at 25°C, albumin was added as an internal standard and samples were loaded on a Superdex 75 PC3.2/30 column (GE Healthcare), equilibrated in 25 mM potassium phosphate, pH 7.5, 150 mM KCl, 1 mM EDTA. Control experiments performed with 0.5 mM reduced DTT demonstrated that CP12 remained oxidized. To test the capability of the different thioredoxin isoforms to reduce CP12-2, samples were prepared under the same conditions except for the inclusion of 0.5 mM reduced DTT and TRXs at variable concentrations. After 10 min incubation, thiols were blocked with 10 mM 2-iodoacetamide for 30 min. Albumin as standard was added immediately before loading the sample on the column and chromatographed as above. The column was calibrated with standard globular proteins: Ribonuclease A, Chymotrypsinogen A, Ovalbumin and Serum Albumin, 5.0 mg ml–1, from the Gel Filtration Calibration Kit (GE Healthcare) (Sparla et al., 2002).
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Work supported by the Italian Ministry of University (FIRB 2003). No conflict of interest declared.
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2 These authors contributed equally to this work.
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Baalmann E, Backhausen JE, Klitzmann C, Scheibe R. Regulation of NADP-dependent glyceraldehyde-3-phosphate dehydrogenase activity in spinach chloroplasts. Bot. Acta. (1994) 107:313–320.[Medline]
Baalmann E, Backhausen JE, Rak C, Vetter S, Scheibe R. Reductive modification and nonreductive activation of purified spinach chloroplast NADP-dependent glyceraldehyde-3-phosphate dehydrogenase. Arch. Biochem. Biophys. (1995) 324:201–208.[CrossRef][Web of Science][Medline]
Baalmann E, Scheibe R, Cerff R, Martin W. Functional studies of chloroplast glyceraldehyde-3-phosphate dehydrogenase subunits A and B expressed in Escherichia coli: formation of highly active A4 and B4 homotetramers and evidence that the aggregation of the B4 complex is mediated by the B-subunit carboxy terminus. Plant Mol. Biol. (1996) 32:505–513.[CrossRef][Web of Science][Medline]
Buchanan BB, Balmer Y. Redox regulation: a broadening horizon. Annu. Rev. Plant Biol. (2005) 56:187–220.[CrossRef][Medline]
Collin V, Issakidis-Bourguet E, Marchand C, Hirasawa M, Lancelin JM, Knaff DB, Miginiac-Maslow M. The Arabidopsis plastidial thioredoxins: new functions and new insights into specificity. J. Biol. Chem. (2003) 278:23747–23752.
Collin V, Lamkemeyer P, Miginiac-Maslow M, Hirasawa M, Knaff DB, Dietz KJ, Issakidis-Bourguet E. Characterization of plastidial thioredoxins from Arabidopsis belonging to the new y-type. Plant Physiol (2004) 136:4088–4095.
Dai S, Friemann R, Glauser DA, Bourquin F, Manieri W, Schürmann P, Eklund H. Structural snapshots along the reaction pathway of ferredoxin-thioredoxin reductase. Nature (2007) 448:92–96.[CrossRef][Web of Science][Medline]
de Dios Barajas-López J, Serrato AJ, Olmedilla A, Chueca A, Sahrawy M. Localization in roots and flowers of pea chloroplastic thioredoxin f and thioredoxin m proteins reveals new roles in nonphotosynthetic organs. Plant Physiol (2007) 145:946–960.
Fermani S, Sparla F, Falini G, Martelli PL, Casadio R, Pupillo P, Ripamonti A, Trost P. The molecular mechanism of thioredoxin regulation in photosynthetic A2B2-glyceraldehyde-3-phosphate dehydrogenase. Proc. Natl Acad. Sci. U S A (2007) 104:11109–11104.
Florencio FJ, Pérez-Pérez ME, López-Maury L, Mata-Cabana A, Lindahl M. The diversity and complexity of the cyanobacterial thioredoxin systems. Photosynth. Res. (2006) 89:157–171.[CrossRef][Web of Science][Medline]
Geck MK, Hartman FC. Kinetic and mutational analyses of the regulation of phosphoribulokinase by thioredoxins. J. Biol. Chem. (2000) 275:18034–18039.
Gontero B, Mulliert G, Rault M, Giudici-Orticoni MT, Ricard J. Structural and functional properties of a multi-enzyme complex from spinach chloroplasts. 2. Modulation of the kinetic properties of enzymes in the aggregated state. Eur. J. Biochem. (1993) 217:1075–1082.[Web of Science][Medline]
Graciet E, Gans P, Wedel N, Lebreton S, Camadro JM, Gontero B. The small protein CP12: a protein linker for supramolecular complex assembly. Biochemistry (2003) 42:8163–8170.[CrossRef][Web of Science][Medline]
Graciet E, Lebreton S, Gontero B. Emergence of new regulatory mechanisms in the Benson-Calvin pathway via protein-protein interactions: a glyceraldehyde-3-phosphate dehydrogenase/CP12/phosphoribulokinase complex. J. Exp. Bot. (2004) 55:1245–1254.
Hirasawa M, Ruelland E, Schepens I, Issakidis-Bourguet E, Miginiac-Maslow M, Knaff DB. Oxidation-reduction properties of the regulatory disulfides of sorghum chloroplast nicotinamide adenine dinucleotide phosphate-malate dehydrogenase. Biochemistry (2000) 28:3344–3350.
Hirasawa M, Schürmann P, Jacquot JP, Manieri W, Jacquot P, Keryer E, Hartman FC, Knaff DB. Oxidation-reduction properties of chloroplast thioredoxins, ferredoxin:thioredoxin reductase, and thioredoxin f-regulated enzymes. Biochemistry (1999) 38:5200–5205.[CrossRef][Web of Science][Medline]
Howard TP, Metodiev M, Lloyd JC, Raines CA. Thioredoxin-mediated reversible dissociation of a stromal multiprotein complex in response to changes in light availability. Proc. Natl Acad. Sci. U S A (2008) 105:4056–4061.
Hutchinson RS, Ort DR. Measurement of equilibrium midpoint potentials of thiol/disulfide regulatory groups on thioredoxin-activated chloroplast enzymes. Methods Enzymol (1995) 252:220–228.[Web of Science][Medline]
Hutchinson RS, Groom Q, Ort DR. Differential effects of chilling-induced photooxidation on the redox regulation of photosynthetic enzymes. Biochemistry (2000) 6:6679–6688.
Lebreton S, Andreescu S, Graciet E, Gontero B. Mapping of the interaction site of CP12 with glyceraldehyde-3-phosphate dehydrogenase from Chlamydomonas reinhardtii: functional consequences for glyceraldehyde-3-phosphate dehydrogenase. FEBS J (2006) 273:3358–3369.[CrossRef][Medline]
Lemaire SD, Collin V, Keryer E, Issakidis-Bourguet E, Lavergne D, Miginiac-Maslow M. Chlamydomonas reinhardtii: a model organism for the study of the thioredoxin family. Plant Physiol. Biochem. (2003) 41:513–521.[CrossRef][Web of Science]
Lemaire SD, et al. NADP-malate dehydrogenase from unicellular green alga Chlamydomonas reinhardtii: a first step toward redox regulation? Plant Physiol (2005) 137:514–521.
Lemaire SD, Michelet L, Zaffagnini M, Massot V, Issakidis-Bourguet E. Thioredoxins in chloroplasts. Curr. Genet. (2007) 51:343–365.[CrossRef][Web of Science][Medline]
Li AD, Anderson LE. Expression and characterization of pea chloroplastic glyceraldehyde-3-phosphate dehydrogenase composed of only the B-subunit. Plant Physiol (1997) 115:1201–1209.[Abstract]
Marri L, Trost P, Pupillo P, Sparla F. Reconstitution and properties of the recombinant glyceraldehyde-3-phosphate dehydrogenase/CP12/phosphoribulokinase supramolecular complex of Arabidopsis. Plant Physiol (2005) 139:1433–1443.
Marri L, Trost P, Trivelli X, Gonnelli L, Pupillo P, Sparla F. Spontaneous assembly of photosynthetic supramolecular complexes as mediated by the intrinsically unstructured protein CP12. J. Biol. Chem. (2008) 283:1831–1838.
Meyer Y, Reichheld JP, Vignols F. Thioredoxins in Arabidopsis and other plants. Photosynth. Res. (2005) 86:419–433.[CrossRef][Web of Science][Medline]
Meyer Y, Riondet C, Constans L, Abdelgawwad MR, Reichheld JP, Vignols F. Evolution of redoxin genes in the green lineage. Photosynth. Res. (2006) 89:179–212.[CrossRef][Web of Science][Medline]
Michelet L, et al. Glutathionylation of chloroplast thioredoxin f is a redox signaling mechanism in plants. Proc. Natl Acad. Sci. U S A (2005) 102:16478–16483.
Michelet L, Zaffagnini M, Massot V, Keryer E, Vanacker H, Miginiac-Maslow M, Issakidis-Bourguet E, Lemaire SD. Thioredoxins, glutaredoxins, and glutathionylation: new crosstalks to explore. Photosynth. Res. (2006) 89:225–245.[CrossRef][Web of Science][Medline]
Navrot N, Collin V, Gualberto J, Gelhaye E, Hirasawa M, Rey P, Knaff DB, Issakidis E, Jacquot JP, Rouhier N. Plant glutathione peroxidases are functional peroxiredoxins distributed in several subcellular compartments and regulated during biotic and abiotic stresses. Plant Physiol (2006) 142:1364–1379.
Oesterhelt C, Klocke S, Holtgrefe S, Linke V, Weber AP, Scheibe R. Redox regulation of chloroplast enzymes in Galdieria sulphuraria in view of eukaryotic evolution. Plant Cell Physiol (2007) 48:1359–1373.
Pohlmeyer K, Paap BK, Soll J, Wedel N. CP12: a small nuclear-encoded chloroplast protein provides novel insights into higher-plant GAPDH evolution. Plant Mol. Biol. (1996) 32:969–978.[CrossRef][Web of Science][Medline]
Porter MA, Stringer CD, Hartman FC. Characterization of the regulatory thioredoxin site of phosphoribulokinase. J. Biol. Chem. (1988) 263:123–129.
Portis AR Jr., Li C, Wang D, Salvucci ME. Regulation of Rubisco activase and its interaction with Rubisco. J. Exp. Bot. (2008) 59:1597–1604.
Scagliarini S, Trost P, Pupillo P. The non-regulatory isoform of NAD(P)-glyceraldehyde-3-phosphate dehydrogenase from spinach chloroplasts. J. Exp. Bot. (1998) 49:1307–1315.
Scagliarini S, Trost P, Pupillo P, Valenti V. Light activation and molecular mass changes of NAD(P)-glyceraldehyde-3-phosphate dehydrogenase of spinach and maize leaves. Planta (1993) 190:313–319.[Web of Science]
Scheibe R, Wedel N, Vetter S, Emmerlich V, Sauermann SM. Co-existence of two regulatory NADP-glyceraldehyde 3-P dehydrogenase complexes in higher plant chloroplasts. Eur. J. Biochem. (2002) 269:5617–5624.[Web of Science][Medline]
Schmid M, Davison TS, Henz SR, Pape UJ, Demar M, Vingron M, Schölkopf B, Weigel D, Lohmann JU. A gene expression map of Arabidopsis thaliana development. Nat Genet. (2005) 37:501–506.[CrossRef][Web of Science][Medline]
Schürmann P, Buchanan BB. The ferredoxin/thioredoxin system of oxygenic photosynthesis. Antioxidants and Redox Signalling (2008) 10:1235–1274.[CrossRef][Web of Science][Medline]
Sparla F, Fermani S, Falini G, Zaffagnini M, Ripamonti A, Sabatino P, Pupillo P, Trost P. Coenzyme site directed mutants of photosynthetic A4-GAPDH show selectively reduced NADPH-dependent catalysis, similar to regulatory AB-GAPDH inhibited by oxidized thioredoxin. J. Mol. Biol. (2004) 340:1025–1037.[CrossRef][Web of Science][Medline]
Sparla F, Pupillo P, Trost P. The C-terminal extension of glyceraldehyde-3-phosphate dehydrogenase subunit B acts as an autoinhibitory domain regulated by thioredoxins and nicotinamide adenine dinucleotide. J. Biol. Chem. (2002) 277:44946–44952.
Sparla F, Zaffagnini M, Wedel N, Scheibe R, Pupillo P, Trost P. Regulation of photosynthetic GAPDH dissected by mutants. Plant Physiol (2005) 138:2210–2219.
Tamoi M, Myazaki T, Fukamizo T, Shigeoka S. The Calvin cycle in cyanobacteria is regulated by CP12 via NAD(H)/NADP(H) ratio under light/dark conditions. Plant J (2005) 42:504–513.[CrossRef][Web of Science][Medline]
Traverso JA, Vignols F, Cazalis R, Serrato AJ, Pulido P, Sahrawy M, Meyer Y, Cejudo FJ, Chueca A. Immunocytochemical localization of Pisum sativum TRXs f and m in non-photosynthetic tissues. J. Exp. Bot. (2008) 59:1267–1277.
Trost P, Fermani S, Marri L, Zaffagnini M, Falini G, Scagliarini S, Pupillo P, Sparla F. Thioredoxin-dependent regulation of photosynthetic glyceraldehyde-3-phosphate dehydrogenase: autonomous vs. CP12-dependent mechanisms. Photosynth. Res. (2006) 89:263–275.[CrossRef][Web of Science][Medline]
Trost P, Scagliarini S, Valenti V, Pupillo P. Activation of spinach chloroplast glyceraldehyde-3-phosphate dehydrogenase: effect of glycerate 1,3-bisphosphate. Planta (1993) 190:320–326.[Web of Science]
Vieira Dos Santos C, Laugier E, Tarrago L, Massot V, Issakidis-Bourguet E, Rouhier N, Rey P. Specificity of thioredoxins and glutaredoxins as electron donors to two distinct classes of Arabidopsis plastidial methionine sulfoxide reductases B. FEBS Lett. (2007) 581:4371–4376.[CrossRef][Web of Science][Medline]
Wedel N, Soll J. Evolutionary conserved light regulation of Calvin cycle activity by NAPDH-mediated reversible phosphoribulokinase/CP12/glyceraldehyde-3-phosphate-dehydrogenase complex dissociation. Proc. Natl Acad. Sci. U S A (1998) 95:9699–9704.
Wedel N, Soll J, Paap BK. CP12 provides a new mode of light regulation of Calvin cycle activity in higher plants. Proc. Natl Acad. Sci. U S A (1997) 94:10479–10484.
Zaffagnini M, Michelet L, Marchand C, Sparla F, Decottignies P, Le Maréchal P, Miginiac-Maslow M, Noctor G, Trost P, Lemaire SD. The thioredoxin independent isoform of photosynthetic glyceraldehyde-3-phosphate dehydrogenase is selectively regulated by glutathionylation. FEBS J (2007) 274:212–226.[CrossRef][Medline]
Zaffagnini M, Michelet L, Massot V, Trost P, Lemaire SD. Biochemical characterization of glutaredoxins from Chlamydomonas reinhardtii reveals the unique properties of a chloroplastic CGFS-type glutaredoxin. J. Biol. Chem. (2008) 283:8868–8876.
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